Leave a comment

M E T H O D S I N M O L E C U L A R M E D I C I N E

Humana Press
M E T H O D S  I N  M O L E C U L A R  M E D I C I N E
Molecular
Pathology
Protocols
Edited by
Anthony A. Killeen
Humana Press
Molecular
Pathology
Protocols
Edited by
Anthony A. Killeen
DNA Extraction from Paraffin-Embedded Tissues 1
1
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
1
DNA Extraction from Paraffin-Embedded Tissues
Hongxin Fan and Margaret L. Gulley
1. Introduction
In routine histopathology, most tissues are fixed in formalin and embedded
in paraffin for long-term preservation. DNA can be extracted from these
tissues for subsequent molecular analysis by amplification methods. We
describe herein a protocol for DNA preparation from paraffin-embedded
tissues based on published procedures (1–3) . In brief, tissue sections are placed
into microfuge tubes, then deparaffinized with xylene. The xylene is removed
with ethanol washes, and the tissue is treated with proteinase K to make DNA
available for amplification.
This protocol is simple, but there are several factors that influence the suc-cess of subsequent DNA amplification assays, including the type of fixative
that is used, the duration of fixation, the age of the paraffin block, and the
length of the DNA segment to be amplified (see Note 1). Ethanol fixation pre-serves DNA much better than does formalin. Formalin fixation randomly chops
DNA in a duration-dependent manner, resulting in partial degradation. Even
more severe DNA degradation occurs in bone samples subjected to acid decal-cification. Because of this degradation, formalin-fixed tissue is not suitable for
Southern blot analysis or for amplification of large DNA segments. Neverthe-less, polymerase chain reaction (PCR) amplification of segments ranging up to
1300 bp has been reported (2) , and consistent amplification of segments up to
300 bp is commonly achieved from archival fixed tissues. Be aware that partial
degradation of DNA may result in sampling bias, and therefore results should
be interpreted with caution. For example, amplification of DNA from one cell
may produce a PCR product that is not representative of the entire population
of cells in the tissue. For this reason, it is wise to run tests in duplicate and
always with appropriate controls.
2 Fan and Gulley
2. Materials
2.1. Reagents
1. Xylene.
2. 100% Ethanol.
3. Proteinase K stock solution (20 mg/mL).
4. TEN buffer: 10 mM Tris-HCl, pH 8.0, 1 mM EDTA, pH 8.0, 20 mM NaCl.
2.2. Equipment
1. Microcentrifuge.
2. Heating block or water bath to hold 1.5-mL Eppendorf tubes.
3. Microtome.
3. Methods
Preparation of paraffin-embedded tissue for DNA amplification involves
several manual manipulations; therefore precautions should be taken to avoid
contamination, such as changing gloves frequently. When opening and closing
microfuge tubes, do not touch the rim or inside of the cap. (Many laboratory
scientists place a fresh gauze square over their thumb when opening a tube, or
use a cap-opener device or screw-top lids.) Appropriate negative controls must
be used to alert for contamination.
3.1. Cutting and Deparaffinizing Sections
1. Use a microtome to cut five sections, 5–20 µm thick, from a paraffin block, and
place these directly into a 1.5-mL microfuge tube. The thickness of the sections
depends on the size of the biopsy. For a small biopsy (up to 3 mm), 20-µm thick
sections may be required, whereas a large biopsy requires only 5-µm thick sec-tions. Although multiple thin sections can be placed in a single tube, fewer thick
sections are more practical for processing.  See Note 2 for special precautions
against contamination.
2. Add 800  µL of xylene to each tube, close, mix by gentle vortexing, and then
incubate at room temperature for 10 min. Pellet the tissue by centrifugation for
3 min in a microfuge at full speed. Carefully remove and discard the supernatant
using a pipet; do not disturb the tissue. If any translucent white paraffin remains,
repeat the xylene wash one to two more times.
3. Add 800 µL of 100% ethanol to each tube, close the lid, and mix by inverting.
Pellet the tissue by centrifugation for 3 min in a microfuge at full speed, and
carefully remove and discard the supernatant with a pipet. Repeat the ethanol
wash one more time, and remove as much supernatant as possible.
4. Open the tubes and let the residual ethanol evaporate by incubating in a dry heat
block at 55°C for 15–30 min or until the sample is completely dry. (Speed-vacuum drying is not recommended because of the risk of contamination.)
DNA Extraction from Paraffin-Embedded Tissues 3
3.2. Proteinase K Digestion
1. To the dried tissue samples add 100 µL of TEN buffer containing 200 µg/mL of
proteinase K (prepared by mixing 1 µL of proteinase K stock solution in 100 µL
of TEN buffer). Large tissue samples should be resuspended in 200 µL or more
of this solution.
2. Close the tubes and incubate at 55°C for 3 h. (Large tissues should be incubated
overnight.)
3. Spin briefly to remove any liquid from the cap. Cover the caps tightly with cap
locks (PGC Scientifics, Gaithersburg, MD) to prevent them from popping open
during high-temperature incubation. Incubate in a 95°C heat block for exactly 10
min to inactivate the proteinase K (see Note 3). Pellet the tissue in a microfuge at
full speed for 10 min, and then transfer the supernatant to a clean tube and dis-card the pellet. Promptly proceed with PCR amplification.
4. Quantitation of DNA is not recommended; rather, the amount of supernatant
required for subsequent DNA amplification is determined empirically. Try
1- and 10-µL vol of the supernatant as a template for a 100-µL PCR amplifica-tion. If PCR products are not generated, then different volumes can be tried (see
Note 4). A positive control reaction (e.g., `-globin) should be run to ensure that
amplifiable DNA of similar length to the target DNA is present in the sample.
(See Note 5 for modification of thermocycling parameters.)
5. Store DNA at –20°C, and avoid thawing and refreezing. (Freshly prepared
samples are more efficiently amplified than those stored frozen, perhaps because
the freeze-thaw cycle damages DNA.)
4. Notes
1. The most important factor affecting DNA quality is the type of fixative employed
and the duration of fixation. Tissues fixed between 12 and 24 h in ethanol,
acetone, Omnifix, or 10% buffered formalin usually yield good-quality DNA;
but B-5, Zenker’s, or Bouin’s solutions, or duration of fixation longer than 5 d,
are poor prognostic factors for PCR productivity  (2 ,4) . Prior studies showed
that when the length of `-globin amplification product increased (175, 324, and
676 bp), the percentage of fixed tissues containing amplifiable DNA decreased
(100, 69, and 45%) (5) . And when the age of a block increased, PCR productivity
decreased (6 ,7) , although some blocks stored for more than 40 yr were success-fully studied (8) .
2. It is important that no tissue be carried over from one case to the next during
microtomy. Between each block, shift to a fresh part of the blade. Use smooth-edge rather than toothed forceps for transporting sections into tubes. Do not
allow bleach to come in contact with the tissue or the DNA will be destroyed. Ice
cubes used to cool a block should be discarded between cases.
3. Longer incubation at 95°C may damage DNA, whereas shorter incubation may
not fully inactivate proteinase K. Additional time is needed for volumes >500 µL.
4 Fan and Gulley
4. The optimal amount of template for an amplification reaction depends on numer-ous factors specific to each sample, such as DNA concentration and presence of
inhibitors. It is useful to test several concentrations of each template (e.g., 1 and
10 µL of template per 100-µL PCR). Large tissues may necessitate the use of a
smaller fraction of the template (e.g., 0.1  µL). The amount of template that is
“tolerated” in a PCR may be affected by residual fixation chemicals or paraffin,
excessive tissue debris, and other factors. If the first attempt fails, a 10-fold dilu-tion will often reduce inhibitors while still retaining enough DNA to allow
amplification.
5. Amplification of DNA prepared from paraffin-embedded tissue is less efficient
than amplification of DNA from fresh or frozen tissues. To compensate for this
reduced efficiency, consider modifying the thermocycling parameters by increas-ing the number of cycles and lengthening the duration at each temperature within
the cycle (1) .
References
1. Wright, D. K. and Manos, M. M. (1990) Sample preparation from paraffin-embedded tissues, in  PCR Protocols: A Guide to Methods and Applications
(Innis, M. A., Glefand, D. H., and Sninsky, J. J., eds.), Academic, San Diego,
pp. 153–158.
2. Greer, C. E., Wheeler, C. M., and Manos, M. M. (1994) Sample preparation
and PCR amplification from paraffin-embedded tissues. PCR Methods Appl. 3,
S113–S122.
3. Rolfs, A., Schuller, I., Finckh, U., and Weber-Rolfs, I. (1992)  PCR: Clinical
Diagnostics and Research, Springer-Verlag, Berlin, pp. 85–87.
4. Shibata, D. (1994) Extraction of DNA from paraffin-embedded tissue for analysis
by polymerase chain reaction: new tricks from an old friend. Hum. Pathol. 25,
561–563.
5. Liu, J., Johnson, R. M., and Traweek, S. T. (1993) Rearrangement of the BCL-2
gene in follicular lymphoma: detection by PCR in both fresh and fixed tissue
samples. Diagn. Mol. Pathol. 2, 241–247.
6. Limpens, J., Beelen, M., Stad, R., Haverkort, M., van Krieken, J. H., van Ommen,
G. J., and Kluin, P. M. (1993) Detection of the t(14;18) translocation in frozen and
formalin-fixed tissue. Diagn. Mol. Pathol. 2, 99–107.
7. Goelz, S. E., Hamilton, S. R., and Vogelstein, B. (1985) Purification of DNA
from formaldehyde fixed and paraffin embedded human tissue. Biochem. Biophys.
Res. Commun. 130, 118–126.
8. Shibata, D., Martin, W. J., and Arnheim, N. (1988) Analysis of DNA sequences in
forty-year-old paraffin-embedded thin-tissue sections: a bridge between molecu-lar biology and classical histology. Cancer Res. 48, 4564–4566.
DNA Extraction from Fresh or Frozen Tissues 5
2
DNA Extraction from Fresh or Frozen Tissues
Hongxin Fan and Margaret L. Gulley
1. Introduction
The first step in molecular analysis of patient tissues is preparation of puri-fied, high molecular weight DNA. A number of methods and commercial kits
are available for DNA isolation. Traditional organic extraction protocols (1,2)
are based on the fact that DNA is soluble in water whereas lipids are soluble in
phenol. In these protocols, tissues are disaggregated and then treated with
detergent to lyse cell membranes followed by proteinase to digest proteins.
Phenol, an organic solvent, is added to help separate the lipids and protein
remnants from the DNA. Chloroform is then used to facilitate the removal of
phenol. DNA is subsequently concentrated and further purified by precipita-tion in a cold mixture of salt and ethanol. Finally, DNA is resolubilized in
Tris-EDTA buffer.
The traditional organic extraction procedure presented herein is used by
many laboratories to obtain abundant high molecular weight DNA. However,
in recent years, there has been a trend toward adoption of commercial non-organic protocols that are faster and avoid the toxicity inherent with phenol
exposure. A popular nonorganic extraction kit that works particularly well on
blood and marrow samples is the Puregene DNA Extraction Kit (Gentra Sys-tems, Minneapolis, MN). This kit can also be adapted for use on solid tissue
samples for subsequent polymerase chain reaction (PCR) analysis.
2. Materials
2.1. Reagents
1. Lymphocyte Separation Media (ICN Biomedicals Inc., Aurora, OH).
2. 1X phosphate buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 4.3 mM
Na2HPO4, 1.4 mM KH2PO4.
5
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
6 Fan and Gulley
3. Liquid nitrogen.
4. DNA extraction buffer: 10 mM NaCl, 20 mM Tris-HCl, pH 8.0, 1 mM EDTA.
5. 10% sodium dodecyl sulfate (SDS).
6. Proteinase K solution: 10 mg/mL of proteinase K in 50 mM Tris-HCl, pH 7.5;
store at 4°C.
7. Phenol equilibrated with 0.1 M Tris-HCl, pH 8.0.
8. Chloroform: isoamyl alcohol (24 1).
9. 3 M Sodium acetate, pH 5.2.
10. 100% Ethanol.
11. 70% Ethanol.
12. TE buffer: 10 mM Tris-HCl, pH 8.0, 0.1 mM EDTA.
13. Agarose.
14. 1X TAE buffer: 40 mM Tris-acetate, 1 mM EDTA.
15. Ethidium bromide (10 mg/mL).
16. 10X Gel loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol, 15%
Ficoll (type 400) in 10X TAE buffer.
17. DNA molecular weight marker.
2.2. Equipment
1. Mortar and pestle.
2. Water baths at 37, 50, and 55°C.
3. Centrifuge.
4. Spectrophotometer.
5. Horizontal gel electrophoresis apparatus.
6. DC power supply.
3. Methods
3.1. Sample Preparation
3.1.1. Tissue Specimen
1. Mince fresh, solid tissue up to 3 mm3 into small pieces (1–2 mm) with a sterile
scalpel blade. Process the tissue within 2 h of collection, or freeze at –20°C or
colder until the time of DNA extraction (see Note 1).
2. Place the tissue in a clean mortar filled with liquid nitrogen. (See Note 2 for
cleaning instructions.)
3. Using a clean pestle, grind the frozen tissue to a powder while it is submerged in
liquid nitrogen. While grinding, cover the mortar with a paper towel to keep
tissue fragments inside the mortar, and work under a hood to protect yourself
from aerosolized powder.
4. Allow the liquid nitrogen to evaporate, leaving a dry frozen tissue powder in the
mortar.
DNA Extraction from Fresh or Frozen Tissues 7
3.1.2. Ficoll Separation of Mononuclear Cells
from Blood and Marrow Aspirates
Prior to DNA extraction, mononuclear cells are isolated from anticoagu-lated blood or bone marrow aspirates by Ficoll centrifugation. (See Note 3 for
information about sample stability.) About 107 nucleated cells yield 40 µg of
DNA for Southern blot analysis, and 3  × 106 cells yield sufficient DNA for
amplification testing.
1. To a 15-mL conical tube, add 4.5 mL of blood and an equal volume of PBS. For
bone marrow aspirates, use 1 mL of marrow and 8 mL of PBS. If less sample
volume is available, use all of it. If greater sample volume is desired, split the
sample evenly among two or more tubes so that all of it is processed, and then
recombine the samples on collection of the mononuclear cell layer.
2. Using a Pasteur pipet, underlay the diluted blood or marrow with 3 mL of Ficoll
solution (Lymphocyte Separation Media).
3. Cap the tube and centrifuge at 400g for 30 min at room temperature in a swinging
bucket rotor.
4. Use a plastic Pasteur pipet to aspirate the mononuclear cell layer, which is the
fuzzy white layer located between the plasma and the separation medium, into a
clean 15-mL conical tube. Avoid the red cell layer at the bottom of the tube. (If
no mononuclear cell layer is visible, see Note 4.)
5. Resuspend the mononuclear cells in PBS to 12 mL total volume.
6. Centrifuge for 10 min at 1700g at room temperature. Remove and discard the
supernatant by pouring it off.
7. Store the cell pellet at –20°C temporarily or at –70°C long term, or proceed
directly to DNA or RNA extraction.
3.2. DNA Extraction
3.2.1. Cell Lysis and Digestion
The procedure for solid tissue differs from that of blood or marrow mono-nuclear cells only in the first step.
1. For solid tissue, add 920 µL of DNA extraction buffer to the tissue powder in the
mortar, and gently mix with the pestle. If the buffer freezes, wait until it thaws before
proceeding. Then transfer the fluid to a 15-mL conical tube or microfuge tube by
gentle pipeting. For a mononuclear cell pellet (about 107 cells), resuspend the cells in
920 µL of DNA extraction buffer and mix well by gentle pipeting.
2. Add 50 µL of 10% SDS to the mixture and mix well; the solution should become
viscous.
3. Add 30 µL of proteinase K solution to the viscous mixture. Close the cap tightly
and mix vigorously by repeated forceful inversion or vortex.
4. Incubate in a 37°C water bath for at least 6 h or as long as 2 d, or at 55°C for 3 h;
gently invert the tube a few times during incubation.
8 Fan and Gulley
5. The lysed sample should be viscous and relatively clear. This sample may be
stored at 4°C for up to 1 wk before subjecting it to phenol/chloroform extraction
as described in Subheading 3.2.2., or before proceeding with nonorganic extrac-tion as described in Note 5.
3.2.2. Phenol/Chloroform Extraction of DNA
1. Add an equal volume of equilibrated phenol, close the cap tightly, and mix gently
by inversion for 1 min.
2. Spin the tube at 1700g in a swinging bucket rotor at room temperature for 10 min.
3. With a plastic pipet, aspirate the upper clear aqueous layer and transfer it to
another clean labeled tube. This should be done carefully to avoid carrying over
phenol or white proteinaceous material from the interface.
4. Repeat the phenol extraction (steps 1–3) one more time.
5. Next, extract with an equal volume of chloroform instead of phenol, and save the
supernatant to another clean tube after centrifugation.
6. Repeat the chloroform extraction; this helps eliminate all of the phenol from the
DNA sample.
3.2.3. Purification and Precipitation of DNA
1. To the aqueous DNA solution add 0.1 vol of 3 M sodium acetate (pH 5.2), and
mix well by vortexing.
2. Add 2 vol of ice-cold 100% ethanol to the tube. Close the cap tightly and mix by
inversion. A white cotton-like precipitate should form.
3. Use a sterile plastic rod to spool the precipitated DNA. (If no precipitate is vis-ible, then microfuge at full speed for 10 min, rinse the pellet with 70% ethanol,
air-dry for about 10–15 min, and proceed with step 6.)
4. Rinse the spooled DNA thoroughly in 1 mL of cold 70% ethanol by dipping.
5. Remove the DNA-coated plastic rod and allow the precipitate to air-dry until the
white precipitate becomes clear, usually about 5–10 min.
6. Dissolve the precipitate in an appropriate volume of TE buffer (typically about
100–500 µL; targeting an optimal DNA concentration 1 µg/µL), scraping the rod
along the wall of the microfuge tube to help detach the viscous DNA.
7. Allow the DNA to dissolve in the TE buffer for at least 4 h at 50°C, gently shak-ing periodically during incubation. Failure to adequately resolubilize the DNA
will result in uneven distribution of DNA within the solution.
8. The purified DNA sample may be stored for 4 wk at 4°C prior to analysis, or
indefinitely at –20°C.
3.3. DNA Quantitation by Spectrophotometry
1. Mix the DNA sample by gentle vortexing and inversion.
2. Add 5 µL of the DNA sample to 495 µL of sterile water and mix well.
3. Place the diluted sample in a quartz microcuvet and measure the absorbance at
260 and 280 nm against a water blank. (Nucleic acids absorb light maximally at
260 nm whereas proteins absorb strongly at 280 nm.)
DNA Extraction from Fresh or Frozen Tissues 9
4. Compute the DNA concentration based on the concept that an OD260 of 1 corre-sponds to 50  µg/mL of double-stranded DNA, and adjusting for the 100-fold
dilution factor, according to the following formula:
DNA concentration (µg/µL) = OD260 × 5
5. The OD260 OD280 ratio should be between 1.7 and 2.0. Lower values indicate
protein contamination, in which case the DNA can be further purified by addi-tional phenol/chloroform extractions followed by ethanol precipitation.
3.4. Gel Electrophoresis to Analyze DNA Quality
Agarose gel electrophoresis can be used to assess the intactness of purified
DNA. High molecular weight DNA is needed for Southern blot analysis,
whereas partially degraded DNA might be suitable for amplification
procedures.
1. Prepare a 0.7% agarose gel in 1X TAE buffer containing 0.5 µg/mL of ethidium
bromide.
2. Mix an aliquot of the extracted DNA sample with loading buffer, and load into a
submerged well. Control samples representing intact and degraded DNA should
be loaded into adjacent wells.
3. Electrophorese in 1X TAE buffer with 0.5 µg/mL ethidium bromide at 2 V/cm,
until the dye front reaches the end of the gel.
4. View the gel under UV light. High molecular weight DNA is too large to migrate
well under these conditions, whereas degraded DNA contains a spectrum of
smaller fragment sizes that appear as a smear across the lane.
4. Notes
1. Solid tissue samples should be processed immediately or else frozen to minimize
the activity of endogenous nucleases. If frozen tissue immunohistochemistry is
planned, then slice the tissue into pieces no more than 5 mm thick and snap-freeze in liquid nitrogen or in a cryostat. If morphologic preservation is not
needed, then place the tissue in a –70°C freezer indefinitely, or at –20°C for up to
3 d until DNA or RNA isolation.
2. After washing with detergent and rinsing well, soak the mortar and pestle in 50%
nitric acid or 10% bleach to prevent carryover of DNA to the next case, then rinse
well.
3. Peripheral blood or bone marrow aspirate anticoagulated with EDTA or acid
citrate dextrose should be stored at room temperature and processed as soon
as possible. EDTA beneficially chelates ions to inhibit nucleases from
degrading nucleic acid, and consequently DNA and RNA are often stable for
up to 48 h at room temperature. Heparin anticoagulant is not recommended
because residual heparin may interfere with subsequent restriction enzyme or
DNA polymerase activity.
10 Fan and Gulley
4. If no mononuclear cell layer is visible, follow these steps to recover mononuclear
cells from the red cell pellet. (This procedure may be particularly helpful if the
blood has been previously refrigerated, thus causing the white cells to clump and
sediment with the red cells at the bottom of the tube. For this reason, refrigeration
of samples is not recommended prior to Ficoll centrifugation.) Carefully remove
and discard the supernatant with a Pasteur pipet. Avoid disturbing the red cell
pellet. Add 3 vol of Puregene RBC Lysis Solution (Gentra Systems) to the pellet.
Invert to mix several times, and incubate for 10 min at room temperature. Invert
again once during the incubation. Centrifuge at 2000g for 10 min, remove the
supernatant with a Pasteur pipet, and discard it. The mononuclear cell pellet is
not always visible at this step, and if not, you may use half-volumes of the extrac-tion reagents in the subsequent steps of extraction.
5. To avoid the use of organic solvents, numerous nonorganic extraction protocols
have been developed. We recommend the Puregene DNA Extraction Kit (Gentra
Systems) for blood and marrow aspirates. To adapt this kit for PCR analysis of
solid tissues, follow the protocol in this chapter through SDS and proteinase
K digestion. Then apply the Puregene kit according to the manufacturer’s
instructions, skipping the red blood cell lysis and cell lysis steps.
References
1. Strauss, W. M. (1995) Preparation of genomic DNA from mammalian tissue, in
Current Protocols in Molecular Biology (Ausubel, F. M., Brent, R., Kingston,
R. E., et al., eds.), John Wiley & Sons, New York, pp. 2.2.1–2.2.3.
2. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989)  Molecular Cloning: A
Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring
Harbor, NY, pp. 9.14–9.23.
RNA Extraction from Fresh or Frozen Tissues 11
11
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
3
RNA Extraction from Fresh or Frozen Tissues
Hongxin Fan and Margaret L. Gulley
1. Introduction
RNA can be isolated from fresh or frozen tissue, then purified and quanti-fied for subsequent molecular analysis. RNA is quite labile compared to DNA
for good reason: RNA is the transient message that transmits information from
activated genes. Once gene transcription is turned off, no more RNA is pro-duced, and preexisting RNA is rapidly degraded to prevent continued transla-tion of proteins. Because RNA is so labile, special care is required at all steps
of RNA extraction to prevent RNA degradation.
Total RNA can be extracted from cells by treating them with guanidine
isothiocyanate, a chemical that disrupts cellular membranes and also inhibits
endogenous RNase activity (1 ,2). This reagent forms the basis for several com-mercial kits that are available for total RNA isolation. In the protocol described
herein, we use the TRIzol™ reagent (Gibco-BRL, Gaithersburg, MD), which
combines phenol and guanidine isothiocyanate in a single solution to produce
high yields of RNA in a rapid isolation procedure. This method is applicable to
fresh or frozen tissue including blood or bone marrow samples.
2. Materials
2.1. Reagents
1. TRIzol reagent (Gibco-BRL); store at 4°C.
2. Chloroform.
3. Isopropanol.
4. Diethylpyrocarbonate (DEPC)-treated H2O (see Note 1).
5. 75% ethanol made with DEPC-H2O.
6. Ribonuclease inhibitor (40 U/µL) (Promega, Madison, WI).
12 Fan and Gulley
7. 10X DNase I buffer (Gibco-BRL)
8. DNase I, amplification grade (Gibco-BRL).
9. 20 mM EDTA.
10. Phenol chloroform isoamyl alcohol (25 24 1).
11. Chloroform isoamyl alcohol (24 1).
12. 2 M Sodium acetate made with DEPC-H2O, pH 4.5.
13. 100% Ethanol.
2.2. Equipment
1. Mortar and pestle.
2. Centrifuge.
3. Spectrophotometer.
3. Methods
3.1. Sample Preparation
Fresh and frozen tissues including blood and bone marrow aspirates are ini-tially prepared in the same way as for DNA extraction. For example, solid
tissues are frozen and ground into a powder under the liquid nitrogen, whereas
blood and marrow mononuclear cells are isolated by Ficoll centrifugation as
described elsewhere in this volume. Once the tissue powder or mononuclear
cell pellet is obtained, instead of adding DNA lysis buffer, proceed with the
following RNA extraction procedure.
3.2. RNA Extraction
Total RNA is isolated using TRIzol reagent according to the manufacturer’s
directions (Gibco-BRL). The procedure for solid tissues differs from that of
blood or marrow mononuclear cells only in the first step. All of the following
procedures must be carried out using RNase-free solutions and labware (see
Notes 1–3).
1. Cell lysis: For solid tissue samples that have been ground to a powder, add 1 mL
of TRIzol reagent per 50–100 mg of tissue (up to 3 mm3). Gently mix with a
pestle and then transfer to a microfuge tube by pipetting. Pipet up and down three
to five times until tissue clumps are dispersed. If desired, this sample can now be
stored at –70°C for at least 1 mo. Prior to use, the mortar and pestle should be
baked at 150°C for 4 h to remove RNases (see Note 3). For mononuclear cells
obtained from blood or bone marrow, add 1 mL of TRIzol reagent per 106–107
cells and then lyse the cells by pipetting up and down three to five times until
clumps are dispersed. This sample can now be stored at –70°C for at least 1 mo.
2. Phase separation: Incubate the sample for 5 min at room temperature. Add
0.2 mL of chloroform, shake the tube vigorously by hand for 15 s, and incubate
at room temperature for 3 min. Centrifuge the sample at no more than 12,000g at
4°C for 15 min.
RNA Extraction from Fresh or Frozen Tissues 13
3. RNA precipitation: Transfer the upper colorless aqueous phase to a fresh labeled
microfuge tube; the lower chloroform phase can be discarded. Precipitate the
RNA by adding 0.5 mL of isopropanol. Incubate at room temperature for 10 min.
Recover the RNA pellet by centrifugation at no more than 12,000g at 4°C for
10 min.
4. RNA wash: Remove and discard the supernatant by pipetting. Wash the RNA
pellet with 1 mL of cold 75% ethanol, mix by vortexing, and centrifuge at no
more than 7500g at 4°C for 5 min. Then remove as much ethanol supernatant as
possible.
5. Redissolving of RNA: Air-dry the RNA pellet by opening the cap for about
30–60 min, but do not dry it completely because this will greatly decrease its
solubility. Dissolve RNA in about 50  µL of DEPC-H2O (targeting an optimal
RNA concentration of 1 µg/µL) by pipetting up and down a few times and then
incubating at 37°C for 10 min. This RNA sample may be stored temporarily
at –20°C, or add 1 µL of RNasin and store long term at –70°C.
3.3. DNase Treatment
Contaminating DNA in the RNA sample could potentially interfere with
subsequent RNA analysis. For example, genomic DNA might compete with
cDNA in reverse transcriptase polymerase chain reaction if the two primers do
not span an intronic splice. In this case, treatment of the isolated RNA with
DNase is recommended as follows:
1. Add 5 µL of 10X DNase I reaction buffer and 2 µL of 1 U/µL of amplification
grade DNase I (Gibco-BRL) into 43 µL of the RNA sample. Incubate at room
temperature for 30 min.
2. Stop the DNase reaction by adding 5 µL of 20 mM EDTA.
3. Add 45 µL of DEPC-H2O per tube to bring the sample volume to 100 µL, and
mix with 100  µL of 25 24 1 Phenol chloroform isoamyl alcohol. Spin in a
microfuge at 12,000g for 5 min at 4°C, and then transfer the upper aqueous layer
to a fresh labeled tube.
4. Add 100  µL of 24 1 chloroform isoamyl alcohol, mix well, spin for 5 min at
4°C, and transfer the upper aqueous layer to a fresh labeled tube.
5. Add 15 µL of 2 M sodium acetate (pH 4.5) and mix well. Add 2.5 vol of cold
100% ethanol and precipitate at –70°C for at least 30 min.
6. Recover the RNA by centrifuging in a microfuge at 12,000g for 15 min at 4°C.
Wash the pellet with 75% ethanol one time.
7. Air-dry the pellet and dissolve RNA in about 10 µL of DEPC-H2O (optimal RNA
concentration is about 1 µg/µL), add 1 µL of RNasin, and store at –70°C.
3.4. Spectrophotometric RNA Quantitation
1. Add 2 µL of the RNA sample to 398 µL of DEPC-H2O.
2. Place the diluted sample in a quartz microcuvet and measure the absorbance at
260 and 280 nm against a DEPC-H2O blank.
14 Fan and Gulley
3. Compute the RNA concentration using the following formula based on the con-cept that an OD260 of 1 corresponds to 40 µg/mL of RNA, and adjusting for the
200-fold dilution factor:
RNA concentration (µg/µL) = OD260 × 8.
4. Notes
1. All solutions used in RNA preparation should be treated with DEPC. DEPC inac-tivates RNases by covalent modification. Add 0.1% DEPC to each solution, mix
well, and incubate at 37°C overnight (or stir vigorously for at least 2 h), and then
autoclave for 20 min. Unfortunately, DEPC cannot be used to treat solutions con-taining Tris because Tris inactivates DEPC. Therefore, it is recommended that an
RNase-free spatula be used to retrieve Tris crystals from a fresh stock, and these
crystals should be dissolved in H2O previously treated with DEPC.
2. The main factor influencing the success of RNA extraction is inhibition of
endogenous RNase activity and prevention of exogenous RNase contamination.
RNases are ubiquitous, stable enzymes that generally require no cofactors to func-tion. Human skin is a primary source of external RNase contamination, so gloves
should be worn during all steps of the extraction procedure. All solutions, glass-ware, and plastic containers must be RNase free. When feasible, disposable
plasticware should be used and then discarded.
3. Glassware should be baked at 150°C for 4 h to reduce RNase activity. Plastic
items can be soaked in 0.5 M NaOH for 10 min, rinsed thoroughly with water,
and autoclaved. Plasticware taken straight out of the original manufacturer’s
package is generally considered to be free from RNase contamination.
4. The TRIzol manufacturer’s protocol recommends a power homogenizer to dis-perse solid tissues, but we have found it easier to manually grind the tissue to a
powder under liquid nitrogen prior to addition of TRIzol reagent.
References
1. Chomczynski, P. and Sacchi, N. (1987) Single-step method of RNA isolation by
acid guanidinium thiocyanate-phenol-chloroform extraction.  Analyt. Biochem.
162, 156–159.
2. Kingston, R. E., Chomczynski, P., and Sacchi, N. (1995) Guanidine methods
for total RNA preparation, in Current Protocols in Molecular Biology (Ausubel,
F. M., Brent, R., Kingston, R. E., et al., eds.), John Wiley & Sons, New York,
pp. 4.2.1–4.2.9.
SSCP in Exons 4–8 of the TP53 Gene 15
15
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
4
Single-Strand Conformation Polymorphism
Analysis of Mutations in Exons 4–8 of the TP53 Gene
Kirsi H. Vähäkangas, Katariina Castrén, and Judith A. Welsh
1. Introduction
The TP53 tumor suppressor gene coding for a nuclear phosphoprotein
involved in cellular stress responses is the most frequently mutated gene in
human cancers described so far (1–4) . Mutations are found throughout the gene
but most frequently within the highly conserved middle region (exons 5–8)
that encodes for the DNA-binding central region of the gene critical for the
major function of TP53 protein as a transcriptional activator (5). The mutation
spectrum of the TP53 gene varies from one tumor type to another with typical
hot-spot codons for mutations (1–2 ,6) . For instance, codons 157, 248, and 273
are frequently mutated in cigarette smoking–associated lung cancer, whereas
mutations in codon 175 are rare. This codon, on the other hand, is often
mutated in breast and colon cancer. In some cases, typical mutations can be
linked with environmental exposures, such as CCATT double mutation with
UV radiation (7) and codon 249 AGGAAGT mutation with aflatoxin B1 and
hepatitis B virus (8) . These findings, in connection with the fact that one of the
main functions of TP53 protein is putatively the protection of the genome (9,10),
implicate the mutations of the TP53 gene in environmentally induced carcino-genesis in humans and the possible use of TP53-related markers in molecular
epidemiology (11–13) .
Mutations of the p53 gene with the loss of wild-type function also seem to
have clinical importance. The reported tumor types, in which either a  TP53
gene mutation or aberrant expression of p53 protein indicates a worse progno-sis, include breast cancer (14–16) and bladder cancer (17 ,18) . Furthermore, the
known polymorphisms of the p53 gene may be involved in the susceptibility to
cancers and prognosis of the disease (19 ,20) . Proallele carriers of exon 4 codon
16 Vähäkangas, Castrén, and Welsh
72 polymorphism (CGCArg or CCCPro) were found to be overrepresented among
patients who smoke and also among those affected at a younger age (21) , and
an excess of Arg/Arg homozygotes was found in nonsmoking patients with
lung cancers (22) . p53Arg allele displays several differences in comparison to
p53Pro. For instance, it is more susceptible to degradation by human papilloma
virus (HPV) E6 protein  (23 ,24) . There are indications that this translates to
higher susceptibility to HPV-associated cervical cancer (23 ,25) , although stud-ies contradicting these findings also exist (26–30) .
Among the most frequently used methods for the detection of mutation of the
p53 gene are sequencing and single-strand conformation polymorphism (SSCP)
analysis. Although originally SSCP was developed as a radioactive method (31),
currently nonradioactive applications are available and, for obvious reasons, gain-ing more popularity. Further reasons for the increasing popularity are that SSCP
has also been shown to be more sensitive than manual sequencing (32–34) and
because of the introduction of smaller, more manageable gels and temperature-controlled equipment (35–39).
2. Materials
As a negative control, lymphocyte DNA from a healthy volunteer deter-mined to be wild type by sequencing can be used. For positive controls, wild-type lymphocyte DNA is amplified using modified primers to integrate a
change in the sequence. A positive and a negative control are included in each
gel, because slight variation in the performance of the assay may occur from
one piece of equipment to another.
2.1. Purification of DNA
DNA purified for any other purpose serves well as a starting material for
this protocol. However, for those who want to use paraffin-embedded tissue
blocks, we provide a method that has worked well for us:
1. 100 mg of proteinase K in 20 mL of 10% sodium dodecyl sulfate (SDS).
2. Phenol buffered with Tris-HCl, pH 7.5–7.8 (cat. no. 15513-039; Life Technologies).
3. Chloroform isoamyl alcohol (24 1).
4. 0.5 M ammonium acetate.
5. Glycogen suspension (20 mg/mL) (cat. no. 901 393; Boehringer Mannheim).
6. 100% Ethanol.
7. TE buffer: 10 mM Tris-HCl, 1 mM EDTA, pH 8.0.
2.2. Amplification of p53 Exons 4–8
1. Primers (see Table 1).
2. dNTPs (100 mM) (Pharmacia).
3. Dynazyme (Finnzymes) or AmpliTaq (Perkin-Elmer).
4. 10X Dynazyme or AmpliTaq buffer.
5. Sterile water.
SSCP in Exons 4–8 of the TP53 Gene 17
Table 1
Intronic Primers Used in Nonradioactive SSCP for TP53 Exons 4–8a
Exon Location Primers with orientation 5v to 3v
Outer primers
X4 Left ACG TGA ATT CTG AGG ACC TGG TCC TCT GAC
Right ACG TGG ATC CAG AGG AAT CCC AAA GTT CCA
X5 Left ACG TGA ATT CGT TTC TTT GCT GCC GTG TTC
Right ACG TGG ATC CAG GCC TGG GGA CCC TGG GCA
X6 Left ACG TGA ATT CTG GTT GCC CAG GGT CCC CAG
Right ACG TGG ATC CTG GAG GGC CAC TGA CAA CCA
X7 Left ACG TGA ATT CAC CAT CCT GGC TAA CGG TGA
Right ACG TGG ATC CAG GGG TCA GCG GCA AGC AGA
X8 Left ACG TGA ATT CTT GGG AGT AGA TGG AGC CT
Right AGG CAT AAC TGC ACC CTT GG
Inner primers
X4 Left TGC TCT TTT CAC CCA TCT AC
X4A1 Left TGC TCT TTT CAC CCG TCT AC
Right ATA CGG CCA GGC ATT GAA GT
X5 Left TTC AAC TCT GTC TCC TTC CT
X5A5 Left TTG AAC TGT GTC TCG TTC CT
Right CAG CCC TGT CGT CTC TCC AG
X6 Left GCC TCT GAT TCC TCA CTG AT
X6A6 Left GCC TCT GAT TCC TCG CTG AT
Right TTA ACC CCT CCT CCC AGA GA
X7 Left CTT GCC ACA GGT CTC CCC AA
X7A2 Left CTT GCC ACA GCT CTC CCC AA
Right TGT GCA GGG TGG CAA GTG GC
X8 Left TTC CTT ACT GCC TCT TGC TT
X8A1 Left TTC CTT ACT GCC TCG TGC TT
Right CGC TTC TTG TCC TGC TTG CT
a The artificial mutations introduced to positive controls in the second amplification are under-lined and set in boldface. In the second amplification, the left primer is either wild type regular
primer or a changed one (“A” within the code means artificial). The right primer is according to
the unchanged sequence in each case.
2.3. Minigel Analysis and Gel Purification of Amplified Products
1. NuSieve 3 1 agarose (FMC BioProducts, Rockland, ME).
2. 10X Tris-borate EDTA (TBE) buffer: 108 g/L Tris base, 55 g/L boric acid,
4 mL/L 0.5 M EDTA, pH 8.0.
3. Ethidium bromide solution (10 mg/mL) (Research Genetics).
4. DNA size marker (e.g., GelMarker-I, Research Genetics).
5. Bromophenol blue (0.2 mg/50 mL) in 25% Ficoll 400.
6. 0.5 M Ammonium acetate.
18 Vähäkangas, Castrén, and Welsh
For minigels, the Hoefer HE 33 Mini Submarine (Pharmacia Biotech) has
proved quick and practical. For gel purification, the Model 850 gel apparatus
with 8  × 2.5 mm lucite combs from Aquebogue Machine and Repair Shop
(Aquebogue, NY) is used.
2.4. Single-Strand Conformation Polymorphism
1. Pharmacia PhastSystem® equipment.
2. Stop buffer from a Sequenase kit (Sequenase 2.0 from Amersham).
3. Pharmacia reagents for PhastSystem: homogeneous 20% polyacrylamide gels,
PhastGel Native Buffer Strips, PhastGel DNA Silver Staining Kit.
4. Gel-Drying Kit (Promega, Madison, WI).
2.5. Preservation of Gels
1. Clear cellulose film (e.g., gel-drying film; Promega).
2. Gel-drying frames. These are provided by several companies (e.g., Gel-Drying
Kit; Promega).
3. Methods
3.1. Purification of DNA from Paraffin-Embedded Tissue
1. Scrape the tumor tissue into a 1.5-mL conical screw-cap microfuge tube. Add
50 µL of sterile water or TE buffer and break up pieces with a pipet tip.
2. Add 220  µL of water or TE buffer and 30  µL of proteinase K in SDS. Vortex
and incubate in a 37°C heat block for 2 d. After 2 d add another 30 µL of proteinase
solution. If not totally digested after 4 d, another 30 µL can be added for another day.
3. Boil the samples at 100°C for 10 min to inactivate proteinase K.
4. Add 300 µL of phenol, vortex for 20 s, and centrifuge for 2 min. Transfer the
upper (aqueous) layer to a new tube.
5. Add 300 µL of chloroform isoamyl alcohol, vortex for 15 s, and centrifuge for
1 min. Transfer the upper (aqueous) layer to a new tube.
6. Add 100 µL (one-third of volume) of 10 M ammonium acetate, 2 µL of freshly
vortexed glycogen, and 900 µL of cold ethanol (2.5 vol). Vortex and precipitate
DNA at –20°C overnight or at –80°C for at least 6 h.
7. Centrifuge for at least 30 min at high speed (12,800g) in the cold (4°C), wash
pellets once with 70% ethanol for 10–15 min, centrifuge for 2 min, and decant
the supernatant carefully.
8. Dry the pellets in a 42°C heat block for 10 min and dissolve the pellet in
50–100 µL of water.
3.2. Amplification of TP53 Exons
This method is only specific for the primers in Table 1. For a new strand,
new standards must be developed and the experimental conditons optimized.
All reagents should be sterile and reactions done under a laminar flow hood
to avoid contamination. Dedicated pipets for this purpose should be used for
SSCP in Exons 4–8 of the TP53 Gene 19
amplification reagents and a different pipet used for the template. Also, pipet tips
containing an aerosol barrier are important to reduce contamination. A negative
control without a template will confirm the purity of reagents and the procedure.
Regarding polymerase, we have found that Dynazyme and AmpliTaq work well
for us. Two consecutive amplifications are carried out, first with outer primers
and then with inner (nested) primers ([40]; primer sequences in Table 1).
1. Dilute the dNTPs by adding 250  µL of each into 12.33 mL of sterile water.
Divide into about 1-mL portions and store at –20°C.
2. Prepare a mixture of all the reagents for the first polymerase chain reaction
(PCR) (without the enzyme or template). The following mixture is for one reaction: 10 µL
of the 10X enzyme buffer (according to the enzyme used), 16 µL of dNTPs (final
concentration 300 µM), 20 pmol of each of the outer primers, H2O to give 100 µL.
3. Prepare at the same time a mixture (with the enzyme this time) for the second
PCR. The following mixture is for one reaction: 20 µL of 10X enzyme buffer,
32 µL of dNTPs, 40 pmol of each of the inner primers, 1 µL of enzyme, H2O to
give 200 µL. Aliquot 200 µL in each tube and store refrigerated until the first-round reactions are completed.
4. Divide 100 µL of the mixture for the first PCR into the amplification tubes.
5. Add 2–5 µL of template.
6. Heat at 100°C for 10 min and keep at 85°C in a heat block until placed into the
thermocycler.
7. Add 0.5 µL of enzyme.
8. Place tubes in a preheated 95°C thermocycler.
9. Amplify for 35 cycles (94°C for 1 min, 60°C for 1 min, 78°C for 30 s).
10. Add 5 µL from the first reaction tubes to the second tubes as templates. There
should be two controls: one with 5 µL from the control from the first reaction
series as a template, and another with no template to control for the reagents in
the second series (see Note 1).
11. Amplify for another 35 cycles as in step 9.
3.3. Minigel Analysis of Amplified DNA
Prepare a 4% gel (NuSieve 3 1 agarose) with 0.1% ethidium bromide in 1X
TBE buffer and run 5 µL of samples + 3 µL bromophenol blue, and 10 µL of
controls + 4 µL of bromophenol blue, and run with DNA size markers (a marker
with a band at 100–200 is needed) for about 30 min at 160–180 V. Table 2
gives the sizes of the amplified products.
3.4. Gel Purification of Amplified Exons
1. Prepare a 4% gel, about 1 cm thick, with deep wells to hold the 200-µL amplified
product.
2. Add 25 µL of bromophenol blue to the sample.
3. Run the samples for about 2 h at 125 V.
4. View the gel in long-wave UV light to visualize the bands.
20 Vähäkangas, Castrén, and Welsh
5. Using a sterile, disposable scalpel, cut the bands very closely according to known
controls. Cutting closely is critical, because any extra material will show as extra
bands in SSCP.
6. Cut the gel piece into smaller pieces, place in an Eppendorf tube, and add 500 µL
of ammonium acetate.
7. Let the tube rock at 37°C overnight.
8. The next morning carefully pipet the buffer into another tube, add 1 mL of 95%
ethanol, and precipitate at –20°C for about 24 h.
9. Centrifuge at 4°C for 30 min at 12,800g and carefully decant the ethanol.
10. Wash DNA once; gently add 1 µL of cold 70% ethanol to each tube, let sit at
room temperature for 5 min, centrifuge at 12,800g for 3 min, and carefully decant
the ethanol.
11. Let the tube stand open at room temperature (for about 30 min) to evaporate any
remaining ethanol.
12. Dissolve the pellet in 20–50 µL of H2O, depending on the intensity of the DNA
band in the minigel.
13. Store refrigerated, or at –20°C for long-term storage.
3.5. Separation of DNA Strands in PhastSystem
Two temperatures are used for each exon. The lower temperature is 4°C for
all exons and the higher is 20°C for exons 5–8. However, the higher running
temperature for exon 4 is 15°C for the best detection of the codon 72 polymor-phism. This polymorphism is not detectable at 4°C (41) .
1. Program the machine. Select a different method number for each of the tempera-tures (4°C, 15°C, and 20°C) and program according to the following conditions:
a. Prerun: 400 V, 10 mA, 1 W, and 100 average volt-hours (AVh).
b. Sample application: 25 V, 10 mA, 1 W, and 2 AVh.
c. Run: 200 V, 5 mA, 1 W, and 800 AVh.
2. Mix 3 µL of sample and 3 µL of stop buffer from a Sequenase kit (see Note 3).
Table 2
Amplification Products and Their Running
Conditions in SSCP of TP53 Exons 4–8
Running times at different temperaturesa
TP53 exon Size of product 4°C15°C20°C
Exon 4 353 800 AVh          700 AVh
Exon 5 247 600 AVh 500 AVh
Exon 6 180 400 AVh 300 AVh
Exon 7 195 500 AVh 400 AVh
Exon 8 200 600 AVh 500 AVh
aAVh, average V-h.
SSCP in Exons 4–8 of the TP53 Gene 21
3. Set the temperature in the PhastSystem and let it settle.
4. Prepare the gels by bending one corner of the tab (for later removal of the gel),
and cutting one corner from one gel (to distinguish left and right gels). Clean the
gel bed with distilled water and pipet about 0.5 mL of water onto the gel area to
keep the gel properly positioned during the run. Carefully position the gel along
the lines on the gel bed and blot (do not wipe) excess water with a paper towel.
Be sure that no air bubbles remain under the gel. Correct the position if needed.
5. Remove the cellophane covering the gel and save it for later use.
6. Replace the buffer strip holder carefully, so as not to disturb the gel. After the
buffer strip holder is in place, insert the two buffer strips. Ensure the even contact
between the buffer and the gel by applying light pressure over the strips.
7. Lower the sample applicator arm. Ensure an even contact between the electrode
and the buffer strip again by applying light pressure over the electrodes. Close
the chamber.
8. Prerun the gels for buffer equilibration.
9. When the AVh reaches 80 (about 5 min before the sample applicator drops on the
gel), boil the samples at 100°C for 4 min and put on ice. Apply 1 µL of each sample
in the grooves of the eight-well comb (note that the ends of the comb do not have
grooves).
10. Place the comb carefully in its place immediately after the sample applicator has
dropped. This has to be done between 100 and 101 AVh (about 4 min) before the
sample applicator rises up again.
11. Stop the run by pressing the button “SEP start/stop” and then “DO” at the AVh
according to the exon and temperature (Table 2) (see Note 3).
3.6. Silver Staining of Gels in PhastSystem
Program and store the developing method prepared according to the method
optimized for native-polyacrylamide gel electrophoresis with PhastGel gradi-ent media. (See table 3 in the PhastGel™ Silver Kit Instruction Manual. Steps
1 and 16, however, are not needed for this protocol.) Prepare the reagents for
silver staining while running the gels (see Note 4).
1. For step 2, make up 100 mL of 50% ethanol + 10% acetic acid in deionized water.
2. For steps 3, 4, 6, and 7, make up a total of 500 mL of 10% ethanol + 5% acetic acid.
3. For steps 8, 9, 11, and 12, set aside 1000 mL of deionized water.
4. For steps 13 and 14 (developer), pipet one ampoule of 2% formaldehyde into one
bottle of sodium carbonate (both of these are provided in the kit). Mix thoroughly.
5. For step 15, prepare the background reducer by adding one packet of sodium
thiosulfate into 100 mL of deionized water. When dissolved, adjust the pH to
5.0–6.0 by adding about one drop of 10% acetic acid.
6. For step 5 the 5% glutaraldehyde and for step 10 the silver stain are ready to use.
7. Place tubes into correct solutions (as defined in table 3 in the instruction manual).
8. For the run, place the two gels, facing each other, in the developing chamber in
the gel holders. Close the lid tightly and start the developing program. The pro-
22 Vähäkangas, Castrén, and Welsh
gram takes about 1 h to run. For easier handling, when staining is finished,
remove the gel and replace the cellophane on the gel.
3.7. Documentation and Preservation of Gels
Documentation of the gels is essential for later analysis, because silver stain
fades with time. This may happen within a few days in daylight. We recom-mend both immediate photography and preservation of the gels (see Note 5).
1. For preservation of the gels, remove the cellophane and soak the gel in deionized
water for 1–2 min.
2. Prewet the clear cellulose film with deionized water.
3. Place the gel in between the wet cellulose sheets and press out any air bubbles.
4. Secure the films tightly in the frame using clamps or bulldog clips.
5. Allow to dry completely at room temperature overnight. If properly done, the
film is sealed and can be cut out with about a 0.5-cm frame for storage.
3.8. Interpretation of Results
To interpret results, the bands of the samples are compared with those of
wild-type controls along the whole gel (Fig. 1). Double-stranded DNA runs
Fig. 1. Band patterns of p53 exons in SSCP. In exon 4, the band patterns for codon
72 polymorphism are shown: a, arginine; p, proline; a/p, heterozygote. The codes for
mutated bands refer to strands amplified using exons listed in Table 1; wt, wild-type
sequence.
SSCP in Exons 4–8 of the TP53 Gene 23
off from the bottom of the gel and does not interfere with the interpretation of
these conditions. Positive results in the analysis can result from a sample with
a mutation, an amplified product with a polymerase error, or misinterpretation
of SSCP owing to, e.g., an incompletely purified amplified sample. Very faint
bands are usually owing to differences in the purification procedure from one
time to another. If positive or suspicious, it is necessary to analyze another
amplified product from the same sample.
4. Notes
1. Because there can be a slight variation among different machines, it is important
to include both negative and positive controls on each gel. Each new set of con-trols must be confirmed to be wild type, either by comparison in the same SSCP
gel with older known controls or by sequencing. This must be done for two
reasons: (1) in each reaction there is a possibility of a polymerase error, and
(2) apparently healthy people may carry a germline p53 mutation.
2. It is not necessary to know the DNA concentration exactly. However, note that as
with sequencing, more is not necessarily better. The optimal concentration of
DNA in the final sample, according to our experience, is 1–10 ng/µL.
3. After every use of the PhastSystem (which includes the separation by electro-phoresis and silver staining), it is essential to run a cleaning program in the
developing chamber. This is best done immediately after each run to avoid
reagents drying in the tubes. Do not forget to place a blank plastic gel support
(which is usually provided with the machine; if not, dedicate one of the unused
gels for this purpose) in the gel holder in the chamber during the cleaning run to
avoid overflowing of the chamber. The PhastSystem will not run without the
blank gel support. After the cleaning program, careful manual cleaning of the
staining chamber with soft paper including the level sensor (by cotton tip appli-cators) is still necessary. The electrodes should be cleaned with a soft brush.
Without these cleaning procedures, dark background in the gels is inevitable and
the problem grows worse with time. The tubing should be changed at least twice
a year if the system is used regularly, and more frequently for heavier use.
4. Although the shelf life of the reagents is usually long (6 mo), it is better not to
store them for a long time at the laboratory, but rather to use fresh reagents.
5. A practical way of storing the gels after drying is to use a plastic pocket sheet for
slides. The dried gels within the cellophane sheets can be cut out to fit the pock-ets. Although the bands will stay visible for at least 1 yr if stored properly, it is
advisable to photograph at least the key experiments. Dried gels break easily and
careful handling is necessary.
Acknowledgments
We thank Mohammed Aslam Khan and Anna-Kaisa Lappi for critical read-ing of the manuscript.
24 Vähäkangas, Castrén, and Welsh
References
1. Hollstein, M., Sidransky, D., Vogelstein, B., and Harris, C. C. (1991) p53 muta-tions in human cancers. Science 253, 49–53.
2. Greenblatt, M. S., Bennett, W. P., Hollstein, M., and Harris, C. C. (1994) Muta-tions in the p53 tumor suppressor gene: clues to cancer etiology and molecular
pathogenesis. Cancer Res. 54, 4855–4878.
3. Hainaut, P., Soussi, T., Shomer, B., Hollstein, M., Greenblatt, M., Hovig, E., Har-ris, C. C., and Montesano, R. (1997) Database of p53 gene somatic mutations in
human tumors and cell lines: updated compilation and future prospects. Nucleic
Acids Res. 25, 151–157.
4. Hussain, S. P. and Harris, C. C. (1998) Molecular epidemiology of human cancer:
contribution of mutation spectra studies of tumor suppressor genes. Cancer Res.
58, 4023–4037.
5. Gottlieb, T. M. and Oren, M. (1996) p53 in growth control and neoplasia. Biochim.
Biophys. Acta 1287, 77–102.
6. Hernandez-Boussard, T. M. and Hainaut, P. (1998) A specific spectrum of p53
mutations in lung cancer from smokers: review of mutations compiled in the IARC
p53 database. Environ. Health Perspect. 106, 385–391.
7. Brash, D. E., Rudolph, J. A., Simon, J. A., Lin, A., McKenna, G. J., Baden, H. P.,
Halperin, A. J., and Ponten, J. (1991) A role for sunlight in skin cancer:
UV-induced p53 mutations in squamous cell carcinoma.  Proc. Natl. Acad. Sci.
USA 88, 10,124–10,128.
8. Ozturk, M. (1991) p53 mutation in hepatocellular carcinoma after aflatoxin expo-sure. Lancet 338, 1356–1359.
9. Kastan, M. B., Onyekwere, O., Sidransky, D., Vogelstein, B., and Craig, R. W.
(1991) Participation of p53 protein in the cellular response to DNA damage. Can-cer Res. 51, 6304–6311.
10. Lane, D. P. (1992) p53, guardian of the genome. Nature 358, 15,16.
11. Harris, C. C. (1996) The 1995 Walter Hubert Lecture—molecular epidemiology
of human cancer: insights from the mutational analysis of the p53 tumour-suppressor gene. Br. J. Cancer 73, 261–269.
12. Perera, F. P., Whyatt, R. M., Jedrychowski, W., Rauh, V., Manchester, D.,
Santella, R. M., and Ottman, R. (1998) Recent developments in molecular epide-miology: a study of the effects of environmental polycyclic aromatic hydrocar-bons on birth outcomes in Poland. Am. J. Epidemiol. 147, 309–314.
13. Hainaut, P. and Vähäkangas, K. (1997) p53 as a sensor of carcinogenic expo-sures: mechanisms of p53 protein induction and lessons from p53 gene mutations.
Pathologie-Biologie 45, 833–844.
14. Bergh, J., Norberg, T., Sjögren, S., Lindgren, A., and Holmberg, L. (1995) Com-plete sequencing of the p53 gene provides prognostic information in breast cancer
patients, particularly in relation to adjuvant systemic therapy and radiotherapy.
Nat. Med. 1, 1029–1034.
15. Borresen, A.-L., Anderson, T. I., Eyfjörd, J. E., Cornelis, R. S., Thorlacius, S.,
Borg, Å., Johansson, U., Theillet, C., Scherneck, S., Hartman, S., Cornelisse, C. J.,
SSCP in Exons 4–8 of the TP53 Gene 25
Hovig, E., and Devilee, P. (1995) TP53 mutations and breast cancer prognosis:
particularly poor survival rates for cases with mutations in the zinc-binding
domains. Genes Chromosomes Cancer 14, 71–75.
16. Valgardsdottir, R., Tryggvadottir, L., Steinarsdottir, M., Olafsdottir, K.,
Jonasdottir, S., Jonasson, J. G., Ögmundsdottir, H. M., and Eyfjörd, J. E.
(1997) Genomic instability and poor prognosis associated with abnormal TP53
in breast carcinomas: molecular and immunohistochemical analysis.  APMIS
105, 121–130.
17. Soini, Y., Turpeenniemi-Hujanen, T., Kamel, D., Autio-Harmainen, H., Risteli,
J., Risteli, L., Nuorva, K., Pääkkö, P., and Vähäkangas, K. (1993) p53 Immuno-histochemistry in transitional cell carcinoma and dysplasia of the urinary bladder
correlates with disease progression. Br. J. Cancer 68, 1029–1035.
18. Uchida, T., Wada, C., Ishida, H., Wang, C., Egawa, S., Yokoyama, E., Kameya,
T., and Koshiba, K. (1999) p53 mutations and prognosis in bladder tumors.
J. Urol. 153, 1097–1104.
19. Peller, S., Halperin, R., Schneider, D., Kopilova, Y., and Rotter, V. (1999) Poly-morphisms of the p53 gene in women with ovarian or endometrial carcinoma.
Oncol. Rep. 6, 193–197.
20. Wang, Y. C., Chen, C. Y., Chen, S. K., Chang, Y. Y., and Lin, P. (1999) p53
codon 72 polymorphism in Taiwanese lung cancer patients: association with lung
cancer susceptibility and prognosis. Clin. Cancer Res. 5, 129–134.
21. Murata, M., Tagawa, M., Kimura, H., Kakisawa, K., Shirasawa, H., and Fujisawa,
T. (1998) Correlation of the mutation of p53 gene and the polymorphism at codon
72 in smoking-related non-small cell lung cancer patients.  Int. J. Oncol. 12,
577–581.
22. Murata, M., Tagawa, M., Kimura, M., Kimura, H., Watanabe, S., and Saisho, H.
(1996) Analysis of a germ line polymorphism of the p53 gene in lung cancer
patients: discrete results with smoking history. Carcinogenesis 17, 261–264.
23. Storey, A., Thomas, M., Kalita, A., Harwood, C., Gardiol, D., Mantovani, F.,
Breuer, J., Leigh, I. M., Matlashewski, G., and Banks, L. (1998) Role of a p53
polymorphism in the development of human papillomavirus-associated cancer.
Nature 393, 229–234.
24. Thomas, M., Kalita, A., Labreque, S., Pim, D., Banks, L., and Matlashewski, G.
(1999) Two polymorphic variants of wild-type p53 differ biochemically and bio-logically. Mol. Cell Biol. 19, 1092–1100.
25. Storey, A., Thomas, M., Kalita, A., Harwood, C., Gardiol, D., Mantovani, F.,
Breuer, J., Leigh, I. M., Matlashewski, G., and Banks, L. (1998) p53 polymor-phism and risk of cervical cancer. Nature 396, 532.
26. Rosenthal, A. N., Ryan, A., Al-Jehani, R. M., Storey, A., Harwood, C. A., and
Jacobs, I. J. (1998) p53 codon 72 polymorphism and risk of cervical cancer in
UK. Lancet 352, 871, 872.
27. Helland, A., Langerod, A., Johnsen, H., Olsen, A. O., Skovlund, E., and Borresen-Dale, A.-L. (1998) p53 polymorphism and risk of cervical cancer.  Nature 396,
530,531.
26 Vähäkangas, Castrén, and Welsh
28. Josefsson, A. M., Magnusson, P. K. E., Ylitalo, N., Quarforth-Tubbin, P., Ponten,
J., Adami, H. O., and Gyllensten, U. B. (1998) p53 polymorphism and risk of
cervical cancer. Nature 396, 531.
29. Hildesheim, A., Schiffman, M., Brinton, L. A., Fraumeni, J. F. Jr., Herrero, R.,
Bratti, M. C., Schwartz, P., Mortel, R., Barnes, W., Greenberg, M., McGowan, L.,
Scott, D. R., Martin, M., Herrera, J. E., and Carrington, M. (1998) p53 polymor-phism and risk of cervical cancer. Nature 396, 531,532.
30. Minaguchi, T., Kanamori, Y., Matsushima, M., Yoshikawa, H., Taketani, Y., and
Nakamura, Y. (1998) No evidence of correlation between polymorphism at codon
72 of p53 and risk of cervical cancer in Japanese patients with human
papillomavirus 16/18 infection. Cancer Res. 58, 4585,4586.
31. Orita, M., Iwahana, H., Kanazawa, H., Hayashi, K., and Sekiya, T. (1989) Detec-tion of polymorphisms of human DNA by gel electrophoresis as single strand
conformation polymorphisms. Proc. Natl. Acad. Sci. USA 86, 2766–2770.
32. Chaubert, P., Beautista, D., and Benhattar J. (1993) An improved method for rapid
screening of DNA mutations by nonradioactive single-strand conformation poly-morphism procedure. BioTechniques 15, 586.
33. Cheng, J. and Haas, M. (1992) Sensitivity of detection of heterozygous point
mutations in p53 cDNAs by direct PCR sequencing.  PCR Methods Appl. 1,
199–201.
34. Wu, J. K., Ye, Z., and Darras, B. T. (1993) Sensitivity of single-strand conforma-tion polymorphism (SSCP) analysis in detecting p53 point mutations in tumors
with mixed cell populations. Am. J. Hum. Genet. 52, 1273–1275.
35. Mohabeer, A. J., Hiti, A. L., and Martin, W. J. (1991) Non-radioactive single-strand conformation polymorphism (SSCP) using the Pharmacia ‘PhastSystem’.
Nucleic Acid Res. 19, 3154.
36. Neubauer, A., Brendel, C., Vogel, D., Schmidt, C. A., Heide, I., and Huhn, D.
(1993) Detection of p53 mutations using nonradioactive SSCP analysis: p53 is
not frequently mutated in myelodysplastic syndromes (MDS). Ann. Hematol. 67,
223–226.
37. Bosari, S., Marchetti, A., Buttitta, F., Graziani, D., Borsani, G., Loda, M.,
Bevilacqua, G., and Coggi, G. (1995) Detection of p53 mutations by single-strand
conformation polymorphisms (SSCP) gel electrophoresis. Diagn. Mol. Pathol. 4,
249–255.
38. Kurvinen, K., Hietanen, S., Syrjänen, K., and Syrjänen, S. (1995) Rapid and
effective detection of mutations in the p53 gene using nonradioactive single-strand
conformation polymorphism (SSCP) technique applied on PhastSystem™.
J. Virol. Methods 51, 43–54.
39. Welsh, J.A., Castrén, K., and Vähäkangas, K. (1997) Single-strand conformation
polymorphism analysis to detect p53 mutations: characterization and develop-ment of controls. Clin. Chem. 43, 2251–2255.
40. Lehman, T. A., Bennett, W. P., Metcalf, R. A., Welsh, J. A., Ecker, J., Modali, R. V.,
Ullrich, S., Romano, J. W., Appella, E., Testa, J. R., Gerwin, B. I., Vogelstein, B.,
SSCP in Exons 4–8 of the TP53 Gene 27
and Harris, C. C. (1991) p53 mutations, ras mutations, and p53-heat shock 70
protein complexes in human lung carcinoma cell lines.  Cancer Res. 51,
4090–4096.
41. Castrén, K., Vähäkangas, K., Heikkinen, E., and Ranki, A. (1998) Absence of p53
mutations in benign and pre-malignant male genital lesions with overexpressed
p53 protein. Int. J. Cancer 77, 674–678.
CFLP Analysis for Genotyping and Mutation Detection 29
29
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
5
Cleavase® Fragment Length Polymorphism
Analysis for Genotyping and Mutation Detection
Laura Heisler and Chao-Hung Lee
1. Introduction
DNA sequencing is the gold standard in DNA diagnostics and is the only
absolute means of establishing the identity of a new mutation. However, the
clinical cost of obtaining this information is often prohibitive, particularly when
large DNA fragments are interrogated for the presence of any of a number of
either known or previously undescribed genetic alterations (1) . Instead, several
mutation scanning methods have been developed to eliminate the need to
sequence every nucleotide when it is only the precise identity of one or a few
nucleotides that is clinically significant. Until now such methods have pro-vided only a “yes” or “no” answer in determining whether a test sample differs
from a known reference. Relatively few methods have been proven capable of
unambiguously identifying unique nucleic acid variants, particularly when mul-tiple sequence changes occur (2) . Consequently, the majority of existing muta-tion scanning methods are unsuitable for PCR-based genotyping applications
in which regions of sequence variability are used to categorize isolates for their
similarities to known variants.
Third Wave Technologies has pioneered a novel mutation and polymor-phism screening method that accurately and precisely distinguishes nucleic acid
variants (3) . This approach relies on enzymatic cleavage of characteristic struc-tures formed by single-stranded nucleic acids. On sequential denaturation and
renaturation, both single-stranded DNA and RNA molecules assume three-dimensional conformations that are a precise reflection of their nucleic acid
sequences (4). This principle is the foundation of several mutation scanning
techniques, such as single-strand conformation polymorphism (SSCP) and
dideoxy fingerprinting (5 ,6) . Instead of relying on direct observation of these
30 Heisler and Lee
structures, e.g., by noting subtle differences in how different DNA strands
migrate through nondenaturing gels, the Third Wave Technologies’ enzyme-based approach uses a structure-specific endonuclease engineered from the
5vnuclease domain of Taq DNA polymerase, dubbed the Cleavase® I enzyme,
to cut DNA strands wherever these structures occur (3)  (Fig. 1). By analogy to
restriction fragment length polymorphism analysis, Third Wave has named
their method Cleavase Fragment Length Polymorphism (CFLP®) analysis.
The Cleavase I enzyme rapidly and specifically cleaves these structures,
many of which are formed on a given DNA fragment, albeit transiently, in
equilibrium with alternative, mutually exclusive structures. The CFLP method
is thus able to elucidate a considerable amount of information about the
sequence content of a DNA fragment without relying on cumbersome high-resolution analysis of each base. Each unique DNA sequence produces a dis-tinctive collection of structures that, in turn, results in the generation of a
singular fingerprint for that sequence. This capability makes the CFLP tech-nology suitable for diverse mutation scanning applications, including
genotyping (1 ,3 ,7–14) . Furthermore, the CFLP reaction is unaffected by the
length of the DNA fragment and can be used to analyze far longer stretches
of DNA than is currently possible with other methods, up to at least 2.7 kb
(unpublished data).
1.2. Visualizing Sequence Differences in CFLP Fingerprints
The CFLP method comprises the steps of separation of DNA strands by
heating, formation of intrastrand structures on cooling with rapid enzymatic
cleavage of these structures before they are disrupted by reannealing of the
complementary strands, and separation and visualization of the resulting “struc-tural fingerprint” (Fig. 2). When closely related DNA fragments, such as a
Fig. 1. Structures recognized by the Cleavase I enzyme. The Cleavase I enzyme is
a structure-specific nuclease that recognizes the junctions between single- and double-stranded regions of nucleic acids, i.e., so-called hairpins or stem loops. Cleavage
occurs on the 5vside of such structures (see arrow). These intrastrand structures are
formed when nucleic acid molecules are denatured and then allowed to cool.
CFLP Analysis for Genotyping and Mutation Detection 31
wild-type and a mutant version of a gene, are compared, the CFLP fingerprints
exhibit strong familial resemblance to one another such that they share the
majority of bands produced. The sequence differences are revealed as changes
in one or several bands. These unique band changes are manifest as mobility
shifts, the appearance or disappearance of bands, and/or significant differences
in band intensity (Fig. 3).
Fig. 2. CFLP reaction. The CFLP reaction itself is a simple three-step procedure
that relies on the use of temperature to change some of the physical characteristics of
DNA molecules.
32 Heisler and Lee
Fig. 3. CFLP analysis of the ITS regions of P. carinii. PCR products spanning the
ITS1 and ITS2 regions of P. carinii, 534 bp long and labeled on the 5vends of both
strands with TET, were subjected to CFLP analysis. Approximately 250 fmol of
labeled PCR product was analyzed in the “ramped” reactions and approx 150 fmol in
the single temperature reactions. The DNA aliquots were supplemented with DNA
dilution buffer. The ramping assay was performed as described in Subheading 3.2.
The genotypes of the samples from which the DNA was amplified are indicated above
the lanes. Lanes marked “mw” contain molecular weight markets with fragment sizes
as indicated. The gel was electrophoresed at constant wattage (20 W) until a bro-mophenol blue market dye, loaded in a far lane (not shown), reached the bottom of the
gel. The gel cassette was scanned on a Hitachi FMBIO-100 fluorescence imager with
a 585-nm emission filter.
CFLP Analysis for Genotyping and Mutation Detection 33
CFLP analysis is exquisitely sensitive to the presence of minor sequence
variations and can detect changes involving one or more bases, including mis-sense mutations, with >95% sensitivity and 100% specificity. Because the
CFLP method results in an easily examined pattern, rather than base-by-base
analysis of each sequence, the value of this approach may be even more pro-nounced in genotyping applications. In these cases, what is sought is the rapid
identification of compound sequence variations occurring throughout an
amplified fragment. Rapid inspection of the patterns generated by CFLP analy-sis of fragments containing multiple, dispersed base changes has proven to be
an effective approach to classifying bacterial and viral sequences according to
genotype (1 ,3 ,11) .
Pneumocystis carinii f. sp. hominis is the leading cause of pneumonia and
the most commonly transmitted life-threatening opportunistic infection among
AIDS patients  (15) . To establish the origin of particular infections, verify
localized outbreaks, and determine whether an individual has sustained mul-tiple, coincident infections, researchers have attempted to classify individual
P. carinii strains based on sequence variability among isolates (16) . Sequence
variation in the internal transcribed spacer (ITS) regions of the rRNA genes of
P. carinii can be used for such genotypic identification  (17) . The region
located between the 18S and 5.8S rRNA genes is called ITS1, and that between
the 5.8S and 26S rRNA genes is ITS2. Among the two regions, approx 60
different ITS sequences have been characterized by direct DNA sequencing
(18) . Sequence variation occurs throughout these 161- and 192-bp regions,
respectively, and the majority of sequence changes within each ITS have been
determined to be significant in establishing type (18) .
The suitability of the CFLP scanning method for differentiating sequences
in the ITS region of five cloned  P. carinii sequences belonging to different
types was investigated. The ITS region was amplified by polymerase chain
reaction (PCR), and the 5v ends of both strands were labeled by using
tetrachlorofluorescein (TET) sense strand labeled primers (see Note 1). The
amplified products were purified and then partially digested with the Cleavase
I enzyme. The samples were analyzed in duplicate sets, one of which was sub-jected to CFLP digestion at a predetermined, optimized reaction temperature,
whereas the other was digested under conditions in which the temperature was
continually increased, or “ramped” (see Subheading 3.2. and Note 2).
The results indicate that the CFLP method is highly effective in reproduc-ibly distinguishing different P. carinii types (Fig. 3). An inspection of the fin-gerprints generated from these samples reveals a high degree of similarity
overall, indicative of the fact that only a few bases are altered in the variants,
with some marked differences that reflect those base changes. In Fig. 3, there
are several examples of bands that appear in some lanes but that are absent in
34 Heisler and Lee
others, as well as bands that appear shifted in some lanes relative to others.
Unique bands, indicated by arrows, are apparent, e.g., in lanes Bm. In other
cases, the most notable difference is a composite shifting downward of a sub-stantial portion of the pattern, indicative of a small deletion, such that the frag-ments are shortened relative to the labeled 5vends (e.g., lanes Ed and Di, as
indicated by brackets).
Note that the patterns generated by the ramping procedure appear to be
enhanced relative to the single temperature procedure in several places. In par-ticular, note the appearance of additional bands between 82 and 118 bp. This
improvement in the richness of the patterns is likely due to the fact that certain
substrate structures may not be as favored at a single temperature, as is used in
the conventional approach, but rather emerge as the temperature changes over
the course of the ramping reaction. In some cases the ramping approach not
only eliminates the need for preliminary optimization steps but may also serve
to improve the sensitivity of the CFLP method.
2. Materials
2.1. Preparation of End-Labeled PCR-Amplified Fragments
2.1.1. PCR Reagents
1. Sterile double-distilled H2O.
2. 10X PCR buffer: 500 mM KCl, 100 mM Tris-HCl, pH 9.0, 15 mM MgCl2.
3. dNTP (deoxynucleotide) mix: 0.2 mM each dNTP in sterile double-distilled H2O.
4. Mineral oil or wax overlay.
5. Oligonucleotide primers at a concentration of 10  µM, at least one of which is
labeled with a fluorescent dye (e.g., tetrachlorofluorescein, fluorescein) or a moi-ety detectable by chemiluminescence (e.g., biotin, digoxyigenin).
2.1.2. Post-PCR Fragment Purification
1. Exonuclease I: Available at 10 U/µL from Amersham Pharmacia Biotech
(Arlington Heights, IL), cat. no. E70073Z, or at 20 U/µL from Epicentre Tech-nologies (Madison, WI), cat. no. X40505K.
2. High Pure PCR Product Purification Kit (HPPPPK), available from Roche
Boehringer Mannheim Biochemical (Indianapolis, IN), cat. no. 1732668.
3. Sterile double-distilled H2O or T10E0.1: 10 mM Tris-HCl, pH 8.0; 0.1 mM EDTA,
pH 8.0.
2.2. CFLP Analysis
1. Cleavase I enzyme (25 U/µL) in Cleavase enzyme dilution buffer: 20 mM Tris-HCl, pH 8.0, 50 mM KCl, 0.05% Tween®-20, 0.05% Nonidet™ P-40, 100 µg/mL
of bovine serum albumin, and 50% (v/v) glycerol.
2. DNA dilution buffer: 5 mM MOPS, pH 7.5.
CFLP Analysis for Genotyping and Mutation Detection 35
3. 10X CFLP buffer: 100 mM MOPS, pH 7.5, 0.5% Tween-20, 0.5% Nonidet P-40.
4. 2 mM MnCl2.
5. 10 mM MgCl2.
6. Stop solution (for nonfluorescent gel-based detection): 95% formamide, 10 mM
EDTA, pH 8.0, 0.05% xylene cylanol, 0.05% bromophenol blue (see Note 3).
7. Stop solution (for fluorescent gel-based detection): 95% formamide, 10 mM
EDTA, pH 8.0, 0.05% crystal violet (see Note 3).
8. Sterile double-distilled H2O.
9. Thin-walled reaction tubes (200 or 500 µL).
2.3. Gel Electrophoresis
1. Gel solution: 6–10% acrylamide bis (19 1) solution, 7 M urea, 0.5X Tris-borate
EDTA (TBE) buffer.
2. 0.5X TBE gel running buffer (pH 8.3): 44.5 mM Tris, 44.5 mM borate, 1 mM
EDTA, pH 8.0.
3. Ammonium persulfate (10% [w/v]).
4. TEMED.
5. Teflon flat-bottomed combs and spacers (0.5 mM thick) (2) .
6. Glass plates (20 × 20 cm), nonfluorescing for use with fluorescence imager or
standard for chemiluminescence detection.
7. Gel electrophoresis support.
8. Power supply capable of supplying up to 2000 V.
2.4. Visualization of CFLP Patterns
2.4.1. Fluorescence Detection
1. Hitachi FMBIO®-100 Fluorescent Method Bio-Image Analyzer (Hitachi Soft-ware, San Bruno, CA) or Molecular Dynamics 595 FluorImager™ (Molecular
Dynamics, Sunnyvale, CA).
2. Lint-free laboratory wipes.
3. Lens paper.
4. Nonfluorescing detergent, e.g., RBS 35 Detergent Concentrate (Pierce, Rock-ford, IL).
2.4.2. Chemiluminescence Detection
1. 10X SAAP: 1 M NaCl, 0.5 M Tris-base, pH 10.0.
2. 1X SAAP, 0.1% sodium dodecyl sulfate (SDS): 100 mM NaCl, 50 mM Tris-Base, pH 10.0, 0.1% SDS (w/v).
3. 1X SAAP, 1 mM MgCl2: 100 mM NaCl, 50 mM Tris-base, pH 10.0, 1 mM MgCl2.
4. Sequenase Images™ 5X Blocking Buffer (cat. no. US75354; Amersham
Pharmacia Biotech).
5. Streptavidin-Alkaline-Phosphatase Conjugate (cat. no. US11687; Amersham
Pharmacia Biotech).
36 Heisler and Lee
6. CDP-Star™ substrate (cat. no. MS250R; Tropix, Bedford, MA).
7. Isopropanol.
8. Latex gloves (powder free).
9. X-ray film.
10. Positively charged nylon membrane, pore size 0.2 µm (e.g., Nytran® Plus Mem-brane, Schleicher and Schuell, Keene, NH).
11. Blotting paper (20 × 20 cm) (cat. no. 28303-100; VWR Scientific).
12. Sealable plastic bags.
13. Forceps.
14. Small plastic containers for processing membranes.
15. Darkroom/film-developing facilities.
16. Permanent laboratory marker.
3. Methods
3.1. Purification of PCR-Generated Fragments (see Note 1)
PCR amplification should be performed according to established protocols
for the particular locus in question. When PCR products are visualized by gel
electrophoresis followed by sensitive detection of the label to be used to visu-alize CFLP products, contamination by labeled primers and prematurely trun-cated single-stranded PCR products is evident. These contaminating DNA
species are effectively removed by the procedures noted. In particular, the
HPPPPK procedure has been proven effective for eliminating lower molecular
weight (i.e., >100 bp) DNA species, whereas Exonuclease I digestion is effec-tive for removing larger DNA species. An alternative to the HPPPPK columns
in conjunction with Exonuclease I digestion is to precipitate with 1 vol of iso-propanol following Exonuclease I digestion.
If a single, labeled product is detected following PCR and HPPPPK and
Exonuclease I digestion, then proceed with CFLP analysis. If more than one
product is detected, then optimization of the PCR reaction or gel isolation of
the desired product is necessary. The following protocol describes the method
for Exonuclease I digestion:
1. Following PCR amplification, incubate the reaction mixture at 70°C for 10 min.
2. Bring the reaction mixture to 37°C.
3. Add 1 U of Exonuclease I/µL of original PCR reaction mixture (e.g., 100 U to a
100-µL reaction mixture).
4. Incubate for 30 min at 37°C.
5. Inactivate the reaction by heating at 70°C for 30 min.
6. Following Exonuclease I digestion, apply the reaction mixtures to the HPPPPK
spin columns according to the manufacturer’s suggested procedures. The sup-plied elution buffer should be replaced with either sterile double-distilled H2O or
T10 E0.1, pH 8.0.
CFLP Analysis for Genotyping and Mutation Detection 37
3.2. Preparation and Performance of CFLP Reactions
Prior to performing CFLP analysis, it is strongly recommended that the qual-ity and quantity of the PCR-generated fragments following purification be
checked. This can be done by visualizing the label used (i.e., by fluorescence
analysis or chemiluminescence detection) on an aliquot of the DNA in a small
denaturing polyacrylamide gel.
As seen in Fig. 3, there are two alternative approaches to be taken in config-uring the CFLP reaction. The initial configuration of the assay involves per-forming the reaction under an abbreviated matrix of reaction times and
temperatures in order to identify the optimal conditions for generation of a
pattern with a broad spectrum of evenly distributed bands (temperature/time
optimization). Alternatively, recent studies have demonstrated that the use of a
programmable thermal cycler enables informative patterns to be generated by
increasing the reaction temperature throughout the course of the reaction, spe-cifically from 25 to 85°C at a rate of 0.1°C/s for a total ramping time of 10 min.
In some genetic systems, such as P. carinii, the ramping approach appears to
generate somewhat more even distributions of fragments and has improved
mutation detection sensitivity. Furthermore, provided suitable thermal cyclers
are available, the ramping approach is simpler and requires less DNA, since
optimization reactions need not be run prior to analysis of test samples. The
following protocol describes the method of performing CFLP analysis utiliz-ing either the single temperature or ramping procedure:
1. Aliquot the desired amount of end-labeled DNA (approx 100–200 fmol) into a
thin-walled reaction tube (200 or 500 µL, depending on the capacity of the ther-mal cycler). Bring the DNA to a final volume of 13 µL with DNA dilution buffer,
if necessary.
2. In a separate tube, prepare an enzyme master mix that contains the following for
each reaction: 2 µL of 10X CFLP buffer, 2 µL 2 mM MnCl2, 1 µL of Cleavase I
enzyme, 2 µL of 10 mM MgCl2 (optional, see Note 4), and DNA dilution buffer
to a final volume of 7 µL (if needed).
3. To denature samples, place tubes containing DNA in a programmable thermal cycler
(or heat block) and incubate at 95°C for 15 s. If the single temperature method is used
proceed to step 4. If the ramping method is to be used proceed to step 5.
4. Temperature/time optimization: After the 15-s denaturation step, set the thermal
cycler to the desired reaction temperature (or place the tubes in a heat block held
at reaction temperature if no thermal cycler is available). Optimal times and tem-peratures can be determined by examining matrices of different reaction times
(e.g., 1, 3, and 5 min) and temperatures (40, 50, and 55°C). Choose the con-ditions that yield the richest and most even pattern (see Note 5). Incubate the
CFLP reactions for the amount of time determined to be optimal, holding the
temperature constant. After the incubation period, stop the reactions with 16 µL
of stop solution. Proceed to Subheading 3.3.
38 Heisler and Lee
5. Ramping: After the 15-s denaturation step, set the thermal cycler to 35°C. As
soon as the thermal cycler reaches 35°C, add 7 µL of the enzyme/buffer mixture.
Mix well by pipetting up and down several times. Incubate the CFLP reactions
for 15 s at 35°C. Program the thermal cycler to increase in temperature at a rate of
0.1°C/s to 85°C, or set to ramp for an 8-min period from 35 to 85°C. On reaching
85°C, stop the reactions with 16 µL stop solution (see Note 3).
3.3. Separation of CFLP Fragments
1. Prepare a denaturing polyacrylamide gel, choosing a percentage of acrylamide
(19 1) appropriate for the size of the fragment being analyzed (see Note 6).
2. Prerun the gel for approx 30 min before loading the samples at sufficient wattage
to warm the gel (e.g., 18–20 W).
3. Heat denature the CFLP reactions at 80°C for 2 min immediately prior to loading
onto the gel. The best resolution is achieved when the samples are fully denatured.
4. Load 5–10 µL of the appropriate CFLP reaction per well. The remainder of the
reactions can be stored at 4°C or –20°C for later analysis.
5. Continue electrophoresis until sufficient separation of the CFLP fragments is
achieved (the time will depend on the fragment size and the percentage of
acrylamide used).
3.4. Visualization of CFLP Patterns
3.4.1. Fluorescence Imaging of CFLP Patterns
1. Following gel electrophoresis, thoroughly wash the outside of the gel plates
using nonfluorescent soap.
2. Dry and wipe clear with lens paper to remove residual debris.
3. Place the gel carefully in the fluorescence scanning unit (Hitachi FMBIO-100 or
Molecular Dynamics 595).
4. Scan using the correct wavelength or filter for the fluorescent group to be detected.
3.4.2. Chemiluminescence Detection of CFLP Patterns
1. After electrophoresis, wearing powder-free latex gloves that have been washed
with isopropanol (see Note 7), carefully separate the glass plates to expose the
acrylamide gel.
2. Cut a piece of Nytran Plus membrane (Schleicher and Schuell) to fit the gel size
and moisten by applying 5–10 mL of 0.5X TBE.
3. Carefully place the moistened membrane onto the gel, avoiding lifting and repo-sitioning the membrane, and smooth out air bubbles with a clean pipet. Transfer
starts immediately, so the membrane should not be picked up and repositioned
once it has come into contact with the gel.
4. Cover the membrane with two pieces of precut blotting paper, cover with a glass
plate, and place a binder clip on each side of the sandwiched gel. Alternatively,
for large gels (i.e., 20 × 20 cm or larger), place an approx 2-kg weight on top of
the sandwich.
5. Allow the DNA to transfer onto the membrane for 4–16 h (e.g., overnight, if
convenient) at room temperature.
CFLP Analysis for Genotyping and Mutation Detection 39
6. After the transfer, disassemble the sandwiched gel and remove the membrane by
carefully moistening it with distilled water. Mark the DNA side (i.e., the side
touching the gel during transfer) using a permanent laboratory marker.
7. Rinse a dish thoroughly with isopropanol (see Note 7) and fill with 0.2 mL/cm2
of 1X blocking buffer (e.g., 100 mL for a 20 × 20 cm membrane).
8. Transfer the membrane to the dish containing the blocking buffer and allow to
rock gently for 15 min.
9. Repeat the 15-min wash with fresh blocking buffer and discard the buffer.
10. Add 2 µL of Streptavidin-Alkaline-Phosphatase Conjugate to 50 mL of fresh block-ing buffer (or add conjugate to the blocking buffer at a volume ratio of 1 4000).
11. Pour the conjugate/blocking buffer mixture onto the blocked membrane and rock
gently for 15 min.
12. Remove the conjugate and rinse for 5 min with 0.1% SDS/1X SAAP buffer,
0.5 mL/cm2 each (200 mL for a 20 × 20 cm membrane). Repeat twice, for a total
of three washes.
13. Remove the SDS and rinse 5 min with 0.5 mL/cm2 1 mM MgCl2/1X SAAP buffer.
Repeat twice, for a total of three washes.
14. Place the membrane in a sealable bag and add 4 mL of CDP-Star (or
0.01 mL/cm2).
15. Seal the bag and spread the CDP-Star gently over the membrane for 3–5 min.
16. Completely remove the CDP-Star and any air bubbles. Transfer the membrane
while still in the bag to a film exposure cassette.
17. In the darkroom, expose the membrane to X-ray film. Initially expose for 30 min.
For subsequent exposures, adjust the time for clarity and intensity (see Note 8).
18. Develop the film.
4. Notes
1. Depending on the objective of the analysis in question, the DNA can be labeled
on either strand or on both strands using this approach. Single end labeling, i.e.,
of the sense or the antisense strand, permits some degree of localization of the
base change(s) corresponding to the observed pattern changes (3) . The sensitiv-ity of the CFLP method is approx 90% for single-stranded analysis and >95% for
two-strand analysis. While double end labeling precludes this localization, it
affords more sensitive mutational analysis.
2. It has been determined empirically that samples analyzed according to the ramp-ing procedure require approximately twofold more DNA than do those analyzed
by the conventional method. This is likely because in the ramping procedure,
digestion occurs throughout the course of the temperature increase and optimally
cleaves different hairpins at different temperatures.
3. The choice of dyes used in the stop solution depends on the system used to visu-alize the CFLP patterns. If chemiluminescence detection is used, then the stop
solution should include 0.05% bromophenol blue and 0.05% xylene cyanol (Sub-heading 2.2., item 6). If fluorescent scanning is used, then a dye that migrates
40 Heisler and Lee
with opposite polarity, such as crystal violet (0.05%), is preferable, because dyes
that migrate into the gel fluoresce at the wavelengths used to detect the fluores-cent dyes, thereby obscuring a portion of the CFLP pattern. Note that when a dye
with opposite polarity is used, it is advisable to load 3–5  µL of stop solution
containing bromophenol blue and xylene cyanol in a lane that does not contain
CFLP reactions in order to monitor the progress of gel electrophoresis.
4. MgCl2 dramatically reduces the rate of cleavage in the CFLP reaction. When MgCl2
is added to a final concentration of 1 mM in the presence of standard
MnCl2 concentrations of 0.2 mM, the rate of cleavage is slowed by as much
as 10-fold. This reduced reaction rate can be useful for analysis of DNA frag-ments that assume highly favored secondary structures that are rapidly
cleaved in the CFLP reaction. The presence of such structures is readily iden-tified by the appearance of a structural fingerprint comprising one or two
prominent bands. When 1 mM MgCl2 is added, the optimal time and tempera-ture of digestion should be reevaluated to reflect the reduced rate of cleavage
(see Note 5).
5. The structural fingerprint produced by CFLP digestion is a collection of frag-ments resulting from partial digestion, usually of 5v end labeled fragments.
Because the CFLP reaction is a partial digestion and because the formation of the
substrate secondary structures depends on reaction temperature, it is possible to
modulate the extent of digestion through variations in the duration and tempera-ture of the reaction. Specifically, lower temperatures stabilize secondary struc-ture formation whereas higher temperatures reduce the number of structures
formed by a given molecule. Similarly, longer reaction times lead to increased
accumulation of smaller cleavage products. The most informative fingerprints
are those that contain a relatively even distribution of low and high molecular
weight products, including a fraction of full-length, uncut DNA. Ensuring that
the entire size distribution of cleavage products is visible increases the likelihood
of detecting the products that reflect the presence of a polymorphism.
6. The percentage of polyacrylimide to be used is dictated by the size of the PCR
fragment being analyzed. Appropriate percentages of polyacrylamide for various
size ranges are well established (19) .
7. The objective of this step is to minimize carryover of alkaline phosphatase from
previous reactions and from exogenous sources (e.g., skin). Throughout this pro-cedure, it is of paramount importance to minimize contamination by this ubiqui-tous enzyme.
8. An alkaline phosphatase reaction with the chemiluminescence substrate produces
a long-lived signal, especially on membranes. Light emission increases of >300-fold are seen in the first 2 h on application of the substrate onto nylon mem-branes, with the chemiluminescence signal persisting up to several days. Because
film exposure times range from minutes to several hours, multiple images may
be acquired. Varying film exposure times enables the user to optimize signal to
noise.
CFLP Analysis for Genotyping and Mutation Detection 41
Acknowledgments
We wish to acknowledge the efforts of Mary Oldenburg, Senior Technical
Scientist, Third Wave Technologies, in performing much of the CFLP reaction
optimization as well as in providing critical commentary on the manuscript.
References
1. Sreevatsan, S., Bookout, J. B., Ringpis, F. M., Pottathil, M. R., Marshall, D. J.,
de Arruda, M., Murvine, C., Fors, L., Pottathil, R. M., and Barathur, R. R.
(1998) Algorithmic approach to high-throughput molecular screening for alpha
interferon-resistant genotypes in hepatitis C patients.  J. Clin. Microbiol. 36,
1895–1901.
2. Cotton, R. G. H. (1997) Slowly but surely towards better scanning for mutations.
Trends Genet. 13, 43–46.
3. Brow, M. A., Oldenberg, M., Lyamichev, V., Heisler, L., Lyamicheva, N., Hall,
J., Eagan, N., Olive, D. M., Smith, L., Fors, L., and Dahlberg, J. (1996) Differen-tiation of bacterial 16S rRNA genes and intergenic regions and Mycobacterium
tuberculosis katG genes by structure-specific endonuclease cleavage.  J. Clin.
Microbiol. 34, 3129–3137.
4. Orita, M., Suzuki, Y., Sekiya, T., and Hayashi, K. (1989) Rapid and sensitive
detection of point mutations and DNA polymorphisms using the polymerase chain
reaction. Genomics 5, 874–879.
5. Hayashi, K. (1991) PCR-SSCP: a simple and sensitive method for detection of
mutations in the genomic DNA. PCR Methods Applica. 1, 34–38.
6. Sarkar, G., Yoon, H., and Sommer, S. S. (1992) Dideoxy fingerprinting (ddE):
a rapid and efficient screen for the presence of mutations. Genomics 13, 441–443.
7. Sreevatsan, S., Bookout, J. B., Ringpis, F. M., Mogazeh, S. L., Kreiswirth, B. N.,
Pottathil, R. R., and Barathur, R. R. (1998) Comparative evaluation of cleavase
fragment length polymorphism with PCR-SSCP and PCR-RFLP to detect antimi-crobial agent resistance in Mycobacterium tuberculosis. Mol. Diagn. 3, 81–91.
8. Tahar, R. and Basco, L. K. (1997) Analysis of Plasmodium falciparum multidrug-resistance (pfmdr1) gene mutation by hairpin-dependent cleavage fragment length
polymorphism. Mol. Biochem. Parasitol. 88, 243–247.
9. Wartiovaara, K., Hytonen, M., Vuori, M., Paulin, L., Rinne, J., and Sariola, H.
(1998) Mutation analysis of the glial cell line-derived neurotrophic factor gene in
Parkinson’s disease. Exp. Neurol. 152, 307–309.
10. Schlamp, C., Poulsen, G. L., Nork, M., and Nickells, R. W. (1997) Nuclear exclu-sion of wild-type p53 in immortalized human retinoblastoma cells. J. Natl. Can-cer Inst. 89, 1530–1536.
11. Marshall, D. J., Heisler, L. M., Lyamichev, V., Murvine, C., Olive, D. M., Ehrlich,
G. D., Neri, B. P., and de Arruda, M. (1997) Determination of hepatitis C virus
genotypes in the United States by Cleavase Fragment Length Polymorphism
analysis. J. Clin. Microbiol. 35, 3156–3162.
42 Heisler and Lee
12. Rainaldi, G., Marchetti, S., Capecchi, B., Meneveri, R., Piras, A., and Leuzzi, R.
(1998) Absence of mutations in the highest mutability region of the p53 gene in
tumour-derived CHEF18 Chinese hamster cells. Mutagenesis 13, 153–155.
13. Rossetti, S., Englisch, S., Bresin, E., Pignatti, P. F., and Turco, A. E. (1997)
Detection of mutations in human genes by a new rapid method: cleavage frag-ment length polymorphism analysis (CFLPA). Mol. Cell. Probes 11, 155–160.
14. Eisinger, F., Jacquemier, J., Charpin, C., Stoppa-Lyonnet, D., Bressac-de Paillerets, B., Peyrat, J.-P., Longy, M., Guinebretiere, J.-M., Sauvan, R.,
Noguichi, T., Birnbaum, D., and Sobol, H. (1998) Mutations at BRCA1: the med-ullary breast carcinoma revisited. Cancer Res. 58, 1588–1592.
15. Centers for Disease Control and Prevention (1989) AIDS Weekly Surveillance
Report, Centers for Disease Control and Prevention, Atlanta, GA.
16. Latouche, S., Ortona, E., Masers, E., Margutti, P., Tamburrini, E., Siracusano, A.,
Guyot, K., Nigou, M., and Roux, P. (1997) Biodiversity of Pneumocystis carinii
hominis: typing with different DNA regions. J. Clin. Microbiol. 35, 383–387.
17. Lu, J.-J., Bartlett, M. S., Shaw, M. M., Queener, S. F., Smith, J. W., Ortiz-Rivera,
M., Leibowitz, M. J., and Lee, C.-H. (1994) Typing of Pneumocystis carinii strains
that infect humans based on nucleotide sequence variations of internal transcribed
spacers of rRNA genes. J. Clin. Microbiol. 32, 2904–2912.
18. Lee, C.-H., Tang, X., Jin, S., Li, B., Bartlett, M. S., Helweg-Larsen, J., Olsson,
M., Lucas, S. B., Roux, P., Cargnel, A., Atzori, C., Matos, O., and Smith, J. W.
(1998) Update on Pneumocystis carinii f. sp. hominis typing based on nucleotide
sequence variations in internal transcribed spacer regions of rRNA genes. J. Clin.
Microbiol. 36, 734–741.
19. Sambrook, J., Fritsch, E. F., and Maniatis, T. (eds.) (1989) Molecular Cloning:
A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring
Harbor, NY.
Detection of Telomerase by In Situ Hybridization and TRAP 43
6
Detection of Telomerase by In Situ
Hybridization and by Polymerase Chain
Reaction-Based Telomerase Activity Assay
Carmela P. Morales and Shawn E. Holt
1. Introduction
The onset of human cancer typically requires numerous genetic mutations,
generally specific for the tissue type from which the cancer originates. Thus, it
has been difficult to screen all tumor types for a single mutation. In recent
years, telomerase activity has been associated with at least 85% of human
malignancies as well as with some lesions considered preneoplastic by tradi-tional cytology (1 ,2) . Telomerase appears to be ubiquitously associated with
a wide array of human cancers from a variety of tissue sources. Therefore,
detection of telomerase activity relative to human cancer development is likely
to be an important and novel method, in combination with cytology, for cancer
diagnosis.
Telomerase is a ribonucleoprotein, minimally composed of both an RNA
component (hTR) and a catalytic protein component (hTERT) (3) . Most nor-mal somatic cells and tissues express hTR at low levels but not hTERT, and are
devoid of telomerase activity. Without a mechanism to ensure telomeric integ-rity and stability, telomeric repeats are lost with successive cellular divisions
owing to the inability of conventional DNA polymerases to replicate the ends
of linear chromosomes (4,5) . This loss of telomeric DNA signals a DNA dam-age response mechanism, requiring the tumor suppressor molecules p53 and
pRB, which triggers an irreversible growth-arrested phenotype known as cellu-lar senescence (6–8) . Those cells that are capable of bypassing senescence by
blocking the normal functions of p53 and pRB continue telomere shortening
until the cells reach a period of crisis (9–11) . During crisis, cells either die or in
43
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
44 Morales and Holt
rare instances, immortalize, and this immortalization event is almost always
concomitant with activation of telomerase activity (2) . Telomerase is neces-sary for the continued proliferation of immortal and tumor cells, as inhibition
results in reprogramming of the cellular senescence program (12) . Telomerase
activation requires a significant increase in the expression of the integral RNA
component (hTR) and the activation of the catalytic subunit (hTERT). Thus,
determination of telomerase activity in human cancer can be accomplished by
detecting either the upregulation of hTR or activity using a polymerase chain
reaction (PCR)-based assay. In this chapter, we describe both the  in situ
hybridization technique and the PCR-based telomeric repeat amplification
protocol (TRAP) assay.
1.1. In Situ Hybridization for RNA Component of Telomerase
The utility of  in situ hybridization for the RNA component of human
telomerase (hTR) for cancer diagnosis has been recently evaluated  (13 ,14) .
Whereas the detection of telomerase activity using the TRAP assay requires
fresh or fresh-frozen tumor biopsies, analysis of hTR can be accomplished
using archival, formalin-fixed, paraffin-embedded specimens, as well as fro-zen sections and cultured cells (15–17) . In lung cancer, an excellent correlation
between in situ hybridization for telomerase RNA and relative telomerase
activity levels has been demonstrated (18) . The excellent morphologic preser-vation of tissue with this technique makes it a useful alternative to the PCR-based TRAP assay.
1.2. Telomerase Activity Detection Using the TRAP Assay
The TRAP assay is a highly sensitive, PCR-based method for accurately
determining levels of telomerase activity in a sample from fresh or frozen tis-sue or from cultured cells (Fig. 1) (2 ,19 ,20) . This technique has been used to
detect telomerase activity in a wide variety of human tumors from varied tissue
sources (for a review, see ref. 1 ).
2. Materials
2.1. Telomerase Detection Using In Situ hTR Assay
2.1.1. General Reagents
1. Distilled, deionized autoclaved water.
2. Diethylpyrocarbonate (DEPC)-treated water: distilled, deionized water treated
with 0.1% DEPC.
3. 10X Phosphate-buffered saline (PBS).
4. 100% Ethanol, room temperature and –20°C.
5. 1 M Tris-HCl, pH 7.8.
Detection of Telomerase by In Situ Hybridization and TRAP 45
6. 0.5 M EDTA, pH 8.0.
7. 10 N Sodium hydroxide.
8. Phenol chloroform (1 1).
9. 1 M Dithiothreitol (DTT).
10. 3 M Sodium acetate, pH 6.0.
11. 20X Saline sodium citrate (SSC): 3 M NaCl, 0.3 M sodium citrate.
12. 70% Ethanol.
Fig.1. Schematic representing the TRAP assay. Telomerase samples are processed
by mild detergent lysis. A nontelomeric primer that serves as the telomerase substrate
(TS primer) is combined with water, buffer, dNTPs,  Taq polymerase, the reverse
primer (RP/ACX), and two primers that serve to amplify an internal control (TSNT
and NT). After this cocktail is made, the tissue or cell extract is added and incubated at
room temperature for 30 min for telomerase to extend the TS primer. Telomerase-mediated extension occurs in six base increments, corresponding to the integral RNA
component of telomerase (hTR) and the telomeric repeat sequences (TTAGGG). PCR
amplification of the telomerase products and the internal standard utilizes both the TS
and RP/ACX primers. The internal control primer (TSNT) utilizes both the NT and the
TS primers, hence a competition for TS primer in the amplification of both telomerase
products and TSNT. The internal control measures 36 bp after amplification and serves
as a quantitative standard to normalize sample-to-sample variation.
46 Morales and Holt
2.1.2. Pretreatment of Paraffin-Embedded Sections
1. Fresh, filtered 4% paraformaldehyde, pH 7.5.  (Note: paraformaldehyde is
highly toxic and should be made fresh for each use in a fume hood.)
2. 0.85% NaCl.
3. Xylene.
4. Proteinase K (20 mg/mL).
5. TE buffer: 100 mM Tris-HCl, pH 7.5, 50 mM EDTA.
6. 1 M Triethanolamine.
7. 98% Acetic anhydride.
2.1.3. Pretreatment of Cultured Cells
1. Same as paraffin-embedded sections (except xylene).
2. Triton X-100.
3. 2 N HCl.
4. 1 M glycine.
2.1.4. Preparation of Antisense hTR DNA Template
1. EcoRV and NaeI restriction enzymes.
2. TE buffer (see Subheading 2.1.2.).
2.1.5. Preparation and Labeling of Probe
1. SP6 RNA polymerase (5 U/µL).
2. ATP, CTP, and GTP (10 mM ).
3. 35S-UTP (20 mCi/mL and 1000 Ci/mmol).
4. 10X SP6 RNA polymerase transcription buffer.
5. Hydrolysis solution: 0.01  M DTT, 0.08  M sodium bicarbonate, and 0.12  M
sodium carbonate.
6. Neutralization solution: 0.2  M sodium acetate, pH 6.0, 1% glacial acetic acid,
and 0.01 M DTT.
7. RNase inhibitor (5 U/µL).
8. DNase I (2 U/µL).
9. Quick Spin™ columns, G50 Sephadex columns for radiolabeled RNA (Boeh-ringer Mannheim).
10. Yeast tRNA (20 mg/mL).
2.1.6. In Situ Hybridization
1. Cocktail solution: 3 M NaCl, 0.2 M sodium acetate, pH 6.0, 50 mM EDTA, pH 8.0,
50% deionized formamide, 1X Denhardt’s, 10% dextran sulfate, 0.1 M DTT.
2. Clean glass cover slips.
2.1.7. Washing and Detection
1. Formamide solution: 2X SSC, 50% formamide, 10 mM DTT.
2. 5X SSC solution: 5X SSC, 10 mM DTT.
Detection of Telomerase by In Situ Hybridization and TRAP 47
3. 10X Wash solution: 0.4 M NaCl, 10 mM Tris-HCl, pH 7.8, 5 mM EDTA.
4. Pancreatic RNase A (10 mg/mL).
5. Kodak emulsion type NTB-2 (cat. no. 165 4433).
2.1.8. Developing of Sections
1. Kodak Dektol developer (cat. no. 146 4700).
2. Kodak fixer (cat. no. 197 1720).
3. Hematoxylin.
4. Permount™ mounting medium.
2.1.9. Equipment
1. Glass slide chambers, 250-mL capacity.
2. Scintillation counter.
3. Tissue-Tek™ slide staining set (VWR Scientific).
4. Dip Miser (Electron Microscopy Services).
2.2. Telomerase Activity Using TRAP Assay
2.2.1. General Items and Reagents
1. DEPC-treated water: distilled, deionized water treated with 0.1% DEPC.
2. TRAP-eze™ Telomerase Detection Kit (Intergen).
3. Aerosol-resistant tips (USA/Scientific).
4. Beta-shield for protection from radioisotope.
5. Acrylic tube racks (24 place; USA/Scientific).
2.2.2. Preparation of Cell Extracts
from Cultured Cells and Clinical Material
1. 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS)
lysis buffer: 10 mM Tris-HCl, pH 7.5, 1 mM EGTA, 1 mM MgCl2, 5 mM
`-mercaptoethanol, 0.5% CHAPS, 10% glycerol, 1 mM 4-(2-aminoethyl)-benzenesulfonyl fluoride hydrochloride (AEBSF), 0.5 U/µL of RNase inhibitor
(Boehringer Mannheim).
2. NP-40 Lysis buffer: 10 mM Tris-HCl, pH 7.5, 1 mM EGTA, 1 mM MgCl2, 5 mM
`-mercaptoethanol, 1% NP-40 (renamed IPGAL), 0.25 mM sodium deoxycho-late, 150 mM NaCl, 10% glycerol, 1 mM AEBSF.
3. 1.5-mL Microfuge tubes, RNase/DNase free.
4. Microfuge for speeds to 6000g (Eppendorf).
5. Refrigerated centrifuge for speeds to 14,000g (Eppendorf).
6. Kontes homogenization tubes with matching pestles (VWR Scientific).
7. Rechargeable, low-speed drill (CD1000; Black and Decker).
8. BCA assay for protein concentration (Pierce).
9. Liquid nitrogen.
10. Ice buckets.
48 Morales and Holt
2.2.3. Primer Labeling
1. TS primer (high-performance liquid chromatography [HPLC] purified), sequence
5v-AAT CCG TCG AGC AGA GTT-3v.
2. Redivue™ [a-32P]ATP (6000 Ci/mmol) (Amersham).
3. T4 polynucleotide kinase (10 U/µL) (Life Technologies).
4. 5X Kinase buffer (Life Technologies).
2.2.4. Telomerase Reactions Using TRAP Assay
1. Reverse primer (RP or ACX) (HPLC purified), sequence 5v-GCG CGG CTT ACC
CTT ACC CTT ACC CTA ACC-3v.
2. 36-bp Internal standard primers (HPLC purified): 36-bp template sequence
(TSNT) 5v-AAT CCG TCG AGC AGA GTT AAA AGG CCG AGA AGC GAT-3v;
return primer (NT) sequence 5v-ATC GCT TCT CGG CCT TTT-3v.
3. 10X TRAP reaction buffer: 200 mM Tris-HCl, pH 8.3, 10 mM EGTA, 15 mM
MgCl2, 630 mM KCl, 0.5% Tween-20.
4. 50X dNTPs: 2.5 mM dATP, TTP, dGTP, dCTP each.
5. Labeled TS primer.
6. Taq DNA polymerase (5 U/µL) (Life Technologies).
7. 0.5-mL Microfuge tubes (USA/Scientific).
8. MJ Research PCR machine with heated lid (PTC-100).
2.2.5. Gel Electrophoresis and Detection
1. TRAP loading buffer: 50 mM EDTA, 50% glycerol, 0.25% bromophenol blue,
0.25% xylene cyanol.
2. 40% Acrylamide (19 1, acrylamide bisacrylamide ratio) (Bio-Rad).
3. 10% Ammonium persulfate.
4. TEMED.
5. 10X TBE running buffer: 50 mM Tris-borate, 1 mM EDTA, pH 8.3.
6. Vertical Gel Electrophoresis System (Life Technologies).
7. Plastic disposable transfer pipet.
8. Power supply to 300 V with timer (power pac 3000; Bio-Rad).
9. Fix solution: 0.5 M NaCl, 50% ethanol, 40 mM sodium acetate, pH 4.2.
10. Plastic wrap.
11. Phosphorimage cassettes.
12. Phosphorimager (Molecular Dynamics).
3. Methods
3.1. In Situ Detection of Telomerase
The following procedure outlines the technique required for in situ hybrid-ization of telomerase RNA. Figure 2 shows the final hTR in situ hybridization
result for an esophageal adenocarcinoma, along with a hematoxylin & eosin
(H&E) stain. Because RNA probes are used, great care must be taken to main-
Detection of Telomerase by In Situ Hybridization and TRAP 49
tain probe integrity and avoid degradation by RNases. All solutions and dilu-tions should be prepared with DEPC-treated water, and all glassware should be
cleaned with an appropriate RNase-free detergent. Unless otherwise indicated,
all reactions are done at room temperature.
3.1.1. Pretreatment of Paraffin-Embedded Sections
1. Cut 4-µm sections from blocks and place on SuperFrost™ Plus glass slides (see
Note 1).
2. Bake slides for 20 min at 80°C.
3. Deparaffinize in xylene two times for 10 min each.
4. Rehydrate in a graded ethanol series: 100% ethanol twice for 2 min each fol-lowed by 95, 85, 70, 50, and 30% ethanol for 1 min each.
5. Rinse once for 5 min with 0.85% NaCl.
6. Rinse once for 5 min with PBS.
7. Fix tissue at room temperature for 20 min with 4% paraformaldehyde.
8. Wash two times for 5 min each with PBS.
9. Permeabilize sections for 35 min at 37°C with TE buffer containing 20 µg/mL of
proteinase K (see Note 2).
10. Wash once for 5 min with PBS.
11. Postfix in 4% paraformaldehyde for 5 min (reused from step 7).
Fig. 2. In situ hybridization for the RNA component of human telomerase in meta-static esophageal adenocarcinoma.  (A)  An H&E stain demonstrates malignant cells
adjacent to a lymphoid follicle and vascular structure, both uninvolved. (B) Cells are
considered to be positive for telomerase RNA if they display nuclear grains in the
overlying emulsion. Hybridization of a serial section with the antisense probe demon-strates an intense telomerase signal in the malignant cells whereas the endothelial cells
demonstrate no detectable telomerase RNA. Note that cells of the germinal center of
the lymphoid follicle are weakly positive for telomerase RNA.
50 Morales and Holt
12. Rinse in DEPC-treated water for 30 s.
13. Acetylate sections for 10 min on a rocking platform in 0.1  M triethanolamine
containing 0.25% acetic anhydride (see Note 3).
14. Wash once for 5 min with PBS.
15. Wash once for 5 min with 0.85% NaCl.
16. Dehydrate the slides in a graded ethanol series: 30, 50, 70, 85, 95, and 100% for
1 min each.
17. Air-dry slides for 30 min. The slides may be kept in an airtight container at room
temperature for up to 1 wk prior to hybridization.
3.1.2. Pretreatment of Cultured Cells
1. Grow cultured cells on sterile glass Chamber slides overnight at 37°C.
2. Wash two times for 5 min each with PBS.
3. Fix according to either of these procedures: 10 min in cold acetone and air-dried;
or 10 min in cold 95% isopropanol and air-dried. Proceed directly to step 4.
4. Incubate for 10 min in 4% paraformaldehyde at room temperature.
5. Wash two times for 5 min each with PBS.
6. Permeabilize for 3 min with PBS containing 0.05% Triton X-100.
7. Wash two times for 5 min each with PBS.
8. Deproteinize for 10 min with 0.2 N HCl.
9. Wash two times for 5 min each with PBS.
10. Permeabilize sections for 5–15 min at 37°C with TE buffer containing 1–10 µg/mL
of proteinase K. We regularly use 5 µg/mL for 5 min.
11. Wash two times for 5 min each in PBS containing 2 mg/mL of glycine (to inacti-vate the enzyme).
12. Postfix for 5 min in 4% paraformaldehyde.
13. Wash two times for 5 min each with PBS.
14. Acetylate sections for 10 min on a rocking platform in 0.1  M triethanolamine
containing 0.25% acetic anhydride (see note 3).
15. Dehydrate the slides in a graded ethanol series: 30, 50, 70, 85, 95, and 100% for
1 min each.
16. Air-dry slides for 30 min. The slides may be kept in an airtight container at room
temperature for up to 1 wk prior to hybridization.
3.1.3. Preparation of Antisense hTR DNA Template
hTR is the human telomerase template RNA sequence utilized by telomerase
to add TTAGGG repeats to the 3vends of chromosomes (21) . The 560 nucle-otide cDNA sequence is cloned into the SacI site of the pGEM-5Zf(+) expres-sion vector (Boehringer Mannheim). After linearization with either EcoRV or
NaeI, runoff in vitro transcription of the hTR expression plasmid with either
SP6 or T7 yields the antisense (SP6) or sense (T7) RNA probe, respectively.
1. In a standard digestion at 37°C, linearize hTR expression plasmid with EcoRV.
If desired, set up a parallel digestion with NaeI.
Detection of Telomerase by In Situ Hybridization and TRAP 51
2. Purify the linearized template by standard phenol-chloroform extraction and etha-nol precipitation (see Note 4).
3. Resuspend the pellet at approx 1 µg/mL in TE buffer, and quantitate by running
a 200-ng equivalent on a 0.8% agarose gel with an appropriate DNA mass ladder.
3.1.4. Labeling of Antisense RNA Probe
1. Set up an in vitro transcription of the hTR template as follows: 6 µL of DEPC-treated water; 1 µL of RNase inhibitor (5 U/µL); 2 µL of 10X transcription buffer;
1 µL of ATP, CTP, and GTP (all 10 mM); 5 µL 35S-UTP; 1 µL of hTR template
(1 µg/µL); and 2 µL of SP6 RNA polymerase (5 U/µL).
2. Add 1 µL of DNase (1 U/µL) for 15 min at 37°C.
3. Perform alkaline hydrolysis at 60°C by adding 50 µL of hydrolysis solution to
the transcription reaction for 15 min.
4. Neutralize on ice by adding 50 µL of neutralizing solution.
5. While the tube is on ice, prepare Quick Spin columns according to the
manufacturer’s instructions.
6. Keeping the columns upright, carefully apply the entire RNA sample to the cen-ter of the column bed.
7. Centrifuge for 4 min at 1100g.
8. Save the eluate and discard the column into a radioactive waste container.
9. Add 3 µL of yeast tRNA (20 mg/mL) to the eluate and mix well.
10. Add 1/10 volume of 3  M sodium acetate (pH 6.0) and 2 vol of chilled 100%
ethanol and precipitate for 20 min at –80°C.
11. Centrifuge for 25 min at 12,800g.
12. Carefully draw off the supernatant and discard.
13. Gently add 2 vol of chilled 70% ethanol.
14. Centrifuge for 20 min at 14,000g.
15. Draw off the supernatant and allow the pellet to air-dry.
16. Resuspend the pellet in 30 µL of 100 µM DTT; count 1 µL of the final solution.
Using this protocol, we usually obtain counts of approx 2 to 3 × 106 cpm/µL.
3.1.5. In Situ Hybridization
1. Combine probe mix and cocktail solution, vortex, and heat to 90°C for 5 min.
The final hybridization solution contains probe mix and cocktail solution in a
0.14 0.86 ratio. Once the required volume of final hybridization solution is
determined based on the number of slides, the amount of probe mix can be calcu-lated. Probe mix is made of 50% yeast tRNA plus probe and DEPC-treated water
for the remaining 50%. The probe mix is then added to the appropriate volume of
cocktail solution. We utilize a final probe concentration of 50,000 cpm/µL and
typically apply 50  µL of hybridization solution to each slide. For example, to
hybridize 10 slides, 430  µL of hybridization cocktail and 70 µL of probe mix
would be required. If the probe count were 2 × 106/µL, then the probe mix would
be prepared with 12.5 µL of probe, 35 µL of yeast tRNA, and 22.5 µL of DEPC-treated water.
52 Morales and Holt
2. Quickly place hybridization solution on ice.
3. Apply 50  µL of hybridization solution over each section and cover slip. Take
care not to trap air bubbles underneath the cover slip.
4. Place the slides horizontally into a clean, upright slide container (25-slide capacity).
5. Place several Kimwipes moistened with DEPC-treated water at the bottom of
each box, and wrap the boxes in clear plastic wrap followed by aluminum foil.
6. Keeping the box upright, hybridize slides overnight at 50°C.
3.1.6. Washing and Detection
1. In glass staining chambers, prewarm 250 mL each of the 5X SSC solution to
50°C and the formamide wash solution to 65°C.
2. After hybridization, unwrap the box and carefully remove the cover slip from
each slide.
3. Wash the slides at 50°C for 30 min in 5X SSC solution with agitation.
4. Transfer the slides to the formamide solution at 65°C for 30 min with agitation.
5. Wash two times for 10 min each at 37°C with 1X wash solution.
6. Incubate the slides for 30 min at 37°C in 250 mL of 1X wash solution containing
500 µL of RNase A.
7. Rinse once for 5 min with 1X wash solution at 37°C.
8. Wash in 2X SSC for 1 h at 37°C.
9. Wash in 0.1X SSC for 1 h at 37°C.
10. Dehydrate for 1 min each in a graded ethanol series: 30, 50, 70, 85, 95, and
100%.
11. Air-dry the slides for 30 min and proceed with emulsion.
12. Warm the emulsion in a 45°C water bath with occasional gentle swirling.
13. In a dark room, pour a small amount of emulsion in a Dip Miser or other suitable
container.
14. Gently dip a blank slide in the emulsion and inspect for bubbles. If bubbles are
present, allow them to clear for 10 min.
15. Dip the slides into the emulsion in a consistent fashion to minimize slide-to-slide
variability (we perform two 1-s dips in rapid succession).
16. Place the slides vertically in a 50-mL conical styrofoam container and air-dry in
the dark for 1 h.
17. Place the dried slides in a slide box containing desiccant wrapped in a Kimwipe,
and wrap the box in two layers of foil.
18. Place the box upright at –80°C for 2–3 wk (see note 5).
3.1.7. Developing Sections
1. Prepare developer and fixer per manufacturer’s directions and cool to 15°C.
2. Bring the slides to room temperature for several hours.
3. In the dark, load the slides onto a clean slide holder, while trying not to touch the
emulsion.
4. Soak in full-strength developer for 3 min without agitation.
5. Rinse in distilled water for 30 s with mild agitation.
Detection of Telomerase by In Situ Hybridization and TRAP 53
6. Fix the slides two times for 3 min each.
7. Place the slides in distilled water cooled to 15°C (see Note 6).
8. Rinse the slides twice in distilled water.
9. Using a single-edge razor blade, remove the emulsion from the back of each
slide.
10. Counterstain with H&E for 2 min.
11. Rinse in running tap water for 1 min.
12. Dehydrate the slides in a graded ethanol series, clear with xylene, and cover slip.
3.2. TRAP Assay
Because the TRAP assay is PCR based, it is important to take every precau-tion to preserve sample integrity, prevent cross contamination, and avoid slop-piness. Separate work areas and separate pipets for each step of the assay should
be used to lessen the likelihood of contamination. Gloves must be worn at all
times during the assay and a surgical mask is highly recommended. Reagents
should be made with DEPC-treated water, and all equipment should be cleaned
with 70% ethanol prior to use (ice buckets, centrifuges, tube racks, and so on).
All reagents should be used only for the TRAP assay, aliquoted in small
amounts, and kept separate from general laboratory stocks. The TRAP assay is
highly sensitive, which could lead to false positive results if the necessary pre-cautions are not taken. It is absolutely required that each of the following steps
be done using aerosol-resistant tips to avoid possible contamination from
sample to sample.
3.2.1. Preparation of Cell Extracts
from Cultured Cells and Clinical Material
3.2.1.1. CULTURED CELLS
1. Harvest and count cells.
2. Put an aliquot corresponding to 100,000 cells into a sterile 1.5-mL microfuge
tube.
3. Pellet cells at 6000g for 5 min and remove the supernatant.
4. Either lyse samples immediately or store at –80°C until lysis.
5. Add 100–200  µL of ice-cold NP-40 lysis buffer and mechanically lyse the
samples (by pipeting the pellet up and down a few times, avoiding bubbles). The
lysis buffer is normally made with all the ingredients except for AEBSF and
stored at 4°C until use. AEBSF is added fresh with each round of lysis.
6. Save a small amount of lysis buffer for use as a negative control in the TRAP
reactions.
7. Incubate samples on ice for 20–30 min.
8. Centrifuge lysates at 14,000g for 20 min at 4°C.
9. Remove 80% of the lysate (do not disturb pellet) and flash-freeze in liquid nitrogen.
10. Store the samples at –80°C until use in the TRAP assay.
54 Morales and Holt
3.2.1.2. CLINICAL MATERIAL
1. Mechanically disperse partially thawed tissue samples on disposable Petri dishes
using sterile, disposable scalpel blades.
2. Immediately transfer tissue shavings to a sterile 1.5-mL Kontes tube.
3. Add 200 µL of ice-cold CHAPS lysis (4°C) buffer to the tumor samples. Again,
AEBSF is added fresh.
4. Save a small amount of lysis buffer to be used as a negative control in the TRAP
reactions.
5. Homogenize samples until the tissue is well dispersed using disposable pestles
(matching the Kontes tubes) and a drill, with rotation no faster than 450 rpm to
preserve enzyme integrity.
6. Centrifuge lysates at 14,000g for 20 min at 4°C.
7. Remove 80% of the lysate (do not disturb the pellet).
8. Take an aliquot for assessment of protein concentration using the Pierce BCA
assay.
9. Dilute lysates to 3 µg of protein/µL.
10. Flash-freeze the samples and store at –80°C.
3.2.2. Primer Labeling
Kinase reactions can be done at any time prior to making the TRAP master mix
(see Subheading 3.2.3., step 1); the amount of primer used is 100 ng/reaction.
Reactions consist of TS primer (100 ng/sample), 5X kinase buffer, DEPC water,
2.5 µCi of [a-32P]ATP/sample, and 0.25 U of T4 polynucleotide kinase/sample.
1. Incubate at 37°C for 20 min followed by heat inactivation of the kinase at 85°C
for 5 min. This is generally done in a thermocycler.
2. Prior to addition to the TRAP master mix, cool the sample on ice for at least 5
min. If it is not to be used immediately, labeled primer may be stored at –20°C
for up to 1 wk.
3.2.3. Telomerase Reactions Using TRAP Assay
A general schematic of the TRAP reaction is shown in Fig. 1.
1. Make a master mix that includes all the components necessary for TRAP, in a
tube that is RNase/DNase free and able to hold the appropriate volume (see Notes
7 and 8).
2. Use 48-µL reaction volumes (to which 2 µL of sample will be added) in sterile
0.5-mL tubes. Each reaction consists of 38.6 µL of DEPC-treated water, 5 µL of
10X TRAP reaction buffer, 1 µL of 50X dNTPs, 100 ng of TS primer (generally
2-µL vol of end-labeled primer), 100 ng of RP/ACX primer, 100 ng of NT primer,
0.01 amol of TSNT internal control template, and 0.4  µL of  Taq polymerase
(2 U). Ten to 20 samples are normally done per TRAP setup, and combining all
reagents into a TRAP master mix and aliquoting to each individual tube is rec-ommended. Because of frequent pipetting error, master mix volumes should be
the number of samples plus two.
Detection of Telomerase by In Situ Hybridization and TRAP 55
3. Add 2  µL of sample (corresponding to 100–1000-cell equivalents for cultured
cells or 0.1–6 µg of protein for clinical material) to each tube and write down the
tube number and what the sample is.
4. Mix by flicking the tube. Do not vortex or overmix, and make certain that the
entire sample remains in the bottom of the tube.
5. As a negative control, use 2 µL of lysis buffer used for making the sample lysates. In
addition, use as a positive control a telomerase-positive cell line (typically 1000-and 100-cell equivalents of 293 or H1299 cells).
6. Incubate TRAP reactions at room temperature (22–25°C) for no more than
30 min.
7. Preheat the PCR machine with a heated lid to 94°C and insert the samples (i.e., a
“hot start” reaction is used).
8. Proceed with the PCR at an initial hot start at 94°C for 3 min, followed by two-step PCR (94°C for 30 s, 60°C for 30 s) for 27–32 cycles. Twenty-seven or 28
cycles are sufficient for cells in culture; 30–32 cycles may be necessary for
tumors. PCR takes about 75 min from start to finish.
9. Use a separate rack for storage of the samples after PCR than was used for the
TRAP setup, because cross contamination of PCR products can occur.
10. Add 5 µL of TRAP loading buffer to each sample.
11. Load the samples directly onto the gel or store at 4°C or –20°C for up to 5 d with-out a decline in signal intensity.
3.2.4. Gel Electrophoresis and Detection
1. During the PCR, pour a 10% polyacrylamide gel (normally 20 wells) for electro-phoresis of samples. Gel mixtures consist of 2.5 mL of 10X TBE, 12.5 mL of
40% acrylamide, 35 mL of deionized water, 130  µL of 10% ammonium
persulfate, and 45 µL of TEMED.
2. Allow the poured gel to polymerize for at least 1 h.
3. After polymerization, remove 20-well combs from the gel and rinse the wells
with deionized water to remove any unpolymerized acrylamide.
4. Submerge the gel in 0.5X TBE running buffer and flush the wells out again with
running buffer and a plastic disposable transfer pipet.
5. Load 20–25  µL of each PCR sample onto the gel using aerosol-resistant gel-loading tips.
6. Run the gel at 300 V for 2 h. The bromophenol blue dye should migrate off of the
gel, and the xylene cyanol should be approximately halfway. The bottom portion
of the gel box contains buffer that will be highly radioactive and should be dis-posed of properly.
7. Remove the gel from the apparatus, remove the wells with a scalpel, and notch
the top left corner (corresponding to lane 1) for orientation purposes.
8. Fix the gel for 30 min in fix solution at room temperature.
9. After fixing, drain excess fix solution from the gel using paper towels.
10. Wrap the gel in plastic wrap and smooth out all the bubbles. There is no need to
dry the gel unless compelled to do so.
56 Morales and Holt
11. For detection of TRAP ladders, expose the gels to a phosphorimage cassette.
Exposure is generally done for at least 1 h and sometimes overnight.
12. Develop the gel per the manufacturer’s protocol (Molecular Dynamics). A typi-cal TRAP gel showing the 36 bp-internal standard and the characteristic 6-bp
ladder corresponding to telomerase activity is shown in Fig. 3 (see Note 8).
13. Quantitate telomerase activity relative to the internal standard by determining the
ratio of the entire telomerase ladder to the internal standard, relative to positive
control samples.
4. Notes
1. The slides can be stored at 4°C for up to 3 mo prior to pretreatment. Longer
storage may result in attenuation of the hybridization signal.
Fig. 3. Representative TRAP gel showing telomerase activity. On PCR amplifica-tion, the TRAP reactions are electrophoresed on 10% polyacrylamide gel electrophore-sis. In samples that contain telomerase activity, the characteristic telomerase ladder is
observed in 6-bp increments. Lane 1 serves as a negative control and contains only the
lysis buffer used for sample extraction. Lanes 2–4 show varying degrees of telomerase
activity, from high levels (lane 2) to lower levels (lane 3) to undetectable (lane 4).
Telomerase activity is quantified by taking the ratio of the internal standard to the
entire telomerase-specific ladder. Note that the difference between the amount of
telomerase activity in lanes 2 and 3 is based mainly on a comparison of the amplifica-tion of the internal standard relative to the TRAP ladder (see Note 9).
Detection of Telomerase by In Situ Hybridization and TRAP 57
2. The concentration of proteinase K varies among different tissue types. When
performing the technique for the first time, we recommend varying the protein-ase K concentrations (5–20 µg/mL) to determine the optimal conditions.
3. Acetic anhydride is highly unstable and should be added to the triethanolamine
immediately prior to incubation.
4. The linearized template must be completely free of all phenol. As an optional step,
we perform a chloroformisoamyl (241) extraction after the phenol extraction.
5. The exact duration of this step varies considerably from 2 wk to 4 mo. We find
that 3 wk is optimal for tissue sections and 2 wk is optimal for cultured cells.
6. We find that exposure of the slides to prolonged periods in water at room tem-perature causes the emulsion to lift off of the sections.
7. The RP/ACX, NT, and TSNT primers can be combined and stored at –20°C, in
such a way that 1 µL is equivalent to 100 ng of RP/ACX primer, 100 ng of NT
primer, and 0.01 amol of TSNT.
8. The TRAP assay is available as a convenient kit from Intergen (formerly Oncor)
called the TRAP-eze Telomerase Detection Kit. It contains all the components
for sterile detection telomerase activity in tissue samples and cultured cells using
the TRAP assay.
9. For some tumor samples, PCR inhibition can occur, which is easily visible by the
disappearance of the 36-bp internal standard. In cases of PCR inhibition, one of
two things can be done: (1) reextraction of the tumor sample, which can result in
elimination of PCR inhibitors and detectable telomerase activity; or (2) the
telomerase extension, followed by phenol chloroform extraction, ethanol pre-cipitation, and PCR amplification of telomerase extended products. Option 1 is
significantly easier than option 2, but it is also less reliable.
References
1. Shay, J. W. and Bacchetti, S. (1997) A survey of telomerase activity in human
cancer. Eur. J. Cancer 33, 787–791.
2. Kim, N. W., Piatyszek, M. A., Prowse, K. R., Harley, C. B., West, M. D., Ho,
P. L., Coviello, G. M., Wright, W. E., Weinrich, S. L., and Shay, J. W. (1994)
Specific association of human telomerase activity with immortal cells and cancer.
Science 266, 2011–2015.
3. Weinrich, S. L., Pruzan, R., Ma, L., et al. (1997) Reconstitution of human
telomerase with the template RNA component hTR and the catalytic protein sub-unit hTRT. Nat. Genet. 17, 498–502.
4. Olovnikov, A. M. (1971) Principle of marginotomy in template synthesis of poly-nucleotides. Doklady Biochem. 201, 394–397.
5. Watson, J. D. (1972) Origin of concatameric T4 DNA. Nat. New Biol. 239, 197–201.
6. Harley, C. B., Futcher, A. B., and Greider, C. W. (1990) Telomeres shorten with
aging. Nature 345, 458–460.
7. Hastie, N. D., Dempster, M., Dunlop, M. G., Thompson, A. M., Green, D. K., and
Allshire, R. C. (1990) Telomere reduction in human colorectal carcinoma and
with ageing. Nature 346, 866–868.
58 Morales and Holt
8. Lindsey, J., McGill, N., Lindsey, L., Green, D., and Cooke, H. (1991) In vivo loss
of telomeric repeats with age in humans. Mutat. Res. 256, 45–48.
9. Shay, J. W. and Wright, W. E. (1989) Quantitation of the frequency of immortal-ization of normal human diploid fibroblasts by SV40 large T-antigen. Exp. Cell
Res. 184, 109–118.
10. Shay, J. W., Pereira-Smith, O. M., and Wright, W. E. (1991) A role for both
RB and p53 in the regulation of human cellular senescence. Exp. Cell Res. 196,
33–39.
11. Wright, W. E. and Shay, J. W. (1992) The two-stage mechanism controlling cel-lular senescence and immortalization, Exp. Gerontol. 27, 383–389.
12. Ohmura, H., Tahara, H., Suzuki, M., Yoshida, M. A., Tahara, E., Shay, J. W.,
Barrett, J. C., and Oshimura, M. (1995) Restoration of the cellular senescence
program and repression of telomerase by human chromosome 3. Jpn. J. Cancer
Res. 86, 899–904.
13. Yashima, K., Piatyszek, M. A., Saboorian, H. M., Virmani, A. K., Brown, D.,
Shay, J. W., and Gazdar, A. F. (1997) Telomerase activity and in situ telomerase
RNA expression in malignant and non-malignant lymph nodes, J. Clin. Pathol.
50, 110–117.
14. Soder, A. I., Hoare, S. F., Muir, S., Going, J. J., Parkinson, E. K., and Keith, W. N.
(1997) Amplification, increased dosage and in situ expression of the telomerase
RNA gene in human cancer. Oncogene 14, 1013–1021.
15. Morales, C. P., Lee, E. L., and Shay, J. W. (1998) In situ hybridization for the
detection of telomerase RNA in the progression from Barrett’s esophagus to
esophageal adenocarcinoma. Cancer 83, 652–659.
16. Morales, C. P., Burdick, J. S., Saboorian, M. H., Wright, W. E., and Shay, J. W.
(1998) In situ hybridization for telomerase RNA in routine cytologic brushings
for the diagnosis of pancreaticobiliary malignancies.  Gastrointest. Endosc. 48,
402–405.
17. Ogoshi, M., Le, T., Shay, J. W., and Taylor, R. S. (1998)  In situ hybridization
analysis of the expression of human telomerase RNA in normal and pathologic
conditions of the skin. J. Invest. Dermatol. 110, 818–823.
18. Yashima, K., Litzky, L. A., Kaiser, L., Rogers, T., Lam, S., Wistuba, I. I.,
Milchgrub, S., Srivastava, S., Piatyszek, M. A., Shay, J. W., and Gazdar, A. F.
(1997) Telomerase expression in respiratory epithelium during the multistage
pathogenesis of lung carcinomas. Cancer Res. 57, 2373–2377.
19. Piatyszek, M. A., Kim, N. W., Weinrich, S. L., Keiko, H., Hiyama, E., Wright,
W. E., and Shay, J. W. (1995) Detection of telomerase activity in human cells and
tumors by a telomeric repeat amplification protocol (TRAP). Methods Cell Sci.
17, 1–15.
20. Holt, S. E., Norton, J. C, Wright, W. E., and Shay, J. W. (1996) Comparison of the
telomeric repeat amplification protocol (TRAP) to the new TRAP-eze telomerase
detection kit. Methods Cell Sci. 18, 237–248.
21. Feng, J., Funk, W. D., Wang, S. S., et al. (1995) The RNA component of human
telomerase. Science 269, 1236–1241.
Detection of Microsatellite Instability 59
59
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
7
Detection of Microsatellite Instability
Karin D. Berg, Constance A. Griffin, and James R. Eshleman
1. Introduction
In 1993, three groups independently discovered that the lengths of
microsatellites in tumors could vary from the normally constant pattern
defined at birth (1–3)  (see review in ref. 4 ). This discovery has been desig-nated either microsatellite instability (MSI) or replication errors (RER). A
recent international consensus conference convened by the National Cancer
Institute defined MSI/RER as “a change in length due to either insertion or
deletion of repeating units, in a microsatellite within a tumor when compared
to normal tissue” (5) . Microsatellites are regions of repetitive DNA in which
the repeating unit is small, varying in length from 1 to 6 nucleotides, and in
which the number of repeating units in a microsatellite can vary from 10–60
(6–7) . Because microsatellite lengths generally vary from person to person,
they have received widespread use in forensics, gene mapping, parentage test-ing, bone marrow engraftment testing, military remains testing, and so on.
Microsatellite loci (markers) are generally noncoding, lying in introns and
between genes. Analysis of microsatellites requires visualization of the num-ber of repeating units that a person has at a given microsatellite locus in his or
her genome (Fig. 1). Once a microsatellite marker is chosen for analysis, poly-merase chain reaction (PCR) primers are designed in which the first anneals
upstream of the microsatellite (forward primer) and the second anneals down-stream of the microsatellite (reverse primer). When a person’s DNA is ampli-fied by PCR using these primers, two bands are seen on a gel if the person is a
heterozygote, reflecting both paternal and maternal alleles. If the person is a
homozygote, possibly because the locus is invariant in the population (e.g.,
BAT25 and BAT26), only a single band will be seen. By comparing to molecu-lar weight markers (not shown in Fig. 1), one can extrapolate from the relative
band size to the precise number of repeating units in the microsatellite.
60 Berg et al.
During chromosome mapping experiments, microsatellite length was
serendipitously discovered to have deviated from normal in the tumors of cer-tain patients. As illustrated in Fig. 2A in a “normal” tumor, the alleles are the
same size in both the normal and the tumor DNA samples. By contrast, a tumor
with microsatellite instability shows novel alleles. Faint germline alleles are
often still visible owing to residual stromal cells despite microdissection of the
tumor from adjacent stromal tissue to enrich for tumor cell DNA. Stutter or
shadow bands arise during PCR owing to slippage (see Fig. 2B) because Taq
polymerase lacks proofreading function and DNA repair systems are not
present. Note that the difference between the parental band and the stutter bands
is exactly the number of bases of the repeating unit within the microsatellite
(e.g., microsatellites with dinucleotide repeats give rise to –2 and –4 base stut-ter bands). On first consideration, stutter bands seem distracting and could
obscure true MSI in which the shift in the tumor is small (e.g., –2 bases). How-ever, stutter bands are probably helpful because, for a given band, they serve as
a “signature” of what is contained within the PCR primers. For example, note
that in Fig. 2A, the true instability in the tumor sample from patient #2 can be
distinguished from the spurious band by the presence or absence of the signa-ture stutter bands. Interestingly, stutter is an in vitro artifact resulting from a
molecular mechanism somewhat analogous to the in vivo MSI/RER phenom-enon arising from defective mismatch repair (MMR).
Fig. 1. A theoretical “microsatellite” locus is shown in which an individual has
inherited a large paternal allele containing 4 CA repeats and a smaller maternal allele
containing 2 CA repeats. Forward (F) and reverse (R) primers are designed to anneal
at fixed genomic positions upstream and downstream of the microsatellite. Following
amplification, the number of repeating units within the microsatellite determines the
size of the PCR product and therefore the position of migration on a gel. The size of
the bands on the gel can thus be precisely correlated with the number of repeating
units within an individual’s alleles at any microsatellite locus. Very short
“microsatellites” are used in this illustration for clarity. Research Genetics (Hunts-ville, AL; http://www.resgen.com) currently has more than 5500 microsatellite loci
listed in their on-line catalog.
Detection of Microsatellite Instability 61
Microsatellite instability in tumors is a relatively recently discovered phe-nomenon whose diagnostic utility continues to expand (8) . It is a functional
surrogate marker for MMR defective cells (9 ,10) . MMR, first identified in yeast
and bacteria, is responsible for the rapid repair of mistakes made by DNA poly-merases. The principle clinical utility of the MSI/RER assay is currently for
the discovery of families with hereditary nonpolyposis colorectal cancer
Fig. 2. (A) Microsatellite assay (radioactive version, see text): Paired normal (N) and
tumor (T) samples are amplified from two patients at a single microsatellite locus. Patient
#1 demonstrates the pattern seen in a microsatellite stable (MSS) tumor, and patient #2
shows that seen with an MSI or RER tumor. Arrowheads, germline bands; dots, stutter
bands; arrows, shifted allele lengths, indicative of microsatellite instability; asterisk, non-specific PCR product. (B) Template strand slippage during synthesis of a CA repeat from
a 4 GT dinucleotide template repeat. The third GT is identified by an asterisk. The large
arrow symbolizes the DNA polymerase during active replication in which three of the CA
repeats have already been synthesized. Before the polymerase finishes synthesis, the third
GT of the template strand undergoes slippage and the third CA is now annealing with the
fourth GT. In the presence of competent DNA repair, this intermediate is recognized and
repaired such that both strands now contain the full-length repeat. If repair is defective, one
product contains a shortened length repeat.
62 Berg et al.
(HNPCC), which is owing to defects in MMR. Additionally, in colorectal can-cer, the presence of MSI in a tumor probably portends a better prognosis (2 ,11)
and may ultimately prove a predictor of tumor chemosensitivity. The follow-ing protocol to assay for MSI/RER is written primarily for use with a fluores-cent primer format, but the assay can also be performed using radiolabeled
PCR primers.
2. Materials
All materials used in specimen acquisition, DNA extraction, and PCR must
be handled using DNA/PCR precautions. Reagents should be PCR grade. Aero-sol-resistant tips should be used throughout.
2.1. Specimen Acquisition
1. Paraffin-embedded specimens:
a. N: normal block, no histologically identifiable lesions (the block containing
the margin of resection is often the best source) or a peripheral blood sample.
b. T: tumor block, histologically verified section of tumor.
2. Clean microdissection area containing the following equipment:
a. UV illumination source.
b. 37°C water bath.
c. Heating block to hold Eppendorf tubes.
d. Sterile Eppendorf tubes.
e. Sterile razor blades.
f. Microfuge.
g. Kimwipes.
2.2. DNA Extraction
1. Xylene.
2. 100% Ethanol.
3. Digestion buffer at the following final concentrations: 50 mM Tris-HCl (pH 8.5),
1 mM EDTA, 0.5% Tween-20, 200  µg/mL of proteinase K (Boehringer
Mannheim). Store buffer as single-use aliquots at –20°C.
2.3. Polymerase Chain Reaction
1. Thermocycler (PE 9600 or PE 9700).
2. Reagents: AmpliTaq Gold™ Polymerase (5 U/µL; Perkin Elmer/Roche),
AmpliTaq Gold Buffer containing 1.5 mM MgCl (Perkin Elmer/Roche), PCR-grade
water, 10 mM dNTPs (Perkin Elmer/Roche).
3. PCR Primers (Research Genetics; see Note 1 and Table 1). Fluorescent: forward
primer 5v labeled as indicated in  Table 1; radioactive: forward primer 5v end-labeled with 32P or 33P.
Detection of Microsatellite Instability 63
Table 1
Microsatellite Marker Characteristicsa
Genome Data Base/ Product
Locus (Fluor) primer sequence Genbank numbers size (bp)
BAT 25 (Tet)
Forward:
5′-TCGCCTCCAAGAATGTAAGT-3′ GDB:9834508/ 120d
Reverse: U63834
5′-TCTGCATTTTAACTATGGCTC-3′
BAT 26 (Hex)
Forward:
5′-TGACTACTTTTGACTTCAGCC-3′ GDB:9834505/ 116d
Reverse: U41210
5′-AACCATTCAACATTTTTAACCC-3’b
D2S123 (Hex)
Forward:
5′-AAACAGGATGCCTGCCTTTA-3′ GDB:187953/ 197–227e
Reverse: Z16551
5′-GGACTTTCCACCTATGGGAC-3′
D5S346 (Fam)
Forward:
5′-ACTCACTCTAGTGATAAATCGGG-3′ GDB:181171/   96–122e
Reverse: M73547
5′-AGCAGATAAGACAGTATTACTAGTT-3′
D17S250c (Fam)
Forward:
5′-GGAAGAATCAAATAGACAAT-3′ GDB:177030/ 151–169e
Reverse: X54562
5′-GCTGGCCATATATATATTTAAACC-3′
aSizes listed do not reflect addition of the nontemplated adenine. The primer sequences were
obtained from GDB. Single genbank numbers are listed even though the microsatellites may be
present in multiple entries.
bNote that this reverse primer has a one-base mispair from the gene sequence found in the
genbank database.
cThe locus D17S250 is also known as Mfd15.
dBAT25 and BAT26 (Big A-Tract) are generally invariant in size because these amplicons
have not been reported to be polymorphic in humans.
eThe base pair size ranges for the dinucleotide microsatellites do not in all cases include the
full range of sizes in the population. We have noted several germline alleles that are outside of
these ranges.
64 Berg et al.
2.4. Detection
1. Fluorescent: ABI 310 Genetic Analyzer or comparable capillary electrophoresis
apparatus, deionized formamide.
2. Radioactive: Standard DNA sequencing gel reagents and apparatus, autoradiog-raphy supplies.
3. Methods
Preventing cross contamination of cases is essential (individual disposable
pipet tips for each specimen must be used at each step).
3.1. Specimen Acquisition
1. Cut five sections from blocks submitted as normal (N, no tumor) and tumor (T). Sub-mit the first and last sections (5-µm sections) from each set for standard hematoxylin
& eosin (H&E) staining. Mount the three “sandwiched” middle sections (5–10 µm thick)
on standard histology slides unstained, without a cover slip (see Note 2).
2. Mark (“dot”) normal (N) and tumor (T) areas of the H&E slides (this should be
done by an anatomic pathologist).
3.2. Microdissection
1. Perform microdissection in a laminar flow hood, after the empty hood has under-gone UV treatment for at least 20 min.
2. Label 1.5-mL Eppendorf tubes: N for normal, T for tumor, and the specimen
number for each case.
3. Match the first unstained slide (level 2) grossly by tissue shape to the first (level
1) H&E-stained slide. Superimpose the unstained slide exactly over the stained,
marked slide, and using a razor or scalpel blade, microdissect the relevant (dot-ted) areas of the H&E slide from the unstained slide. Save levels 3 and 4 for
additional analyses if needed.
4. Place the section directly from the razor blade into the appropriately labeled 1.5-mL
Eppendorf tube after microdissection and seal the lid. Xylene may prevent static
electricity from affecting the tissue (see Note 3). Make certain that no paraffin or
tissue is left on the external aspect of the Eppendorf tube; wipe the closed tube
surfaces with xylene.
5. Clean all surfaces thoroughly with xylene between samples, and use fresh
Kimwipes and a new razor blade for each specimen. At no time should material
from one specimen (even from the same case) come in contact with another speci-men. Accurate results rely on a meticulous, clean microdissection technique.
6. Centrifuge the tissue to the bottom of the Eppendorf tubes, and microfuge for
2 min at the maximum setting (e.g., 12,800g for the Eppendorf 5414).
3.3. DNA Extraction
1. Remove paraffin using serial xylene washes. (Caution: xylene solubilizes most
inks. It is important to verify frequently that the specimen number is intact.) Add
1 mL of xylene to each Eppendorf tube containing microdissected tissue.
Detection of Microsatellite Instability 65
2. Mix the specimen tube contents by gently inverting each closed tube for 3 min.
3. Centrifuge in a microfuge at the maximum setting for 5 min.
4. Identify the pellet (be careful, it is usually translucent, and may be floating), and
pipet off the xylene.
5. Repeat xylene wash steps (steps 1–4) two additional times.
6. After the third xylene wash has been removed, add 1 mL of absolute (100%)
ethanol to each tube (to remove the residual xylene).
7. Mix the tube contents by gentle inversion for 3 min. The pellet should break up to
some extent.
8. Centrifuge in a microfuge as in step 3 for 5 min.
9. Remove the ethanol supernatant carefully (the pellet may still be difficult to
identify).
10. Repeat the ethanol wash one additional time (steps 6–9).
11. Remove the last ethanol wash and cover each uncapped tube with parafilm. Poke
several small holes in the parafilm (use an individual, sterile needle, and do not
touch the sides).
12. Dry the pellets approx 5 min in a speed-vac (Savant). Do not overdry, because
this can make DNA difficult to resuspend.
13. Depending on the size of the pellet, add 100–200 µL of thawed digestion buffer to the
dried pellets. Large samples require closer to 200 µL, and smaller pellets may require
100 µL or less to ensure sufficient postdigestion DNA concentrations.
14. Vortex each tube (see Note 4) to break up the pellet and spin for 10 s. Make sure
all tissue is in the buffer.
15. Incubate the specimens overnight at 37°C or for 3 h at 55°C.
16. Microfuge the tubes for 10 s (maximum setting) to collect the sample at the bot-tom of the tube.
17. Inactivate the proteinase K by boiling the specimen for 10 min or placing in a
95°C heat block for 10 min. Be sure to securely close the tube lids.
18. Microfuge for 1 min at maximum speed to collect the fluid at the bottom of the
tube.
19. Determine the A260 and A280 absorbance using a UV spectrophotometer. The DNA
concentration should ideally be between 30 and 100 µg/mL (see Note 5).
20. Samples should be stored in a non-self-defrosting –20°C freezer.
3.4. PCR Conditions
PCR conditions have been proven reliable using both the PE 9600 and the
PE 9700 thermocyclers.
1. Reaction mix (10  µL total volume, but should be master mixed and aliquoted
prior to sample addition): 4.7 µL of PCR-grade H2O; 1.0 µL of 10X AmpliTaq
Gold PCR buffer; 1.5 µL of forward primer (1.33 mM); 1.5 µL of reverse primer
(20 mM) (see Note 6); 0.2  µL of dNTPs (10 mM); 0.1  µL of AmpliTaq Gold
Polymerase (5 U/µL); 1.0 µL of sample, or control. For a description of primers,
see Table 1 and Notes 1 and 6.
66 Berg et al.
2. Controls (for each locus):
a. Water negative control.
b. “Normal” cell line (transformed lymphoblasts).
c. “Tumor” cell line from the same patient (known to be shifted at that locus).
3. Thermal-cycling conditions:
a. Initial denaturation: 95°C for 9 min.
b. Thirty-five cycles of denaturation at 94°C for 45 s, anneal at 55°C for 45 s,
and extension at 72°C for 1 min.
c. Final extension at 60°C for 45 min (adds nontemplated A).
3.5. Detection
3.5.1. Fluorescent
1. Aliquot to ABI 9600 PCR tubes 12.0  µL of deionized formamide, 0.75  µL of
internal size standard-Tamara (red), and 1.5 µL of PCR product.
2. Denature for 3–5 min at 95°C in a heat block (PCR block works well).
3. Load ABI 310 or other capillary electrophoresis apparatus per manufacturer’s
directions. The instrument settings for the ABI 310 are as follows:
a. Matrix: C.
b. Polymer: GS POP-4.
c. Size marker: Tamara labeled GS350 or GS500.
d. Buffer: 1X GS buffer.
e. Capillary temperature: 60°C.
f. Run time: 20 min.
g. Voltage: 15 kV.
h. Laser: 9.9 mW.
i. Smoothing: light.
j. Peak amplitude threshold: 200.
3.5.2. Radioactive
1. Use standard DNA sequencing gel and autoradiography.
3.6. Interpretation
Figure 3 provides an example of the MSI/RER assay using both the radio-active and fluorescent formats. Interpretation of this assay can appear straight-forward at first glance; however, one must be aware of potential pitfalls to
ensure accurate and consistent interpretation of the data. First, inspect the
water lanes for each primer set. There should be no banding or significant peak
height data in these lanes. Next, inspect the normal sample results (Fig. 3A and
Fig. 3B, top). Provided that there is no contamination, these bands or peaks
represent the germline microsatellite length(s) for the patient for each of the
loci (see Table 1 for expected amplicon size ranges). Note that BAT25 and
BAT26 are thought to be invariant in size in the population. True alleles can be
Detection of Microsatellite Instability 67
identified by a dominant band or peak (Fig. 3) surrounded by stutter (shadow)
peaks of diminishing intensity, which are generally smaller than the predomi-nant peaks. Peaks and bands without stutter should be very carefully evaluated,
because they almost invariably represent spurious amplification.
After identifying, recording, and verifying that the germline (nontumor)
amplicon sizes lie in the appropriate size range for each locus, look at the tu-mor electropherograms and determine the predominant amplicon sizes for each
locus in the tumor specimens (Fig. 3A and Fig. 3B; bottom). A direct compari-son between the germline amplicon sizes and tumor amplicon sizes is now
possible. For example, when one compares 1N to 1T, it is easy to appreciate
that this tumor has shifted from the germline pattern at the BAT25 locus. No
germline bands derived from residual stromal tissue are seen in the electro-pherogram of 1T shown in  Fig. 3A  or 3B, because 1N and 1T are cell line
controls and thus contain pure normal allele length or pure shifted allele length
PCR products. By contrast, residual germline bands are seen in Fig. 3A (2N
and 2T) that were amplified following microdissection of paraffin blocks.
Whereas the normal patient sample (2N) shows normal length BAT25 amplicon
lengths, the tumor tissue (2T) shows both shifted alleles and germline alleles,
the latter likely owing to contaminating germline stromal tissue.
After identifying heterozygous and homozygous alleles identifying the
germline and tumor amplicon sizes for each locus, one then counts the number
of markers showing instability. One may then make the appropriate diagnosis
using the guidelines from the National Cancer Institute (NCI) consensus con-ference (see ref. 5 ):
1. MSI high: two or more of five loci demonstrating instability.
2. MSI low: one of five loci demonstrating instability.
3. MSS/Non-RER: None of the markers showing instability.
The MSI high result most probably indicates a functional MMR defect in
the tumor. The human MMR genes currently known to be defective in tumors
are hMSH2, hMLH1, hPMS1, hPMS2, hMLH3, GTBP/hMSH6, and  hMSH3
(reviewed in refs. 9 and 10 ). However, this is an area of active investigation,
and consequently this list may not be inclusive. The finding of MSI high can be
due to a germline defect, such as exists in an HNPCC family, in combination
with a second hit in the tumor or can arise owing to biallelic inactivation in the
tumor in the absence of a germline defect. As a follow-up to this result, a
detailed family history, immunohistochemistry of the tumor for MMR protein
expression (at least hMSH2 and hMLH1 proteins), and germline DNA sequenc-ing for MMR defects (and testing for methylation status) may all be indicated.
It is probably appropriate to encourage genetic counseling when this result is
obtained (if not already established). While this protocol is written using the
68 Berg et al.
68
Fig. 3.
Detection of Microsatellite Instability 69
five primary loci recommended at the NCI consensus conference, MSI high
also may be diagnosed with a larger number of microsatellites in which the
percentage showing instability is *30–40% (5) .
The clinical implications of finding only one microsatellite shifted (MSI low) is
uncertain for two reasons. First, microsatellites are inherently difficult to replicate
and accordingly display a higher mutation rate than nonrepetitive regions of DNA
in the presence of functional repair. Therefore, having only one marker shifted
sometimes may be of no importance. Second, there are well-established “muted”
phenotypes, such as  GTBP/hMSH6, in which one would expect low-level
microsatellite instability. A defect in  GTBP produces selective mononucleotide
instability in the presence of stable dinucleotides. The study of additional markers
in this setting may be appropriate to assess the significance of one of five markers
shifted. When more markers are analyzed, MSI low is designated when <30–40%
of markers analyzed are found to be shifted (5).
4. Notes
1. Primers that we currently use are those recommended by the NCI consensus con-ference and are synthesized by Research Genetics. The Food and Drug Adminis-tration is interested in ensuring the quality of oligonucleotide primers used in
“home-brew” assays. Accordingly, one may need to purchase oligonucleotides
only from vendors approved to provide these as analyte-specific reagents. Irre-spective of the source of the primers, it is important to run the primers themselves
on the column when they are obtained to exclude contamination resulting in
extraneous bands.
2. The laboratory needs to ensure that tissue and DNA are not contaminated with
any other specimen. If this occurs, novel bands representing foreign alleles could
be inappropriately misinterpreted as demonstrating MSI/RER.
Fig. 3. (see opposite page) (A) Autoradiograph of BAT25 amplified from a tumor
bearing a defect in the mismatch repair gene, hMSH2. The normal sample is derived
from transformed lymphoblasts (L670) from this patient, and the tumor sample is a
cell line (Vaco 670) derived from the patient’s tumor. Both cell lines were generous
gifts from Dr. James Willson at Case Western Reserve University. Samples 2N and 2T
from the second patient consist of normal colonic epithelium (2N) and colonic adeno-carcinoma (2T), prepared from paraffin blocks as per the protocol described herein.
(B) Electropherograms of amplicons of BAT25 amplified from cell lines 1N and 1T as
shown in (A). Red peaks are size standard peaks at 75, 100, 139, 150, and 160 in which
the x-axis size is in base pairs. Peak heights are quantified as random fluorescent units
(RFUs) on the y-axis. The green peaks at and around 120 bp (normal, arrowhead) and
116 bp (tumor shifted, arrow) represent the BAT25 amplicons. The predominant peak
in 1N (arrowhead) represents the expected amplicon length and stutter (owing to slip-page) is visualized as serration extending above and below the predominant peak.
70 Berg et al.
3. One problem encountered during microdissection is static electricity, which can
cause the microdissected tissue to fly off the razor blade. This can be avoided by
applying 15 µL of xylene to the tissue on the slide prior to microdissection.
4. Vortexing solutions containing protein promotes denaturation and generally is
not done. However, vortexing at this step is empirically advantageous presum-ably because fragmenting the tissue outweighs the protein denaturation.
5. A formal A260 / A280 reading is suggested. Because there is no precipitation step in
the protocol, this reading should be inaccurate owing to the presence of amino
acids, which should still contribute to absorbance. It is useful nonetheless
because it is a crude measurement of the approximate amount of DNA that has
been isolated. We have empirically found that a more extensive DNA prepara-tion is not necessary.
6. We have done a substantial number of titration experiments testing various rela-tive primer ratios. At the labeled unlabeled primer ratio of 1 1, one gets many
spurious bands, whereas at 1 40, one begins to see loss of signal. Ratios of 1 15
or 1 20 appear optimal in both regards (lack of spurious amplification and
robustness of specific signal).
References
1. Aaltonen, L., Peltomaki, P., Leach, F., Sistonen, P., Pylkkanen, L., Mecklin, J.,
Jarvinen, H., Powell, S., Jen, J., Hamilton, S., Petersen, G., Kinzler, K.,
Vogelstein, B., and de la Chapelle, A. (1993) Clues to the pathogenesis of familial
colorectal cancer. Science 260, 812–816.
2. Thibodeau, S., Bren, G., and Schaid, D. (1993) Microsatellite instability in cancer
of the proximal colon. Science 260, 816–819.
3. Ionov, Y., Peinado, M., Malkhosyan, S., Shibata, D., and Perucho, M. (1993)
Ubiquitous somatic mutations in simple repeated sequences reveal a new mecha-nism for colonic carcinogenesis. Nature 363, 558–561.
4. Eshleman, J. R. and Markowitz, S. D. (1995) Microsatellite instability in inher-ited and sporadic neoplasms. Curr. Opin. Oncol. 7, 83–89.
5. Boland, C. R., Thibodeau, S. N., Hamilton, S. R., Sidransky, D., Eshleman, J. R.,
Burt, R. W., Meltzer, S. J., Fodde, R., Rodriguez-Bigas, M. A., Fodde, R.,
Ranzani, G. N., and Srivastava, S. (1998) A National Cancer Institute workshop
on microsatellite instability for cancer detection and familial predisposition:
development of international criteria for the determination of microsatellite insta-bility in colorectal cancer. Cancer Res. 58, 5248–5257.
6. Litt, M. and Luty, J. A. (1989) A hypervariable microsatellite revealed by in vitro
amplification of a dinucleotide repeat within the cardiac muscle actin gene. Am. J.
Hum. Genet. 44, 397–401.
7. Weber, J. L. and May, P. E. (1989) Abundant class of human DNA polymor-phisms which can be typed using the polymerase chain reaction,  Am. J. Hum.
Genet. 44, 388–396.
8. Rodriguez-Bigas, M., Boland, C. R., Hamilton, S. R., Henson, D. E., Jass, J. R.,
Khan, P. M., Lynch, H., Perucho, M., Smyrk, T., Sobin, L., and Srivastava, S.
Detection of Microsatellite Instability 71
(1997) A National Cancer Institute workshop on Hereditary Nonpolyposis
Colorectal Cancer syndrome: meeting highlights and Bethesda guidelines. JNCI
89, 1758–1762.
9. Eshleman, J. R. and Markowitz, S. D. (1996) Mismatch repair defects in human
carcinogenesis. Hum. Mol. Genet. 5, 1489–1494.
10. Toft, N. J. and Arends, M. J. (1998) DNA mismatch repair and colorectal cancer.
J. Pathol. 185, 123–129.
11. Sankila, R., Aaltonen, L. A., Jarvinen, H. J., and Mecklin, J. P. (1996) Better
survival rates in patients with  MLH1-associated hereditary colorectal cancer.
Gastroenterology 110, 682–687.
PCR Clonality Analysis Based on X-Linked Genes 73
73
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
8
Polymerase Chain Reaction Clonality
Assays Based on X-Linked Genes
Langxing Pan and Huaizheng Peng
1. Introduction
During embryogenesis in females, in each cell either the paternal or the
maternal X chromosome is randomly inactivated through methylation  (1,2) .
This event is stably inherited by daughter progeny of each cell. Therefore, in
adult women, polyclonal cell populations will comprise a random mixture of
paternally and maternally derived X-inactivated cells, but monoclonal cells
will contain the same inactivated X chromosome as their progenitor. The
paternal and maternal X chromosomes can be distinguished by identification
of polymorphisms at certain alleles, and differences in methylation of DNA
sequences within these alleles can be detected by digestion of the DNA with
methylation-sensitive restriction enzymes. Based on these principles, research-ers have developed several X-linked clonality assays, and most are performed
using Southern blot hybridization (3–5) .
The polymorphic and methylation sites of some X-linked genes are closely
clustered within a short region, which allows clonality analysis by polymerase
chain reaction (PCR) after methylation-sensitive enzyme digestion. One of the
early targets is the phosphoglycerate kinase gene (6) . However, the usefulness
of this locus as a clonal marker is limited by its low frequency of polymor-phism (20–40%) (3) . Recently, two other genes, the human androgen receptor
gene (AR gene or HUMARA)  (7) and monoamine oxidase A gene (MAOA
gene) (8 ,9) have been reported to have a very high rate of heterozygosity
(AR 90%, MAOA 75%) (7 ,9 ,10) . In an earlier study, based on the inactivation
patterns of these genes, we developed two PCR assays for detection of clonality
in female patients (11) . We describe these assays in detail in this chapter.
74 Pan and Peng
2. Materials
2.1. DNA Samples
Prepare high molecular weight DNA using the standard proteinase K/phenol/
chloroform method (12) or a commercial kit (Puregene; Gentra Systems, Min-neapolis, MN). DNA from a male individual can serve as a control for endonu-clease digestion efficiency.
2.2. Restriction Endonuclease
Use HpaII, a methylation-sensitive restriction endonuclease (Boehringer
Mannheim, Mannheim, Germany) for both AR and MAOA PCR analyses.
2.3. Polymerase Chain Reaction
1. 10X PCR buffer (supplied with Taq DNA polymerase or cat. no. M190A; Pro-mega, Madison, WI): 100 mM Tris-HCl (pH 9.0), 500 mM KCl, 0.1% Triton X-100.
2. Taq DNA polymerase (5 U/µL) (cat. no. M1862A; Promega).
3. 25 mM each dNTP (cat. no. U1240; Promega).
4. 25 mM MgCl2 (supplied with the Taq polymerase or cat. no. A351; Promega)
5. Primers: (5v to 3v)
a. AR sense primer (AR1): GCT GTG AAG GTT GCT GTT CCT CAT
b. AR antisense primer (AR2): TCC AGA ATC TGT TCC AGA GCG TGC
c. MAOA sense primer (MAOA1): ACA TTC TAA ACC TAA TAA CTC
d. MAOA antisense primer (MAOA2): CAA TAA ATG TCC TAC ACC TT
e. Inner MAOA antisense primer (MAOA3): GGT AGA CTC CTT TAA GAA
AAG GTT AAA A
2.4. Denaturing Polyacrylamide Gel Electrophoresis
1. SequaGel™ sequencing system consisting of SequaGel™ diluent, SequaGel™
buffer, and SequaGel™ concentration (EC-833; National Diagnostics, Atlanta, GA).
2. Ammonium persulfate (A-9164; Sigma, St. Louis, MO).
3. TEMED (T-7024; Sigma).
4. 10X TBE: 0.9 M Tris-HCl, pH 8.3, 0.9 M boric acid, 0.02 M EDTA.
5. Loading buffer (sequencing stop buffer): 98% formamide, 10 mM NaOH, 20 mM
EDTA, 0.05% bromophenol blue, and 0.05% xylene cyanol FF.
2.5. Background-Free Silver Staining
1. Acetic acid (100016; BDH).
2. Methanol (10158; BDH).
3. Formaldehyde (10113; BDH).
4. Ethanol (10107; BDH).
5. Sodium thiosulfate (72049; Fluka).
6. AgNO3 (S-0139; Sigma).
7. Na2CO3 (10240; BDH).
8. Glass trays (30 × 30 cm or larger).
PCR Clonality Analysis Based on X-Linked Genes 75
9. 3MM filter paper (3030917; Whatman).
10. Cling film.
11. Cellophane membrane, optional.
2.6. Equipment
1. Thermocycler (oil free).
2. Protean II xi vertical electrophoresis cell with gel mold (200  × 240 mm) and
0.75-mm thick spacers and comb (165-1933; Bio-Rad, Hercules, CA).
3. Vacuum gel dryer, optional (Bio-Rad).
3. Methods
Figure 1 illustrates the principles of both methods. Both digested and undi-gested DNA samples are analyzed in parallel. To obtain accurate results, it is
crucial to ensure a complete digestion of DNA samples. Because male indi-viduals have only one X chromosome, DNA from a male can serve as a control
of digestion efficiency (see Subheading 3.6.).
Fig. 1. AR and MAOA gene structure and PCR strategies. PCR primers are
designed to flank the STR and methylation sites of the AR and MAOA genes. Ampli-fication of this region by PCR is performed from both undigested and HpaII-digested
DNA samples in each case. PCR performed on undigested DNA samples amplified
both methylated (inactive) and unmethylated (active) alleles of the gene and disclosed
whether the sample is polymorphic. By contrast, PCR of the digested DNA samples
amplified only the methylated allele of the gene because the unmethylated allele is
destroyed by the restriction enzyme. Amplification of trinucleotide repeat and methy-lation sites of AR genes is performed in a single round of PCR. Amplification of
dinucleotide repeat and methylation sites of MAOA genes is performed in two rounds
of PCR. (Reproduced from ref. 11  with permission. © John Wiley & Sons Limited.)
76 Pan and Peng
3.1. DNA Digestion
1. Digest 1 µg of DNA in a 20-µL vol with 10 U of HpaII at 37°C overnight.
2. Dilute the digested DNA to a concentration of 20 ng/µL with water.
3.2. PCR to Amplify AR Gene (HUMARA)
1. Prepare a master mix (based on a total 25-µL vol for each reaction) containing
1X PCR buffer; 1.5 mM MgCl2; 0.2 mM each of dATP, dCTP, dGTP and dTTP;
0.2 µM of each primer (AR1 and AR2); and an appropriate amount of Taq DNA
polymerase (0.2 U/25 µL of final reaction volume). Place 24 µL of the master
mix into individual microtubes.
2. Add 1 µL of digested or undigested sample DNA (20 ng) to each tube and per-form 30 cycles of denaturation at 95°C for 30 s, primer annealing at 54°C for
30 s, and primer extension at 72°C for 45 s. Conclude the reaction with a final
extension at 72°C for 5 min.
3.3. PCR on MAOA Gene
For the MAOA gene, two rounds of PCR are performed using the same
master mix used for HUMARA with different primers. In the first round,
MAOA1 and MAOA2 primers are used to amplify the fragment containing
both the simple tandem repeat (STR) and methylation sites in the DNA
samples. In the second round of PCR, MAOA2 and MAOA3 primers are used
to amplify the STR site.
1. For the first round of PCR, add 24 µL of the master mix (see Subheading 3.2.,
step 1) with MAOA1 and MAOA2 primers to each tube and add 1 µL (20 ng) of
digested or undigested DNA sample.
2. Perform 20 PCR cycles of denaturation at 95°C for 30 s, primer annealing at
54°C for 30 s, and primer extension at 72°C for 45 s.
3. Dilute a fraction of the first-round PCR products 100-fold with water. Add 1 µL
of the diluted first-round products to each tube containing 24 µL of the master
mix with MAOA2 and MAOA3 primers. Run the second PCR using the same
cycling conditions as for the first round for a further 30 cycles. Conclude the
reaction with a final extension at 72°C for 5 min.
3.4. Polyacrylamide Gel Electrophoresis
1. For a 200 × 240 × 0.75 mm, 8% denaturing polyacrylamide gel, mix 58 mL of
SequaGel diluent, 10 mL of SequaGel buffer, and 32 mL of SequaGel concentra-tion in a beaker.
2. Add 500 µL of 10% ammonium persulfate and 50 µL of TEMED to the beaker
and mix gently.
3. Quickly pour the mixture into the gel mold and allow it to set at room tempera-ture for 1 h.
4. Assemble the electrophoresis apparatus with the gel and fill the tank with 1X
TBE buffer.
PCR Clonality Analysis Based on X-Linked Genes 77
5. Dilute 5 µL of each PCR product with 15 µL of loading buffer and load 10 µL of
sample per track onto the gel.
6. Run the gel at 40 W constant power for 2–3 h at 40°C maintained by flowing hot
tap water through the built-in water jacket of the gel apparatus (see Note 1).
3.5. Background-Free Silver Staining
1. Fix the gel in a solution containing 12% acetic acid, 50% methanol, and 0.02%
formaldehyde for 2–16 h with gentle agitation.
2. Wash the gel with 50% ethanol for 20 min twice.
3. Place the gel into freshly prepared 0.02% sodium thiosulfate solution for 1 min,
and rinse the gel with distilled water three times.
4. Soak the gel in silver solution containing 0.2% AgNO3 and 0.03% formaldehyde
for 20–30 min and then rinse twice with distilled water.
5. Soak the gel in a solution containing 6% Na2CO3, 0.02% formaldehyde, and
0.0005% sodium thiosulfate for 3–5 min, to develop the color.
6. Stop the reaction by placing the gel into a solution containing 50% methanol and
16% acetic acid.
7. Transfer the gel onto a piece of 3MM filter paper or cellophane membrane. Cover
the gel with cling film, and dry it in a vacuum gel dryer at 80°C for 40 min.
3.6. Interpretation of Results
Undigested control male DNA should show a single band for both AR and
MAOA STR sites, whereas completely digested male DNA should show no
PCR products. DNA from homozygous (noninformative) females should show
a single band with or without enzyme digestion. In heterozygous (informative)
females, DNA from polyclonal cell populations shows two bands with
equal intensity with or without enzyme digestion. Undigested DNA of mono-clonal cell populations from an informative female shows two bands with equal
intensity, whereas completely digested DNA shows a single band if the DNA
is from a pure cell population such as a cell line, or two bands with unequal
intensity if the sample contains normal or polyclonal cells (see Fig. 2).
It has been reported that similar PCR methods based on AR gene can be
applied to minute amounts of paraffin-embedded tissue (13) . We find that com-plete digestion of DNA from paraffin section is extremely difficult to achieve.
However, DNA purified from microdissected fragments of frozen tissue sec-tions can be adequately digested in the presence of both  HpaII and  HhaI
restriction enzymes (unpublished data) and used for the AR gene–based PCR
clonality assay. Further experiments for the reliability and reproducibility in
the microdissected frozen materials are needed.
A skewing phenomenon is the major disadvantage of any clonality assay based
on X-linked gene  (14,15) , especially when DNA samples are prepared from
hemopoietic cells of elderly females (16). When dealing with these materials, it is
important to have DNA from samples of the patient’s normal tissue as control.
78 Pan and Peng
4. Note
1. The 8% denaturing polyacrylamide gel may be stained with ethidium bromide.
Electrophoresis may be run at room temperature, but unexpected bands caused
by heteroduplex DNA in the gel may occur and confuse the results.
References
1. Mandel, J. L., Manaco, A. P., Nelson, D. L., Schlessinger, D., and Willard, H.
(1993) Genome analysis and the human x chromosome. Science 258, 103–109.
2. Riggs, A. D. and Pfeifer, G. P. (1992) X-chromosome inactivation and cell
memory. Trends. Genet. 8, 169–174.
3. Vogelstein, B., Fearon, E. R., Hamilton, S. R., Preisinger, A. C., Willard, H. F.,
Michelson, A. M., Riggs, A. D., and Orkin, S. H. (1987) Clonal analysis using
recombinant DNA probes from the X-chromosome. Cancer Res. 47, 4806–4813.
4. Lucas, G. S., Padua, R. A., Masters, G. S., Oscier, D. G., and Jacobs, A. (1989)
The application of X-chromosome gene probes to the diagnosis of myeloprolif-erative disease. Br. J. Haematol. 72, 530–533.
5. Fey, M. F., Peter, H. J., Hinds, H. L., Zimmermann, A., Liechti Gallati, S., Gerber,
H., Studer, H., and Tobler, A. (1992) Clonal analysis of human tumors with M27
beta, a highly informative polymorphic X chromosomal probe. J. Clin. Invest. 89,
1438–1444.
Fig. 2. Clonality detection by (A) MAOA-PCR and (B) AR-PCR. Case 1, control
DNA from a male; case 2, DNA from a female, showing homozygous; case 3, tonsil
DNA from a female, showing polyclonality; and case 4, breast carcinoma DNA from
a female, showing monoclonality. Lanes (–) and (+) show PCR amplification before
and after  HpaII digestion, respectively. (Reproduced from  ref. 11 with permission.
© John Wiley & Sons Limited.)
PCR Clonality Analysis Based on X-Linked Genes 79
6. Gilliland, D. G., Blanchard, K. L., Levy, J., Perrin, S., and Bunn, H. F. (1991)
Clonality in myeloproliferative disorders: analysis by means of the polymerase
chain reaction. Proc. Natl. Acad. Sci. USA 88, 6848–6852.
7. Allen, R. C., Zoghbi, H. Y., Moseley, A. B., Rosenblatt, H. M., and Belmont,
J. W. (1992) Methylation of HpaII and HhaI sites near the polymorphic CAG
repeat in the human androgen-receptor gene correlates with X chromosome inac-tivation. Am. J. Hum. Genet. 51, 1229–1239.
8. Hornstra, I. K. and Yang, T. P. (1992) Multiple in vivo footprints are specific to
the active allele of the X-linked human hypoxanthine phosphoribosyltransferase
gene 5vregion: implications for X chromosome inactivation. Mol. Cell. Biol. 12,
5345–5354.
9. Hendriks, R. W., Chen, Z. Y., Hinds, H., Schuurman, R. K. B., and Craig, I. W.
(1992) An X chromosome inactivation assay based on differential methylation of
a CpG island coupled to a VNTR polymorphism at the 5vend of the monoamine
oxidase A gene. Hum. Mol. Genet. 1(3), 187–194.
10. Willman, C. L., Busque, L., Griffith, B. B., Favar, B. E., McClain, K. L.,
Duncan, M. H., and Gilliland, D. G. (1994) Langerhans’-cell histocytosis
(Histocytosis X)—a clonal proliferative disease. N. Engl. J. Med. 331, 154–160.
11. Peng, H. Z., Du, M. Q., Diss, T. C., Isaacson, P. G., and Pan, L. X. (1997) Clonality
analysis in tumours of women by PCR amplification of X-linked genes. J. Pathol.
181, 223–227.
12. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989)  Molecular Cloning:
A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Har-bor, NY.
13. Mashal, R. D., Lester, S. C., and Sklar, J. (1993) Clonal analysis by study of
X chromosome inactivation in formalin-fixed paraffin-embedded tissue. Cancer
Res. 53, 4676–4679.
14. Gale, R. E. and Wainscoat, J. S. (1993) Clonal analysis using X-linked DNA poly-morphisms. Br. J. Haematol. 85, 2–8.
15. Gale, R. E., Wheadon, H., Boulos, P., and Linch, D. C. (1994) Tissue specificity
of X-chromosome inactivation patterns. Blood 10, 2899–2905.
16. Gale, R. E. and Linch, D. C. (1998) Clonality studies in acute myeloid leukaemia.
Leukaemia 12, 117–120.
Fluorescent In Situ Hybridization 81
81
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
9
Fluorescent In Situ Hybridization
Evaluation for Ploidy and Gene Amplification
Susan Sheldon
1. Introduction
In situ hybridization was first described in the late 1960s by Pardue and Gall
(1) , who hybridized mouse ribosomal DNA sequences to a mouse chromo-some spread. The technique came into broader use with the description of DNA
probes for various viral sequences, and in the late 1980s with the publications
of Lichter and Ward (2) , Pinkel et al. (3) , and others (4–7) on the use of fluo-rescence in situ hybridization (FISH) probes. To perform FISH, or an in situ
hybridization technique, a DNA sequence is prepared with its thymidine tagged
with a compound such as fluorescein (direct labeling), biotin, or digoxigenin to
create a probe for a given sequence located on a specific chromosome. Both
the probe and the fixed cellular DNA are denatured using a combination of
heat and formamide, and allowed to renature together. The nonspecifically
bound probe is removed, and the probe-cellular DNA complex is visualized
directly, with a fluorochrome-labeled avidin or a fluorochrome-tagged anti-digoxigenin. For general reviews of the technique, see refs. 8–10 .
This chapter supplies background information for persons using commer-cially prepared FISH probes. As with many commercial reagents, they do not
always perform as advertised, nor are the directions always complete or cor-rect. To illustrate the differences between using unique sequence and repeated
sequence probes, I discuss two examples. The first is an enumeration of the
copies of several chromosomes using an alpha satellite probe in the placenta
from a patient with a possible partial molar pregnancy. Three copies of mul-
82 Sheldon
tiple chromosomes are suggestive of a triploid (69 chromosome) karyotype
(11 ,12) . This example also illustrates the issues associated with working with
paraffin-embedded tissue  (13 ,14) . The second example involves the
quantitation of the number of copies of the gene for N-myc in a touch imprint
from a patient with neuroblastoma.
In the first example, the distinction between a complete mole vs a partial
mole vs hydropic changes secondary to a fetal demise can be difficult to make
on histologic grounds alone. Yet, the management of the patient is quite differ-ent depending on the final diagnosis. Often the discovery of hydropic villi is
made after the entire specimen is in formalin, precluding standard cytogenetic
analysis. However, the question is raised: Is this a complete mole, with 46
chromosomes, or a partial mole, with 69, or merely a hydropic placenta with a
badly macerated fetus (11) ? FISH, using several alpha satellite probes, can at
least approach the question of how many chromosomes are present. In the study
patient (Fig. 1) an alpha satellite probe for the centromere of chromosome 7
was used; on scoring 250 nuclei, most had two fluorescent signals from the
probe, suggesting either a complete mole or normal tissue.
In the second example, the N-myc gene is normally present on chromosome
2; there are two copies per nucleus. In patients with neuroblastoma, the gene
will often be amplified or present in many copies in the tumor, but not in the
patient’s somatic cells. The DNA sequence within the gene itself is not ampli-fied or highly repeated, however, so the probe used is a unique sequence probe.
Those patients with multiple copies of this gene have an unfavorable progno-sis, regardless of the histology or stage (15–17) . The study patient’s tumor had
multiple copies of the N-myc gene (Fig. 2) in the 20 cells scored. An unfavor-able prognosis is associated with this finding.
There are several different types of DNA probes:
1. Alpha satellites—These sequences are at the centromeres of each chromosome.
They are specific to that chromosome, so, e.g., there is a different alpha satellite
for chromosome 1 and chromosome 10. Their sequences are highly repeated,
making them “sticky” and subject to nonspecific binding. A high-stringency,
posthybridization wash is required to achieve specificity, i.e., so that chromo-some 1 probes do not bind to those of chromosome 10. These probes are among
the easiest to work with, and are readily used for chromosome enumeration, pro-vided that one is not looking for translocations.
2. Painting probes—These contain a collection of unique sequences specific for a
given chromosome. The precise identity of the genes in this mixture of probes is
not usually known. Depending on the density of the given sequences, the
so-called paint may cover the entire chromosome with equal intensity. These
probes are useful for characterizing chromosomal translocations, but are less use-ful for paraffin sections.
Fluorescent In Situ Hybridization 83
3. Unique sequence probes—These are probes for specific genes or diseases closely
linked to a known chromosomal location. They are available for a variety of
microorganisms, genetic diseases (such as DiGeorge syndrome), and malignan-cies (such as  N-myc in neuroblastoma). In general, probes for microdeletions
associated with genetic diseases contain both a probe for the disorder of interest
and a marker probe for the chromosome on which the disease gene is located.
These probes require a posthybridization wash of lower stringency than that used
for the alpha satellite probes.
Fig. 1. Section through a formalin-fixed, paraffin-embedded section of a placenta
showing hydropic changes consistent with a partial mole. Most nuclei show two
brightly fluorescent signals, although not necessarily in the same focal plane.
84 Sheldon
Unlike Southern and polymerase chain reaction-based analyses, much of
the specificity of FISH probes is derived from the posthybridization wash. The
concept of “stringency” has been alluded to. This refers to the ability of DNA
sequences to bind to one another when the sequence homology is not exact; for
instance, sequences that are rich in adenine and thymine (A-T) and highly
repeated will often hybridize to one another. To eliminate this nonspecific bind-ing, one would use a high-stringency wash. Factors that increase the stringency
and make it more difficult for inexactly matched hybrids to remain bound
include increasing the temperature (from 37 to 43°C), increasing the formamide
concentration (from 50 to 70%), and decreasing the salt concentration (from
0.4 to 0.25X saline sodium citrate [SSC]).
Fig. 2. Touch or imprint preparation of a neuroblastoma probed with N-myc. There
are multiple bright signals over the nucleus, extending into the cytoplasm, suggesting
that the gene is highly amplified. All cells should show two signals in the nucleus
corresponding to the location of the normal gene.
Fluorescent In Situ Hybridization 85
2. Materials
2.1. Tissues
For analysis of ploidy in placenta, cut routine 4-µ paraffin sections and place
on positively charged slides, heat in a 60°C oven for no more than 15 min. The
internal controls, both positive and negative, are blood cells present in the
specimen, which should have a diploid number of signals.
For the evaluation of N-myc amplification, make a series of touch or imprint
preparations of the neuroblastoma fresh tissue sample. One slide should be
fixed and stained with Wright-Giemsa to confirm the presence of tumor, and
the remaining slides are fixed as in Subheading 3.1.2., then processed begin-ning with the denaturation step as in  Subheading 3.2. Internal controls
include the presence of two signals in the majority of cells (corresponding to
the normal cellular gene) and the absence of multiple signals in stroma and
nucleated blood cells.
2.2. Fixing Smears and Imprints
1. Methanol.
2. Glacial acetic acid.
3. A mixture of 3 parts methanol to 1 part acetic acid (“acid alcohol”) is prepared
and used within 1 h.
2.3. Dewaxing Paraffin Sections
1. Xylene.
2. Absolute ethanol, 95% ethanol, 80% ethanol, 70% ethanol.
3. Deionized water.
2.4. Thiocyanate Pretreatment of Paraffin Sections
1. 1 M Sodium thiocyanate solution, store protected from light.
2.5. Enzymatic Digestion of Paraffin Sections
Prepare the solutions using sterile reagents and containers; thaw at 37°C and
use immediately.
1. Pepsin (4 mg/mL) in 0.2 M HCl (store stock at –20°C) or proteinase K (25 µg/mL) in
phosphate-buffered saline (store stock at –20°C).
2.6. Denaturation
Prepare the following solutions using sterile water and containers:
1. 20X SSC: 3 M NaCl (175.32 g/L) and 0.3 M sodium citrate (88.23 g/L) in deion-ized water. Dissolve the two salts separately and mix. This is a stock solution.
The pH will need to be adjusted to 6.8–7.0 on dilution.
2. 70% Formamide, pH 7.0, in 2X SSC (see Note 1).
86 Sheldon
2.7. Probes
The probes used are commercially available and should be used with the
manufacturers’ hybridization buffer. The exact ingredients of the hybridiza-tion buffers are proprietary, but they generally contain 50% formamide, 10%
dextran sulfate, 0.01% sheared salmon sperm DNA in 2X SSC. The  N-myc
probe (Oncor, Gaithersburg, MD) does not require dilution. The alpha satellite
or chromosome enumeration probe (Cytocell, Ltd., Oxfordshire, UK) is sup-plied affixed to a plastic cover slip and comes with hybridization buffer.
2.8. Hybridization
1. Hybridization chamber: These are available commercially, or you can use a plas-tic box with a lid or a glass baking dish covered with plastic wrap. Place one or
two damp paper towels in the bottom to maintain humidity.
2. Rubber cement.
3. 37°C incubator without CO2, 37°C oven, or hot plate.
2.9. Posthybridization Wash
1. 50% Formamide in 2X SSC, pH 7.0, or 0.4X SSC, pH 7.0.
2. Phosphate-buffered detergent (PBD): 130 mM NaCl, 7 mM dibasic sodium phos-phate, 3 mM monobasic sodium phosphate, pH 6.8. Dissolve salts in deionized
water in the order given, and then add 0.05% Triton X-100 (0.05 mL/L).
2.10. Localization of Probe
1. Antidigoxigenin antibody labeled with appropriate fluorochrome or avidin
labeled with appropriate fluorochrome as supplied by the manufacturer.
2. Counterstains include 4′,6-diamidino-2-phenylindole (DAPI) or propidium
iodide in Antifade (Sigma, St. Louis, MO).
3. Methods
3.1. Preparing Target Tissue
When working with paraffin-embedded tissue, one must remove the paraf-fin (deparaffinization or dewaxing), the protein that has been crosslinked to the
DNA, and the DNA crosslinks. Many manufacturers of DNA probes recom-mend that the sections not be baked at 60°C; this depends on the probe. It may
be helpful to bake for 15 min or so to improve adhesion of the section to the
slide. Slides should be positively charged, such as silanized slides, to improve
adhesion.
3.1.1. Paraffin Sections
1. Dewax tissues by immersion in a series of Coplin jars at room temperature as
follows: xylene, two changes for 10 min each; xylene ethanol (50 50) 5 min;
ethanol, two changes for 5 min each; air-dry at least 5 min.
Fluorescent In Situ Hybridization 87
2. For thiocyanate pretreatment, incubate slides for 10 min at 80°C in 1 M sodium
thiocyanate. This can be done on a slide warmer. Flood the slide with about 2 mL
of the reagent to avoid evaporation. Following incubation, rinse the slide with
deionized water, two changes for 2 min each. Briefly drain the slides to remove
excess water.
3. Incubate the slide at 37°C for 5–15 min in pepsin solution (this may require up to 45
min; see Note 2). The placental sample in the illustrated case was treated for 20 min.
4. Rinse with deionized water, two changes for 2 min each.
5. Dehydrate through graded alcohols (70, 80, 95, and 100%) and allow to air-dry.
This is critical to avoid diluting the formamide in the denaturation step. Proceed
to Subheading 3.2.
3.1.2. Smears and Imprints
1. Fix cells by placing the slides on a horizontal surface and covering the surface
with acid alcohol for 2 min.
2. Drain excess liquid by tilting the slide.
3. Air-dry for at least 1 h.
3.2. Denaturation
Generally, both the probe and the target DNA will require denaturation, but
this depends on the manufacturer’s instructions. Some probes require no dena-turation, and some will tolerate codenaturation, in which the probe solution is
placed on the target area of the slide, a cover slip is sealed in place with rubber
cement, and then the slide is heated to between 75 and 100°C for 2–10 min.
3.2.1. Denaturation Protocol
1. Place a heated solution of 70% formamide in 2X SSC, pH 7.0 (the pH is critical;
see Note 1) in a Coplin jar as follows (see Note 3):
a. For paraffin-embedded sections, denature for 8–12 min at 85°C. The section
of placenta in the illustrated case (Fig. 1) was denatured for 10 min, although
this step is not in the manufacturer’s directions.
b. For touch preparations, smears, and chromosome spreads, denature for 2 min
at 72°C. The touch preparation of neuroblastoma in the illustrated case
(Fig. 2) was denatured for 2 min.
2. Immediately place the slide in cold (–20°C) 70% ethanol to stop the denatur-ation. After 2 min (see Note 4), transfer to 90, 95, and 100% ethanol for 2 min
each, and then allow to air-dry. Serial dehydration is crucial for paraffin sections,
but immersion in 70% ethanol followed by absolute ethanol is adequate for thin
smears, chromosome preparations, and so on.
3.3. Hybridization
Some probes require dilution into hybridization buffer. This buffer gener-ally contains 70% formamide, carrier DNA, buffers, and dextran sulfate (to
keep the solution in place on the slide). The last ingredient makes it difficult to
88 Sheldon
pipet when cold. Bring the stock solution of probe to room temperature, briefly
vortex, and centrifuge in a microfuge for 2 to 3 s to collect the contents at the
bottom of the tube. If the probe is labeled with a fluorochrome, it may be useful
to work in subdued light at this point (see Note 5). If the probe has passed its
expiration date, altering the dilution is often useful (see Note 6).
Other probes require predenaturation and/or preannealing; the manufacturer
will state this in the package insert. An aliquot is removed to a microcentrifuge
tube and heated to 72°C for 5–10 min. It is then placed on ice for a few minutes
(for alpha satellites, to prevent reannealing) or at 37°C for 30 min (for painting
probes, to allow the repeated sequences to preanneal and prevent their nonspe-cific binding to all chromosomes).
The alpha satellite probe used on the section of placenta is usually denatured
by a codenaturation step, as per the manufacturer’s instructions. Codenaturation
is a procedure by which the probe is placed on the slide, and it and the target
DNA are denatured simultaneously and then allowed to hybridize. This tech-nique must be used with probes supplied affixed to cover slips (Cytocell). Notes
5–7 are applicable.
1. Alpha satellite probe: Place 10 µL of hybridization buffer on the section, apply
the cover slip with the attached probe on the slide, and seal with rubber cement.
Allow to dry for 5 min.
2. Place on a 75°C hot plate for 5 min to denature. Proceed to step 4.
3. N-myc probe: Place 10 µL of N-myc probe on the surface of the slide or smear
and cover with a 22-mm2 cover slip. Seal with rubber cement. This probe does
not require denaturation or dilution.
4. Place the slides in the hybridization chamber, and incubate at 37°C for 30 min to
overnight. For paraffin sections and new probes, it is useful to start with the
longer incubation and decrease the time on subsequent trials. In these examples,
the probe is incubated for 16 h.
3.4. Posthybridization Wash
1. Remove the rubber cement from the cover slip; often the cover slip comes off
too. If it does not, swish the slide in a beaker of PBD to float the cover slip off.
2. Immediately place the slide in one of the following solutions (for both N-myc and
the alpha satellite used in this example, the 50% formamide wash was used):
a. 50% Formamide at 37°C, 15 min (paraffin sections, irrespective of the type of
probe).
b. 65% Formamide at 43°C, 15 min (alpha and beta satellite probes).
c. 0.4X SSC at 72°C, 2 min, for directly labeled probes; this is the rapid wash
technique (see Note 8).
3. Place the slides in 2X SSC at 37°C, 5 min; for the rapid SSC wash, follow with
30 s in PBD. Hybridized slides can be held in PBD at 4°C overnight prior to the
next step.
Fluorescent In Situ Hybridization 89
3.5. Localization of Probe
1. Directly conjugated probes require only a rinse with PBD and mounting with a
glass cover slip (see step 5).
2. Rinse the slides with PBD to remove the SSC. At this point, the placenta section
is counterstained (see step 5).
3. Apply 15–20  µL of antidigoxigenin/fluorochrome conjugate or avidin/fluoro-chrome conjugate and cover with a plastic cover slip (see Note 9), and incubate
at 37°C for 15–30 min. The N-myc probe required a fluorescein-conjugated avi-din incubation for 30 min, but is now available directly conjugated.
4. Working in subdued light, remove the cover slip and rinse with PBD three times
for 2 min each.
5. Apply counterstain such as propidium iodide for fluorescein-labeled probes or
DAPI for probes labeled with a red fluorochrome, in “antifade.” Cover with a
glass cover slip (22 × 40 or 24 × 50 mm).
3.6. Visualization of Probe
A 100-W, mercury vapor light source is recommended, particularly for
unique sequence probes (see Note 11). Locate the sections or cells under low
power; air bubbles are in the same focal plane as the cells. The presence of
label is scored under high power with either a ×100 or ×60 oil objective. For
some larger probes, such as some alpha satellites a ×40 or ×60 water objective
may be adequate. Score a minimum of 250–500 nuclei for interphase karyo-type preparations. This is particularly necessary for tissue sections, because
the entire nucleus may not be in the section. Cells to be scored for  N-myc
amplification should have at least two signals present in the nucleus, one for
each gene on chromosome 2. Slides may be stored either before or after view-ing (see Note 11). If results are unsatisfactory, many specimens can success-fully be rehybridized (see Note 12).
4. Notes
1. Formamide becomes basic as it degrades; adjust the pH of the solution with HCl.
Deionizing the formamide and freezing it in polypropylene tubes will increase
the stability. Ultrapure formamide (BRL, Bethesda, MD) stored frozen will main-tain its pH for years.
2. To determine whether the digestion is adequate, cover slip the wet slide and view
with a fluorescent microscope using a fluorescein isothiocyanate filter. If there is
green fluorescence, more digestion time is needed. If distinct nuclei are not seen,
the tissue is overdigested.
3. For specimens to be denatured in formamide, it is useful to prewarm the slides on
a slide warmer prior to placing them in the formamide. Denature no more than
four slides at a time to maintain the temperature of the formamide, and recheck
the temperature between batches of slides.
90 Sheldon
4. For most specimens, there is a trade-off between denaturation sufficient to allow
the probe to bind and overdenaturation so that morphology is lost. In general, do
not exceed 2 min unless the specimen is paraffin embedded. Older specimens can
be denatured for 2.5–3 min.
5. For most commercial probes, subdued light is more than adequate. When working
with only a few slides, and the fluorochrome is not directly conjugated, the rela-tively brief exposure to room light does not appear to cause loss of fluorescent
signal.
6. Probes that are 6 mo to 1 yr or more past their stated expiration date have been
used successfully by using a higher ratio of probe to diluent (twice the recom-mended amount of probe works well). If the probe does not require dilution, a
larger volume (20 rather than 10 µL for a 22-mm2 area) is often successful.
7. At least one vendor (Cytocell) supplies its probes bound to cover slips. These can
be used quite successfully. After placing the cover slip over the hybridization
solution on the target area of the slide and sealing with rubber cement, it is
important to wait at least 5 min before doing the denaturation step, because the
probe requires this time to dissolve off the cover slip.
8. A variety of posthybridization washes are described. For probes that are conju-gated to a fluorochrome, a “quick wash” is preferable.
9. Some manufacturers of fluorochrome conjugates sell kits that include a variety
of “blocking reagents” designed to reduce nonspecific binding of avidin to the
tissue, buffers for washing, and polypropylene cover slips for use during incuba-tion of the conjugate. “Plastic” cover slips, which reduce scratching of the speci-men during this incubation, can be cut from Parafilm. A 22 × 50 mm surface area
is preferable for this step.
10. Most commercial probes are designed for viewing with a 100- rather than 50-W
mercury vapor lamp or equivalent. Signals may not be visible at 50 W. A 100-W
bulb cannot be put in a 50-W lamp socket.
11. Slides can be stored in the dark in a refrigerator for 3–7 d. Some slides may be
usable for several months if stored in a freezer. In either case, wipe the oil off the
cover slip before storing, because it may be necessary to remount the cover slip
prior to further viewing.
12. Most specimens can be rehybridized at least once. Remove the cover slip, rinse
in PBD, and dehydrate. The existing probe will be removed during denaturation.
References
1. Pardue, M. and Gall, J. (1970) Chromosomal localization of mouse satellite DNA.
Science 168, 1356–1358.
2. Lichter, P. and Ward, D. (1990) Is non-isotopic in situ hybridization finally com-ing of age? Nature 354, 93,94.
3. Pinkel, D., Landegent, J., Collins, C., Fuscoe, J., Seagraves, R., Lucas, J., and
Gray, J. (1988) Fluorescence in situ hybridization with human chromosome spe-cific libraries. Proc. Natl. Acad. Sci. USA 85, 9138–9142.
Fluorescent In Situ Hybridization 91
4. Van Dilla, M., Deaven, L., Albright, K., et al. (1986). Human chromosome-specific DNA libraries: construction and availability. Biotechnology 4, 537–552.
5. Trask, B. (1991) Fluorescence in situ hybridization: applications in cytogenetics
and gene mapping. Trends Genet. 7, 149–154.
6. Nelson, D., Ledbetter, S., Corbo, L., et al. (1989) Alu polymerase chain reaction:
a method for rapid isolation of human-specific sequences from complex DNA
sources. Proc. Natl. Acad. Sci. USA 86, 6686–6690.
7. Ledbetter, S., Nelson, D., Warren, S., and Ledbetter, D. (1990) Rapid isolation
of DNA probes within specific chromosome regions by interspersed repeated
sequence polymerase chain reaction. Genomics 6, 475–481.
8. Mark, H. F., Jenkins, R., and Miller, W. (1997) Current applications of molecular
cytogenetic technologies. Ann. Clin. Lab. Sci. 27, 47–56.
9. Wolman, S. (1995) Applications of fluorescent in situ hybridization (FISH) to
genetic analysis of human tumors. Pathol. Annu. 30(pt. 2), 227–243.
10. Wolfe, K. and Herrington, C. (1997) Interphase cytogenetics and pathology: a
tool for diagnosis and research. J. Pathol. 181, 359–361.
11. Poddighe, P., Ramaekers, F., and Hopman, A. (1992) Interphase cytogenetics of
tumors. J. Pathol. 166, 215–224.
12. Hopman, A., van Hooren, E., van de Kaa, C., Vooijs, P., and Ramaekers, F. (1991)
Detection of numerical chromosome aberrations using in situ hybridization
in paraffin sections of routinely processed bladder cancers.  Mod. Pathol. 4,
503–513.
13. van de Kaa, C., Nelson, K., Ramaekers, F., Vooijs, P., and Hopman, A. (1991)
Interphase cytogenetics in paraffin sections of routinely processed hydatidiform
moles and hydropic abortions. J. Pathol. 165, 281–287.
14. Long, A., Mueller, J., Schwartz, J., Barrett, K., Schwartz, R., and Wolfe, H. (1992)
High specificity in situ hybridization. Diagn. Mol. Pathol. 1, 45–57.
15. Brodeur, G., Seeger, R. C., Schwab, M., et al. (1984) Amplification of N-myc in
untreated human neuroblastomas correlates with advanced disease stage. Science
224, 1121–1124.
16. Seeger, R., Brodeur, G., Sather, H., et al. (1985) Association of multiple copies of
the N-myc oncogene with rapid progression of neuroblastomas. New Engl. J. Med.
313, 1111–1116.
17. Shiloh, Y., Shipley, J., Brodeur, G., et al. (1985) Differential amplification,
assembly, and relocation of multiple DNA sequences in human neuroblastomas
and neuroblastoma cell lines. Proc. Natl. Acad. Sci. USA 82, 3761–3765.
HER-2/ neu Oncogene Amplification 93
10
HER-2/neu Oncogene Amplification
Determined by Fluorescence In Situ Hybridization
Jeffrey S. Ross, Christine E. Sheehan, and Jonathan A. Fletcher
1. Introduction
1.1. Background and Clinical Significance
The proto-oncogene HER-2/neu (C-erbB-2) has been localized to chromo-some 17q and encodes a transmembrane tyrosine kinase growth factor recep-tor. The name for the HER-2 protein is derived from human epidermal growth
factor receptor (EGFR) because it features substantial homology with the
EGFR (1 ,2) . HER-2/neu gene amplification has been associated with the
development of breast cancer in animal models (1) . The HER-2/neu protein is
a component of a four-member family of closely related growth factor recep-tors including EGFR or HER-1 (erb-B1), HER-2 (erb-B2), HER-3 (erb-B3),
and HER-4 (erb-B4) (3) . In addition to its association with disease outcome in
gastrointestinal, pulmonary, genitourinary, and other neoplasms, amplification
of the HER-2/neu gene or overexpression of the HER-2/neu protein has been
identified in from 10 to 34% of breast cancers (4–50) . The techniques used to
evaluate HER-2/neu status in breast cancer have included gene-based assays
such as Southern and slot blotting, polymerase chain reaction methods, and,
more recently,  in situ hybridization featuring both fluorescent and non-fluorescent techniques (4–50) .
Given that Southern and slot-blotting procedures are expensive, and time-consuming, and require fresh or frozen tissue, the fluorescence in situ hybrid-ization (FISH) technique was implemented to measure HER-2/neu gene copy
number on formalin-fixed, archival specimens (Fig. 1). In two previous stud-ies, the FISH method was found to be more sensitive than Southern blotting for
the detection of HER-2/neu gene amplification (48 ,51) . The FISH technique
93
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
94 Ross, Sheehan, and Fletcher
has outperformed a number of solid matrix blotting techniques designed to
detect HER-2/neu DNA and RNA as well as immunohistochemistry designed
to detect HER-2/neu protein in formalin-fixed, paraffin-embedded tissues (48) .
The FISH technique has been described as a rapid, reproducible, and extremely
reliable method of detecting HER-2/neu gene amplification (48) . In addition,
FISH can readily be performed on archived paraffin blocks stored for long
periods and has been successfully applied to fine-needle aspiration biopsies
(52). In a study using the FISH method, Xing et al. (43) reported that HER-2/neu
gene amplification was more predictive than lymph node status and was the
strongest independent predictor of outcome in breast cancer. In a recently pub-lished study, Press et al. (48) successfully utilized FISH on archived specimens
to predict short- and long-term outcome in node-negative breast cancer. This
study employed a two-category system of HER-2/neu amplification status: four
or fewer signals per nucleus = unamplified and greater than four signals per
nucleus = amplified. A recent study derived from a subset of the cases reported
Fig. 1. Four-part photomicrograph of HER-2/neu gene amplification detection by
FISH. (Upper left) Unamplified case with a mean signal count of 1.5 signals per
nucleus; (upper right) breast cancer with a borderline result featuring a mean signal
count of 4.1 signals per nucleus; (lower left) significantly amplified breast tumor with
a mean signal count of 17.5 signals per nucleus. (Lower right) Another example of a
significantly amplified breast cancer with a mean HER-2/neu signal count of 24.9
signals per nucleus.
HER-2/ neu Oncogene Amplification 95
by Press et al. (48) features a three-tiered amplification scoring system with an
equivocal borderline amplified group featuring greater than 3 but less than 10
signals per nucleus (49) . The results of the three-tiered system are similar to
those of the two-tiered system and confirm the significant association of HER-2/neu gene amplification with early recurrence, recurrence at any time, and
disease-related death in node-negative breast cancer initially treated by sur-gery alone. Both the two-tiered and the three-tiered amplification scoring sys-tems similarly found the adverse impact of HER-2/neu gene amplification to
be independent of tumor size, grade, and estrogen receptor status (48 ,49) . In
another study using a two-tiered system, HER-2/neu gene amplification again
predicted disease-related death independent of the original nodal status in a
combined series of node-negative and node-positive patients (50) .
1.2. Comparison of Methods of Detection
of HER-2/ neu Abnormalities
Immunohistochemistry has been the predominant method utilized to mea-sure HER-2/neu protein abnormalities in breast cancer. However, significant
issues can have an impact on immunohistochemistry, especially when per-formed on archival fixed paraffin-embedded tissues. Many laboratories per-form the staining on referred specimens and cannot control the time and nature
of tissue fixation, the method of tissue processing, and the temperature of the
paraffin-embedding procedure, all of which can influence HER-2/neu protein
antigen loss. Prolonged storage can also be a problem, and significant loss of
tumor-marker immunostaining intensity has been identified, particularly when
specimens are stored as unstained slides (53) . The impact of the fixative has
been considered and shown to have a significant impact on HER-2/neu
immunostaining (54) . Using cell-line controls, different antibodies have dif-fering staining patterns depending on how the cells were fixed (54) . Studies of
the various commercially available antibodies have also demonstrated a wide
variety of sensitivity and specificity for fixed paraffin-embedded tissues. In a
study by Busmanis et al. (55) , a panel of six antibodies showed a wide varia-tion in staining patterns including occasional cytoplasmic immunoreactivity (a
pattern considered to be nonspecific by most investigators). In a study of a
large panel of antibodies, Press et al. (56) similarly reported a wide range of
detection rates using a large tissue block containing multiple breast tumors.
The use of nonstandardized antigen retrieval (amplification) techniques fur-ther compounds the problem and introduces the potential for false-positive
staining. The lack of an agreed-upon scoring system for interpreting HER-2/neu
protein immunohistochemistry is another significant issue. Recent attempts to
reach consensus on the interpretation of immunohistochemical staining show
promise for dealing with this issue. Note that immunohistochemistry on frozen
96 Ross, Sheehan, and Fletcher
sections has shown substantial correlation with HER-2/neu gene–based
assays (48) and would be an ideal method of detection if it were not obviously
limited by the general lack of availability of fresh or frozen material for this
approach.
The enzyme-linked immunosorbent assay (ELISA) technique when per-formed on tumor cytosols made from fresh tissue samples avoids the potential
antigen damage associated with fixation, embedding, and uncontrolled stor-age. In three published studies including 315 patients, ELISA-based measure-ments of HER-2/neu protein in tumor cytosols have uniformly correlated with
disease outcome (39 ,41 ,43) . However, the small size of breast cancers diag-nosed in an era of enhanced screening generally precludes tumor tissue ELISA
methods because insufficient tumor tissue is available to produce a cytosol.
Western blotting can also detect HER-2/neu protein overexpression in both
tumor cytosols and archival tissues, but is generally cumbersome and imprac-tical for routine specimens. Southern and slot blotting were the first gene-based
HER-2/neu detection methods used in breast cancer specimens. These meth-ods can be significantly hampered when tumor cell DNA extracted from the
primary carcinoma sample is diluted by DNA from benign breast tissue and
inflammatory cells. The FISH technique allows simultaneous morphologic
assessment, such that evaluation of gene amplification can be restricted to
invasive carcinoma cells. This approach has been proven to be more sensitive
than Southern analysis for the detection of HER-2/neu abnormalities in breast
cancer (48) . Of the 15 published gene-based studies of HER-2/neu in breast
cancer, the 2 noncorrelating studies used Southern (50 patients) and slot-blotting (362 patients) methods. As already mentioned, the FISH-based assays
of HER-2/neu gene amplification have uniformly predicted an adverse disease
outcome.
In summary, both HER-2/neu gene amplification and protein overexpression
have been associated with an adverse outcome in breast cancer with gene-based
methods and protein detection on fresh or frozen samples obtaining the most
consistent results. Many of the published studies have used a relatively short
clinical follow-up period for a disease prone to late recurrences. Thus, abnor-malities of HER-2/neu may actually identify patients at greater risk for early
disease relapse, and evidence confirming the ability of the marker to predict
overall relapse rates must await continuing long-term studies of tumor
outcome.
The following technique for the determination of amplification of the
HER-2/neu gene in breast cancer by FISH is based on the Oncor® INFORM™
HER-2/neu Gene Detection System using a unique sequence biotinylated probe
(Oncor, Gaithersburg, MD)  (26) . Many of the materials are included in this
system.
HER-2/ neu Oncogene Amplification 97
2. Materials
2.1. Buffers and Solutions
1. Pretreatment powder.
2. Protein digesting enzyme (proteinase K).
3. 20X saline sodium citrate (SSC): 3 M NaCl, 0.3 M sodium citrate.
4. 10X phosphate-buffered detergent (PBD).
5. Biotinylated HER-2/neu DNA probe.
6. Detection reagent (fluorescein-labeled avidin).
7. 4v,6-Diamidino-2-phenylindole (DAPI)/antifade counterstain.
8. Antifade.
9. Plastic cover slips.
10. Distilled deionized water.
11. Sterile deionized water.
12. Ethanol: 70, 80, 90, and 100%.
13. Xylene.
14. 6 N HCl.
2.2. Laboratory Supplies
1. Silanized slides.
2. Glass Coplin jars.
3. Glass cover slips.
4. Micropipettor tips.
5. Microcentrifuge tubes.
6. Microcentrifuge tube rack and float.
7. Graduated cylinders.
8. 50-mL Polypropylene tubes.
2.3. Equipment
1. Fluorescence microscope equipped with the following:
a. 100-W Mercury arc light source.
b. DAPI/fluorescein isothiocyanate (FITC)/Texas Red triple band-pass filter set.
c. DAPI filter set.
d. FITC/Texas Red dual band-pass filter set.
e. ×10 dry, ×40 dry, and ×100 oil fluorescence objectives.
f. Nonfluorescing immersion oil.
2. Humidified chambers.
3. Incubator at 37 ± 2°C.
4. Oven at 65 ± 5°C.
5. Water baths and ice baths.
6. Calibrated thermometers.
7. Adjustable micropipettors.
8. Vortex.
9. pH meter.
10. Balance.
11. Microcentrifuge.
98 Ross, Sheehan, and Fletcher
2.4. Preparation of Reagent
1. 2X SSC: Add 180 mL of 20X SSC to 1620 mL of deionized water. Adjust the pH
of the 2X SSC to 7.0 with 6 N HCI. The reagent may be prepared in advance and
stored in a glass or plastic vessel at 18 to 25°C until the expiration date of the
20X SSC. Check and adjust the pH to 7.0 before use.
2. Protein-digesting enzyme stock solution (25 mg/mL): Add 4 mL of sterile dis-tilled water to 100 mg of lyophilized protein-digesting enzyme. Aliquot 400 mL
each into microcentrifuge tubes. Store this stock solution at –20°C until expira-tion of powder.
3. Protein-digesting enzyme working solution (0.25 mg/mL): Add 400 mL of
protein-digesting enzyme stock solution to 40 mL 2X SSC. This solution must be
used the same day as it is diluted and heated.
4. 1X PBD: Add 200 mL of thoroughly mixed 10X PBD to 1800 mL of distilled
water. Store at 4°C until the expiration date of the stock.
3. Methods
3.1. Preparation of Specimen Slides
1. Cut 4-µm sections and apply to silanized or positively charged slides.
2. Allow to air-dry. Bake at 65 ± 5°C overnight.
3. Deparaffinize in xylene.
4. Wash in 100% ethanol two times for 2 min each.
5. Air-dry.
3.2. Pretreatment
1. Immerse a maximum of four slides in a Coplin jar containing 40 mL of
prewarmed 30% pretreatment solution in a 43 ± 2°C water bath for 15 min.
2. Wash twice in 40 mL of 2X SSC at room temperature for 1 min.
3. Dehydrate in 70, 80, 90, and 100% ethanol for 2 min each.
4. Air-dry.
3.3. Protein Digestion
1. Immerse 40 mL of prewarmed protein digesting enzyme working solution in a
37°C water bath for 10 min.
2. Wash twice in 40 mL of 2X SSC at room temperature for 1 min.
3. Dehydrate in 70, 80, 90, and 100% ethanol for 2 min each.
4. Air-dry.
3.4. Denaturation and Hybridization of Probe
1. Prewarm the HER-2/neu probe for 5 min at 37°C.
2. Apply 10 µL of probe to each denatured tissue section and cover with a 25 × 25 mm
cover slip.
3. Hybridize at 37°C in a humidified chamber for 12–16 h.
HER-2/ neu Oncogene Amplification 99
3.5. Posthybridization Wash
1. Carefully remove the cover slip.
2. Immerse in 2X SSC at 72°C for 5 min (see Note 1).
3. Transfer to 40 mL of 1X PBD at room temperature.
3.6. Detection
1. Remove the slide from the 1X PBD and drain excess fluid without drying the
section.
2. Add 60 µL of detection reagent. Cover with a plastic cover slip.
3. Perform fluorescence detection and counterstaining with fluorescein-labeled avi-din, antiavidin antibody, and DAPI/antifade as described in the Oncor INFORM
HER-2/neu Gene Detection System package insert.
4. Wash three times in 40 mL of 1X PBD at room temperature for 2 min each.
3.7. Nuclear Counterstaining and Storage
1. Remove the slide from the PBD and drain excess fluid without drying the
section.
2. Add 20 µL of DAPI/antifade to each slide.
3. Cover with a glass cover slip.
4. Store in the dark at –15 to –25°C for up to 5 d before scoring.
3.8. Scoring
1. Use the DAPI filter set and ×10 or ×40 objective to confirm that the tissue section
contains areas of invasive breast carcinoma.
2. Use the DAPI/FITC/Texas Red triple band-pass filter set and ×100 oil immersion
objective to confirm the FITC signal is present (see Note 2).
3. Record the number of signals over 20 nonoverlapping tumor cell nuclei from
each of 2 noncontinguous fields.
4. Calculate the mean number of signals per tumor cell nucleus.
3.9. Reporting Results
If the mean signal count is  )4, report as unamplified (see Note 3). If the
mean signal count is >4, report as amplified.
4. Notes
1. Place a clean thermometer in the solution for accurate temperature monitoring.
The temperature will drop approx 1°C for each slide placed in the solution. There-fore, adjust the initial temperature accordingly and do not remove the slides from
37°C hybridization until ready to place immediately into 2X SSC, to prevent
slides from cooling down and resulting in greater temperature drop.
2. Use the DAPI filter set to select non-overlapping nuclei. Slide to dual or triple
band pass to count the signals. Background signals are generally smaller and
100 Ross, Sheehan, and Fletcher
finer (“dustlike”) whereas actual signals are slightly larger and brighter, as well
as on a different plane of focus. To begin signal counting, focus very slowly
through the background plane until the nuclear border and signal come into
focus, begin counting, and count all the signals as you focus through the nucleus.
Some cells with no signal are expected in cut sections. Record more than 20
signals as 20+, but use 20 in the calculation of the mean.
3. Although the test is reported as “not amplified” if the mean signal per nucleus
score is 4.0 or less and “amplified” if the mean score is greater than 4.0, it is
customary to add a comment when the mean score is between 3.5 and 4.5 signals
per nucleus. This comment may include a statement such as “Borderline result.
Please interpret in concert with other breast cancer prognostic factors such as
tumor size, tumor grade, and hormone receptor status.” For lymph node–negative
breast cancer, it is anticipated that approx 5–8% of the tumors will fall into the
“borderline” 3.5–4.5 signals per nucleus range. Of the lymph node–negative
tumors with mean signal counts outside the borderline range, approx 18–28%
will be amplified and 72–82% will be unamplified.
References
1. Slamon, D. J. and Clark, G. M. (1988) Amplification of C-ERB-B2 and aggres-sive breast tumors? Science 240, 1795–1798.
2. De Potter, C. R. (1994) The neu oncogene: more than a prognostic indicator?
Hum. Pathol. 25, 1264–1268.
3. Lupu, R., Cardillo, M., Harris, L., et al. (1995) Interaction between ERB-receptors
and heregulin in breast cancer tumor progression and drug resistance. Semin. Can-cer Biol. 6, 135–145.
4. Slamon, D. J., Clark, G. M., Wong, S. G., et al. (1987) Human breast cancer:
correlation of relapse and survival with amplification of the Her-2/neu oncogene.
Science 235, 177–182.
5. Berger, M. S., Locher, G. W., Saurer, S., et al. (1988) Correlation of c-erb B2
gene amplification and protein expression in human breast carcinoma with nodal
status and nuclear grading. Cancer Res. 48, 1238–1243.
6. van de Vivjer, M. J., Peterse, J. L., Mooi, W. J., et al. (1988)  Neu-protein
overexpression in breast cancer. N. Engl. J. Med. 319, 1239–1245.
7. Heintz, N. H., Leslie, K. O., Rogers, L. A., and Howard, P. L. (1990) Amplifica-tion of the c-erb B-2 oncogene in prognosis of breast adenocarcinoma.  Arch.
Pathol. Lab. Med. 114, 160–163.
8. Tsuda, H., Hirohashi, S., Shimosato, Y., et al. (1990) Correlation between histo-logic grade of malignancy and copy number of  c-erbB-2 gene in breast carci-noma: a retrospective analysis of 176 cases. Cancer 65, 1794–1800.
9. Borg, A., Tandon, A. K., Sigurdsson, H., et al. (1990) HER-2/neu amplification
predicts poor survival in node-positive breast cancer. Cancer Res. 50, 4332–4337.
10. Paik, S., Hazan, R., Fisher, E. R., et al. (1990) Pathologic findings from the
National Surgical Adjuvant Breast and Bowel Project: prognostic significance of
erb B2 protein overexpression in primary breast cancer. J. Clin. Oncol. 8, 103–112.
HER-2/ neu Oncogene Amplification 101
11. Battifora, H., Gaffey, M., Esteban, J., et al. (1991) Immunohistochemical assay of
neu/c-erb B-2 oncogene product in paraffin-embedded tissues in early breast cancer:
retrospective follow-up study of 245 stage I and II cases. Mod. Pathol. 4, 466–474.
12. Kallioniemi, O. P., Holli, K., Visakorpi, T., et al. (1991) Association of C-erb B2
protein over-expression with high rate of cell proliferation, increased risk of vis-ceral metastasis and poor long-term survival in breast cancer. Int. J. Cancer 49,
650–655.
13. Clark, G. M. and McGuire, W. L. (1991) Follow-up study of HER-2/neu amplifi-cation in primary breast cancer. Cancer Res. 51, 944–948.
14. Lovekin, C., Ellis, I. O., Locker, A., et al. (1991) C-erb B2 oncoprotein expres-sion in primary and advanced breast cancer. Br. J. Cancer 63, 439–443.
15. McCann, A. H., DeDervan, T. A., O’Regan, M., et al. (1991) Prognostic signifi-cance of C-erb B2 and estrogen receptor status in human breast cancer. Cancer
Res. 51, 3296–3303.
16. Dykens, R., Corbett, I. P., Henry, J., et al. (1991) Long term survival in breast
cancer related to overexpression of the C-erb B2 oncoprotein: an immunohisto-chemical study using monoclonal antibody NCL-CB11. J. Pathol. 163, 105–110.
17. Rilke, F., Colnaghi, M. I., Cascinelli, N., et al. Prognostic significance of HER-2/
neu expression in breast cancer and its relationship to other prognostic factors.
Int. J. Cancer 49, 44–49.
18. Winstanley, J., Cooke, T., Murray, G. D., et al. (1991) The long term prognostic
significance of C-erb B2 in primary breast cancer. Br. J. Cancer 63, 447–450.
19. O’Reilly, S. M., Barnes, D. M., Camplejohn, R. S., et al. (1991) The relationship
between C-erb B2 expression, and s-phase fraction in prognosis in breast cancer.
Br. J. Cancer 63, 444–446.
20. Paterson, M. C., Dietrich, K. D., Danyluk, J., et al. (1991) Correlation between C-erb B2 amplification and risk of recurrent disease in node-negative breast cancer.
Cancer Res. 51, 556–567.
21. Toikkanen, S., Helin, H., Isola, J., et al. (1992) Prognostic significance of Her-2
oncoprotein expression in breast cancer: a 30-year follow up. J. Clin. Oncol. 10,
1044–1048.
22. Molina, R., Ciocca, D. R., Candon, A. K., et al. (1992) Expression of HER-2/neu
oncoprotein in breast cancer: a comparison of immunohistochemical and western
blot techniques. Anticancer Res. 12, 1965–1991.
23. Noguchi, M., Koyasaki, M., Ohta, N., et al. (1992) c-erb B-2 oncoprotein expres-sion versus internal mammary lymph node metastases as additional prognostic
factors in patients with axillary lymph node-positive breast cancer.  Cancer 69,
2953–2960.
24. Allred, D. C., Clark, G. M., Tandon, A. K., et al. (1992) HER-2/neu node-negative breast cancer: prognostic significance of overexpression influenced by
the presence of in-situ carcinoma. J. Clin. Oncol. 10, 599–605.
25. Babiak, J., Hugh, J., and Poppeme, S. (1992) Significance of c-erb B-2 amplifica-tion in DNA aneuploidy: analysis in 78 patients with node-negative breast cancer.
Cancer 70, 770–776.
102 Ross, Sheehan, and Fletcher
26. Tiwari, R. K., Borgen, P. I., Wong, G. Y., et al. (1992) HER-2/neu amplification
and overexpression in primary human breast cancer is associated with early
metastasis. Anticancer Res. 12, 419–426.
27. Gusterson, B. A., Gelber, R. D., Goldhirsch, A., et al. (1992) Prognostic impor-tance of C-erb B2 expression in breast cancer. J. Clin. Oncol. 10, 1049–1056.
28. Bianchi, S., Paglierani, M., Zampi, G., et al. (1993) Prognostic significance
of C-erb B2 expression in node negative breast cancer.  Brit. J. Cancer 67,
625–629.
29. Press, M. F., Pike, M. C., Chazin, V. R., et al. (1993) Her-2/neu expression in
node-negative breast cancer: direct tissue quantification by computerized image
analysis and association of overexpression with increased risk of recurrent dis-ease. Cancer Res. 53, 4960–4970.
30. Seshadri, R., Firgaira, F. A., Horsfall, D. J., et al. (1993) Clinical significance of
Her-2/neu oncogene amplification in primary breast cancer.  J. Clin. Oncol. 11,
1936–1942.
31. Descotes, F., Pavy, J. J., and Adessi, G. L. (1993) Human breast cancer: correla-tion study between Her-2/neu amplification and prognostic factors in an
unselected population. Anticancer Res. 13, 119–124.
32. Giai, M., Roagna, R., Ponzone, R., et al. (1994) Prognostic and predictive rel-evance of C-erb B2 and ras expression in node positive and negative breast can-cer. Anticancer Res. 14, 1441–1450.
33. Muss, H. B., Thor, A. D., Berry, D. A., et al. (1994) C-erb-B2 expression
and response to adjuvant therapy in women with node-positive early breast can-cer. N. Engl. J. Med. 330, 1260–1266.
34. Tetu, B. and Brisson, J. (1994) Prognostic significance of Her-2/neu oncogene
expression in node-positive breast cancer: the influence of the pattern of
immunostaining and adjuvant therapy. Cancer 73, 2359–2365.
35. Hartmann, L. C., Ingle, J. N., Wold, L. E., et al. (1994) Prognostic value of CerbB2
overexpression in axillary lymph node positive breast cancer: results from a ran-domized adjuvant treatment protocol. Cancer 74, 2956–2963.
36. Jacquemeier, J., Penault-Llorca, P., Viens, P., et al. (1994) Breast cancer response
to adjuvant chemotherapy in correlation with erb B2 and p53 expression. Anti-cancer Res. 14, 2773–2778.
37. Marks, J. R., Humphrey, P. A., Wu, K., et al. (1994) Overexpression of p53 and
Her-2/neu proteins as prognostic markers in early stage breast cancer. Ann. Surg.
219, 332–341.
38. Rosen, P. P., Lesser, M. L., Arroyo, C. D., et al. (1995) Immunohistochemical
detection of Her-2/neu expression in patients with axillary lymph node-negative
breast carcinoma: a study of epidemiologic risk factors, histologic features and
prognosis. Cancer 75, 1320–1326.
39. Quenel, N., Wafflart, J., Bonichon, F., et al. (1995) The prognostic value of
c-erbB2 in primary breast carcinomas: a study on 942 cases. Breast Cancer Res.
Treat. 35, 283–291.
HER-2/ neu Oncogene Amplification 103
40. Sundblad, A. S., Pellicer, E. M., and Ricci, L. (1996) Carcinoembryonic expres-sion in stages I and II breast cancer: its relationship with clinicopathologic fac-tors. Hum. Pathol. 27, 297–300.
41. O’Malley, F. P., Saad, Z., Kerkvliet, N., et al. (1996) The predictive power of
semiquantitative immunohistochemical assessment of p53 and C-erbB2 in lymph
node-negative breast cancer. Hum. Pathol. 27, 955–963.
42. Hieken, T. J., Mehta, R. R., Shilkaitis, A., et al. (1996) Her-2/neu and p53 expression
in breast cancer: valid prognostic markers when assessed by direct immunoassay,
but not by immunochemistry. Proc. Annu. Meet. Am. Soc. Clin. Oncol. 15, A113.
43. Xing, W. R., Gilchrist, K. W., Harris, C. P., et al. (1996) FISH detection of HER-2/neu oncogene amplification in early onset breast cancer.  Breast Cancer Res.
Treat. 39, 203–212.
44. Dittadi, R., Brazzale, A., Pappagallo, G., et al. (1997) ErbB2 assay in breast can-cer: possibly improved clinical information using a quantitative method. Antican-cer Res. 17, 1245–1247.
45. Fernandez Acenero, M. J., Farina Gonzalez, J., and Arangoncillo Ballesteros, P.
(1997) Immunohistochemical expression of p53 and c-erbB-2 in breast carcinoma:
relation with epidemiologic factors, histologic features and prognosis. Gen. Diagn.
Pathol. 142, 289–296.
46. Eissa, S., Khalifa, A., el-Gharib, A., et al. (1997) Multivariate analysis of DNA
ploidy, p53, c-erbB-2 proteins, EGFR, and steroid hormone receptors for short-term prognosis in breast cancer. Anticancer Res. 17, 3091–3097.
47. Charpin, C., Garcia, S., Bouvier, C., et al. (1997) c-erbB-2 oncoprotein detected
by automated quantitative immunocytochemistry in breast carcinomas correlates
with patients’ overall and disease-free survival. Br. J. Cancer 75, 1667–1673.
48. Press, M. J., Bernstein, L., Thomas, P. A., et al. (1997) Her-2/neu gene amplifica-tion characterized by fluorescence in situ hybridization: poor prognosis in node-negative breast carcinomas. J. Clin. Oncol. 15, 2894–2904.
49. Ross, J. S., Muraca, P. J., Jaffe, D., et al. (1998) Multivariate analysis of prognos-tic factors in lymph node negative breast cancer. Mod. Pathol. 11, 26A.
50. Depowski, P. L., Brien, T. P., Sheehan, C. E., et al. (1998) Prognostic significance
of p34cdc2 cyclin dependent kinase and MIB1 overexpression, and HER-2/neu
gene amplification detected by fluorescence in-situ hybridization in breast can-cer. Mod. Pathol. 11, 18A.
51. Pauletti, G., Godolphin, W., Press, M. F., et al. (1996) Detection and quantitation
of HER-2/neu gene amplification in human breast cancer archival material using
fluorescence in situ hybridization. Oncogene 13, 63–72.
52. Sauter, G., Feichter, G., Torhorst, J., et al.(1996) Fluorescence in-situ hybridiza-tion for detecting ERBB-2 amplification in breast tumor fine needle aspiration
biopsies. Acta Cytol. 40, 164–173.
53. Jacobs, T. W., Prioleau, J. E., Stillman, I. E., et al. (1996) Loss of tumor marker-immunostaining intensity on stored paraffin slides of breast cancer. J. Natl. Can-cer Inst. 88, 1054–1059.
104 Ross, Sheehan, and Fletcher
54. Penault-Llorca, F., Adelaide, J., Houvenaeghel, G., et al. (1994) Optimization of
immunohistochemical detection of ERBB2 in human breast cancer: impact of fixa-tion. J. Pathol. 173, 65–75.
55. Busmanis, I., Feleppa, F., Jones, A., et al. (1994) Analysis of C-erb B2 expression
using a panel of six commercially available antibodies. Pathology 26, 261–267.
56. Press, M. F., Hung, G., Godolphin, W., et al. (1994) Sensitivity of Her-2/neu
antibodies in archival tissue samples: potential source of error in immunohis-tochemical studies of oncogene expression. Cancer Res. 54, 2771–2777.
Nested RT-PCR Assay to Detect BCR/ abl 105
11
A Nested Reverse Transcriptase-Polymerase
Chain Reaction Assay to Detect BCR/abl
Linda M. Wasserman
1. Introduction
Chronic myelogenous leukemia (CML), a clonal myeloproliferative disor-der in adults, and some pediatric and adult acute lymphoblastic leukemias
(ALLs) are characterized by the presence of a Philadelphia chromosome,
t(9;22)(q34;q11) (1) . In this chromosomal translocation, exons from a major
breakpoint cluster region (M-bcr), located on chromosome 22q11, are joined
to the c-abl proto-oncogene, located on chromosome 9q34. When this chromo-somal translocation occurs in a hematopoietic stem cell, the resulting BCR/abl
fusion protein has increased tyrosine kinase activity and a transforming capac-ity that is critical to the pathogenesis of these leukemic disorders.
The Philadelphia chromosome can be detected in 95% of adults with CML
and 23–50% of adult ALL and 11% of childhood ALL (2) . In the M-bcr, most
translocations occur within intronic sequences between the second and fourth
exons. Most translocations in c-abl occur across a large region 5v to c-abl
exon II. Although breakpoints in the M-bcr and c-abl can be widely distributed
along their respective chromosomes, mRNA processing of the fusion transcript
consistently links either M-bcr exons b1–b3 or exons b1 and b2 to c-abl exon
II, or occasionally to c-abl exon Ia (see Fig. 1). Because of the consistent pro-cessing of the fusion transcript, reverse transcriptase-polymerase chain reac-tion (RT-PCR) assays, which use mRNA as a starting material, can readily
detect almost all Philadelphia chromosome translocations despite the variabil-ity in the location of chromosomal breakpoints.
Detection of the BCR/abl fusion transcript by RT-PCR is used clinically
either to confirm a CML or ALL diagnosis or to detect and monitor the pres-105
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
106 Wasserman
ence of minimal residual disease in leukemic patients following treatment (3) .
The sensitivity of an RT-PCR assay, particularly if it is a nested, two-step PCR,
typically enables the detection of one positive cell within a background of
105–107 normal cells. In the multiplexed, nested RT-PCR assay described
herein, RNA is extracted either from peripheral blood leukocytes or from bone
marrow hematopoietic cells. The RNA is first reverse transcribed and then sub-jected to two rounds of PCR. The final PCR product is electrophoresed through
an ethidium bromide–stained agarose gel and detected by fluorescence under
ultraviolet light.
The multiplexed PCR reactions described in this chapter contain primers
for the BCR/abl translocation (4) as well as a set of control primers for the
`-catenin gene  (5) to confirm the quality and amplifiability of the cDNAs
being tested. The positive control `-catenin primers ensure that the cDNA in
each specimen is intact and that the RNA extraction and reverse transcription
procedures are successful. A BCR/abl cDNA containing M-Bcr exons b1–b3
generates a 194-bp second-round PCR product whereas a BCR/abl cDNA con-taining only M-Bcr exons b1 and b2 generates a 119-bp second-round PCR
product. Presence of the second-round `-catenin PCR amplicon is detected by
the presence of a 372-bp product. Table 1 gives the round 1 and round 2 PCR
primer sequences for both BCR/abl and `-catenin. In the round 2 PCR, an
aliquot of the PCR product from round 1 is amplified using BCR/abl primers
that are both internal to the round 1 primers and  `- catenin primers that are
internal to the round 1 antisense primer.
2. Materials
2.1. Positive Control Cell Line
The K562 cell line is available from ATCC (cat. no. CCL 243) (http://
http://www.atcc.org) and contains a BCR/abl translocation containing M-bcr exons
b1–b3.
Fig. 1. The Philadelphia chromosome.
Nested RT-PCR Assay to Detect BCR/ abl 107
2.2. RNA Extraction (see Note 1)
1. 1.5-mL sterile plastic microcentrifuge tubes.
2. Sterilized aerosol-resistant pipet tips.
3. Purescript RNA Isolation Kit (cat. no. R-5000; Gentra Systems, Minneapolis, MN).
4. Sterile, double-deionized water treated with diethylpyrocarbonate (DEPC) and
autoclaved (see Note 2).
5. Glycogen (20 mg/mL) (cat. no. 901 393; Boehringer Mannheim).
6. 100% Isopropanol, dedicated to only RNA extraction use, stored at 4°C.
7. 70% Ethanol, dedicated to only RNA extraction use, stored at 4°C.
8. Agarose DNA Grade (cat. no. BP164-100; Fisher).
9. Ethidium bromide (10 mg/mL).
10. 1X Tris-borate EDTA (TBE) prepared with DEPC-treated water.
11. 80% Glycerol/bromophenol blue loading buffer.
2.3. Reverse Transcription
1. GeneAmp RNA PCR Core Kit (cat. no. N808-0143; Perkin Elmer).
2. 200-µL Sterile microcentrifuge tubes.
3. Sterilized aerosol-resistant pipet tips.
4. RNA extracted from the K562 cell line.
5. RNA extracted from a “normal” subject, i.e., an individual with no evidence of
leukemia, to use as a negative control.
6. RNA extracted from patient sample(s).
2.4. PCR Rounds 1 and 2
1. GeneAmp RNA PCR Core Kit (Perkin Elmer), which includes dNTPs, Taq poly-merase, PCR Buffer II, and 25 mM MgCl2).
2. 200-µL Sterile microcentrifuge tubes.
Table 1
BCR/abl and `-Catenin Primer Sequences
Sequence Genset code no.
BCR/abl
Oligo A: 5v-ggA gCT gCA gAT gCT gAC CAA C-3v Genset HG099
Oligo B: 5v-CTg Agg CTC AAA gTC AgA Tg-3v Genset HG101
Oligo C: 5v-gCT TCT CCC TgA CAT CCg Tg-3v Genset HG098
Oligo D: 5v-CgA gCg gCT TCA CTC AgA CC-3v Genset HG100
`-Catenin:
Sense 4: 5v-TTC CAC gAC TAg TTC AgT TgC-3v
Antisense 4: 5v-CTA CAg gCC AAT CAC AAT gC-3v
Antisense 3: 5v-AAC AgC AgC TgC ATA TgT Cg-3v
108 Wasserman
3. Sterilized aerosol-resistant pipet tips.
4. Patient and control cDNAs.
5. Sterile double-deionized water.
2.5. Gel Electrophoresis
1. NuSieve 3 1 Agarose (cat. no. 50090; FMC BioProducts).
2. 1X TBE.
3. Ethidium bromide (10 mg/mL).
4. 80% Glycerol–bromophenol blue loading buffer: 80% glycerol, 10 mM EDTA,
0.1% bromophenol blue.
5. DNA size standards: either pBR322/HaeIII (cat. no. D 9655; Sigma, St. Louis,
MO) or q X174 DNA/HaeIII (cat. no. D 0672; Sigma).
3. Methods
3.1. RNA Extraction (see Note 3)
RNA is more labile than DNA and degrades relatively quickly. Thus, RNA
should be extracted from blood or bone marrow specimens as soon as possible
after the specimen is received and accessioned in your laboratory. The speci-men should be stored at 4°C from the time it is received in the laboratory until
the RNA is extracted. The following method uses components of the Purescript
RNA Isolation Kit (Gentra Systems) and includes some modifications of the
original protocol found useful in our laboratory to increase RNA yield.
1. For best results, use at least 3 mL of peripheral blood or bone marrow. If the
patient’s specimen is ❤ mL, use the entire specimen.
2. Use pipets dedicated to nucleic acid extraction, ideally dedicated to RNA extrac-tion. Do not use these pipets when assembling the master mixes for the reverse
transcription and PCR steps.
3. For each milliliter of peripheral blood or bone marrow, add 3 mL of red blood
cell lysis solution in a Corning 15-mL centrifuge tube. Cap the tube and invert
once to mix. Incubate at room temperature for 10 min, inverting the tube once
more in the middle of the incubation.
4. Centrifuge at high speed in a bench-top centrifuge for 1 min to pellet the white
cells.
5. Discard the supernatant and vortex the pellet to loosen it from the bottom of the
tube.
6. Add 300 µL of cell lysis solution and pipet up and down no more than three times
to lyse the white cells. Place the cell lysate in a 1.5-mL microcentrifuge tube.
7. Add 175 µL of protein precipitation solution, cap the tube, and invert it gently 10
times to mix. Place the microcentrifuge tube on ice for 5 min (see Note 4).
8. Centrifuge at 12,800g for 3 min.
9. Carefully remove the supernatant with a pipet and place it in a new
microcentrifuge tube. Using the pipet, measure the approximate volume of the
supernatant.
Nested RT-PCR Assay to Detect BCR/ abl 109
10. Add an equal volume of 100% isopropanol and 1 µL of glycogen (20 mg/mL).
Cap the tube and invert gently 50 times.
11. Centrifuge at 12,800g for 5 min.
12. Carefully pour off the isopropanol and drain the tube on clean paper towels or
absorbent paper.
13. Carefully pipet 300  µL of 70% ethanol into the microcentrifuge, directing the
stream of fluid along the side of the tube rather than directly onto the pellet, in
order to avoid dislodging it. Centrifuge at highest speed for 1 min.
14. Either carefully pour off the 70% ethanol without dislodging the pellet or remove
most of it with a pipet, and allow the remainder of the ethanol to drain out of the
microcentrifuge tube onto paper towels.
15. When no drops of ethanol remain on the sides of the microcentrifuge tube, rehy-drate the pellet with 20–40 µL of RNA hydration solution (see Note 5).
3.2. Quantifying RNA Yield
There are two methods of quantifying RNA yield: spectrophotometric analysis
of the absorption at 260 and 280 nm, and visual inspection on an agarose gel.
Determining the ratio of absorption at 260280 nm is used to calculate the concen-tration (micrograms/microliter) of nucleic acid in the preparation but is subject
to positive interference from DNA. Electrophoresis of an aliquot of RNA in an
agarose gel with ethidium bromide staining demonstrates the 18S and 28S bands
of RNA, and is a visual check of the presence of high molecular weight RNA.
3.2.1. Absorption at 260 nm
Dilute 6 µL of RNA into 600 µL of DEPC-treated sterile double-deionized
water. The RNA concentration in nanograms/microliter is obtained by multiply-ing the absorption at 260 nm by 4000. The ratio of absorptions at 260 280 nm
should be *1.8.
3.2.2. Agarose Gel Analysis
1. Make a 1% agarose gel using 1X TBE prepared with DEPC-treated double-deionized water (ddH2O).
2. For a small gel, add 0.4 g of agarose to 40 mL of TBE and melt in a microwave
oven.
3. Add 4 µL of ethidium bromide (10 mg/mL) to the molten agarose prior to pour-ing the gel.
4. When the gel sets, submerge with 1X TBE and remove the comb.
5. Mix 3 µL of RNA with 3 µL of 80% glycerol/bromophenol blue loading buffer
and load into the gel.
6. Run at 80 V until the dye front is two thirds of the way down the gel.
7. Photograph the gel under UV light. 18S RNA has an electrophoretic migration
equivalent to a DNA molecule of 1.9 kb. 28S RNA migrates at the equivalent of
a 4.7-kb DNA molecule.
110 Wasserman
3.3. Storage of RNA
RNA can be either frozen at –80°C indefinitely until needed for reverse
transcription or used immediately in the reverse transcription step. If the RNA
is frozen prior to reverse transcription, it should be thawed on ice.
3.4. Reverse Transcription
1. Prepare a master mix for reverse transcription according to the kit directions,
calculating the total volume needed based on the number of patient and control
reactions needed. Volumes of ingredients per reaction are as follows: 4  µL of
25 mM MgCl2; 2 µL of 10X PCR Buffer II; 2 µL of each dNTP, at 10 mM initial
concentration; 1 µL of RNAsin (20 U/µL); 1 µL of MULV reverse transcriptase
(50 U/µL); and 1 µL of either random hexamers (50 µM) or oligo dT (50 µM).
2. Set up the reactions in a biosafety hood, if available, or in a space in your labora-tory dedicated to assembling PCR ingredients. Use pipets dedicated to setting up
PCR reactions.
3. Make duplicate reverse transcription reactions for each patient sample, one posi-tive control reaction each for the BCR/abl exons b1–b3 translocation (i.e., K562)
and the BCR/abl exons b1 and b2 translocation, if available, and one reaction
for the negative RNA control (i.e., an RNA known not to contain a BCR/abl
translocation).
4. Pipet 17  µL of master mix into each 200-µL microcentrifuge tube. Each tube
should be and remain capped after addition of the master mix and should be
opened only when pipeting in the appropriate RNA aliquot.
5. Add 3 µL of RNA to each patient sample, positive and negative control tubes,
recapping each tube. Allow the tubes to sit at room temperature for 10 min to
enhance binding of the random hexamers or oligo dT primer to the RNA.
6. Place the tubes in a thermocycler. Program the thermocycler for 1 cycle at 42°C
for 60 min followed by 1 cycle at 90°C for 5 min to inactivate the reverse
transcriptase.
7. Either use the cDNA immediately for the first PCR round or store at –20°C until
needed.
3.5. PCR Round 1
1. Prepare the first round PCR master mix with dedicated PCR reagents, pipets, and
consumable supplies in a location in your laboratory dedicated to preparing PCR
reactions. Include duplicate tubes for each patient specimen, single tubes for the
positive and negative control reactions, and a single tube for a water-alone con-trol (see Notes 6–8). Prepare the round 1 PCR master mix according to the fol-lowing reaction volumes (a total of 40.0 µL):
a. 23.5 µL of sterile double deionized water.
b. 3.0 µL of 25 mM MgCl2 (1.5 mM final concentration).
c. 5.0 µL of 10X PCR Buffer II (1X final concentration).
d. 1.0 µL of 10 mM dATP (200 µM final concentration).
Nested RT-PCR Assay to Detect BCR/ abl 111
e. 1.0 µL of 10 mM dGTP (200 µM final concentration).
f. 1.0 µL of 10 mM dCTP (200 µM final concentration).
g. 1.0 µL of 10 mM dTTP (200 µM final concentration).
h. 1.0 µL of 10 mM `-catenin sense 4 primer (200 nM final concentration).
i. 1.0 µL of 10 mM `-catenin antisense 4 primer (200 nM final concentration)
j. 1.0 µL of 25 mM Oligo C (500 nM final concentration).
k. 1.0 µL of 25 mM Oligo D (500 nM final concentration).
l. 0.5 µL of Taq (5 U/µL) (2.5 U per reaction).
2. Aliquot 40  µL of master mix into each reaction tube. Cap the tubes and carry
them to the area in your laboratory dedicated to the addition of DNA, cDNA, or
PCR products to PCR reactions.
3. Using pipets dedicated to the addition of DNA, cDNA, or PCR reaction products
and aerosol-resistant pipet tips, add 10 µL of the appropriate cDNA to each PCR
tube or 10  µL of sterile double-deionized water to the water-alone control,
uncapping each tube only to add the appropriate cDNA or sterile water and then
recapping each tube prior to uncapping the next tube.
4. Vortex or pulse spin the tubes and place them in the thermocycler.
5. Use the following PCR cycle conditions:
a. Step 1: 94°C for 10 min for 1 cycle.
b. Step 2: 94°C for 1 min, 50°C for 1 min, 72°C for 1 min.
c. Step 3: 72°C for 10 min for 1 cycle.
6. Repeat step 5b for 30 cycles for PCR round 1.
7. Allow the tubes to cool to room temperature. Store at 4°C until ready to assemble
PCR round 2.
3.6. PCR Round 2
PCR Round 2 is the nested PCR reaction and uses an aliquot of the PCR
round 1 product.
1. Prepare the PCR round 2 master mix in your laboratory’s PCR setup area using
dedicated reagents, pipets, and aerosol-resistant pipet tips according to the fol-lowing reaction volumes (a total of 46.0 µL):
a. 29.5 µL of Sterile double-deionized water.
b. 3.0 µL of 25 mM MgCl2 (1.5 m× final concentration)
c. 5.0 µL of 10X PCR Buffer II (1X final concentration).
d. 1.0 µL of 10 mM dATP (200 µM final concentration).
e. 1.0 µL of 10 mM dGTP (200 µM final concentration).
f. 1.0 µL of 10 mM dCTP (200 µM final concentration).
g. 1.0 µL of 10 mM dTTP (200 µM final concentration).
h. 1.0 µL of 10 µM `-catenin sense 4 primer (200 nM final concentration).
i. 1.0 µL of 10 µM `-catenin antisense 3 primer (200 nM final concentration).
j. 1.0 µL of 25 µM Oligo A (500 nM final concentration).
k. 1.0 µL of 25 mM Oligo B (500 nM final concentration).
l. 0.5 µL of Taq (5 U/µL) (2.5 U per reaction).
112 Wasserman
2. Aliquot 46 µL into each reaction tube. Cap each tube and carry to the area in your
laboratory dedicated to the addition of DNA or PCR products. Using dedicated
pipets and aerosol-resistant pipet tips, add 4 µL of PCR round 1 product to each
tube, recapping each tube immediately afterward.
3. Vortex or pulse spin each tube and insert in the thermocycler.
4. Use the same PCR cycling conditions as for round 1.
5. Allow the tubes to cool to room temperature and store at 4°C until ready for gel
electrophoresis.
3.7. Gel Electrophoresis
1. For 100 mL of molten agarose, add 9 g of NuSieve 3 1 to 100 mL of 1X TBE and
melt in a microwave oven to make a 3% gel. Add 10 µL of ethidium bromide
(10 mg/mL) and pour into a gel apparatus, using a wide-tooth comb.
2. When the gel is set, add enough 1X TBE to just cover the agarose, and carefully
remove the comb.
3. Mix 20 µL of each PCR round 2 product with 4 µL of 80% glycerol/bromophenol
blue loading buffer and load into the gel.
4. Use either pBR322/HaeIII (1 µg/µL) (Sigma) or qX174 DNA/HaeIII (1 µg/µL)
(Sigma) as DNA size markers.
4. Notes
1. All buffers and consumable plastics needed for this assay should be kept dedi-cated to work with RNA. They should be kept separate from similar laboratory
supplies used for work with DNA and should be labeled “FOR RNA USE
ONLY.”
2. All glassware used to measure reagents or to contain buffers and any sterile
double-deionized water used to work with RNA should first be treated with 0.1%
DEPC (cat. no. D 5758, Sigma) to inactivate RNases. One milliliter of DEPC is
added to each liter of sterile ddH2O, allowed to sit for at least 12 h at room tem-perature, and then autoclaved for 15 min on the liquid cycle. Glassware can be
filled to the brim with sterile ddH2O, an appropriate amount of DEPC added and
allowed to sit in the vessel for 12 h at room temperature. The DEPC-H2O is
removed from the glassware prior to autoclaving. DEPC is unstable in the pres-ence of Tris buffers and breaks down to ethanol and carbon dioxide. Thus, when
preparing the 5X TBE buffer that will be used to check the quality of the RNA on
an agarose gel prior to the reverse transcriptase step, prepare the buffer in DEPC-treated glassware, using DEPC-treated ddH2O rather than adding DEPC directly
to the 5X TBE.
3. If you have never extracted RNA before, it is wise to practice on specimens of
peripheral blood prior to extracting a patient’s sample in order to acquaint your-self with the extraction protocol you are using, to become familiar with manipu-
Nested RT-PCR Assay to Detect BCR/ abl 113
lating the RNA pellet following isopropanol precipitation and ethanol wash, and
to become familiar with rehydrating the RNA pellet following the ethanol wash.
The RNA extraction step is critical to the success of the assay because it is the
starting material on which the remainder of the assay depends. Many RNA
extraction kits are available and you may wish to try several to determine which
kit or extraction method gives the best results in your laboratory. No matter which
kit or RNA extraction protocol you use, it is wise to keep the white RNA cell
pellet on ice, to complete the extraction protocol without interruption once you
start it; and when precipitating the RNA, to use equal volumes of supernatant to
isopropanol.
4. After completing the protein-DNA precipitation step using the Purescript
reagents, the red-brown pellet, consisting of residual red cell debris, protein, and
DNA, should be tight and well compacted at the bottom of the microcentrifuge
tube. If the pellet is rather loose and not well compacted, adding additional pro-tein precipitation reagent may be beneficial. When the pellet is loose and has a
mucous-like appearance, it can be difficult to remove the supernatant, and the
volume of supernatant recovered is often reduced. Loss of supernatant reduces
the ultimate RNA yield.
5. If the RNA pellet appears small, you might wish to rehydrate it in half the recom-mended volume of rehydration solution as recommended in order to maximize
the concentration of RNA that goes into the reverse transcription step. Because
the volume of RNA in the reverse transcription step is fixed, it is advantageous to
maximize the RNA concentration being reverse transcribed, in order to detect the
BCR/abl transcript, should it be rare in your specimen.
6. Awareness of the ways in which a PCR reaction can become contaminated, as
evidenced by a PCR product seen in the water-alone control reaction, is always
required when amplifying DNA. However, when a PCR reaction is nested, such
as in this protocol, contamination can occur much more easily and must be more
vigorously anticipated and guarded against. Because an aliquot of the water-alone
control reaction is reamplified in round 2, any slight, otherwise undetectable con-tamination occurring in the round 1 step could be detected when an aliquot is
reamplified in round 2.
7. Ways to avoid contamination include the following:
a. Assembling the PCR mixes in a laboratory area dedicated to that purpose,
using dedicated pipetmen and aerosol-resistant pipet tips.
b. Capping each tube after aliquoting the PCR master mix, keeping each tube
capped except when adding each respective cDNA or PCR aliquot, and then
recapping each tube prior to uncapping the next.
c. Adding cDNA or PCR product only in a laboratory area dedicated to that
purpose, using dedicated pipetmen and aerosol-resistant pipet tips.
8. Should contamination be detected in the blank lane, it is imperative that all solu-tions or reagents, including double-deionized water, dNTPs, primer dilutions,
and Taq, be discarded and new dilutions prepared.
114 Wasserman
References
1. Melo, J. V. (1996) The molecular biology of chronic myeloid leukemia. Leukemia
10, 751–756.
2. McClure, J. S. and Litz, C. E. (1994) Chronic myelogenous leukemia: molecular
diagnostic considerations. Hum. Pathol. 25, 594–597.
3. Campana, D. and Pui, C.-H. (1995) Detection of minimal residual disease in acute
leukemia: methodological advances and clinical significance.  Blood 85,
1416–1434.
4. Genset Corp. (1991) Human Genes. Oligonucleotide Handbook, vol. 2, La Jolla, CA.
5. Oyama, T., Kanai, Y., Ochiai, A., Akimoto, S., Oda, T., Yanagihara, K.,
Nagafuchi, A., Tsukita S., Shibamoto, S., Ito, F., Takeichi, M., Matsuda, H., and
Hirohashi, S. (1994) A truncated  `-catenin disrupts the interaction between
E-cadherin and _-catenin: a cause of loss of intercellular adhesiveness in human
cancer cell lines. Cancer Res. 54, 6282–6287.
Anomalies in Acute Myeloid Leukemias 115
12
Detection of t(15;17)(q24;q21),
inv(16)/t(16;16)(p13;q22), and t(8;21)(q22;q22)
Anomalies in Acute Myeloid Leukemias
David S. Viswanatha
1. Introduction
The acute myeloid leukemias (AMLs) are a relatively heterogeneous group
of diseases. However, there is growing awareness that the clinical features and
subclassification of morphologic leukemia types is often highly correlated with
tumor genetics. Furthermore, distinct genetic subgroups of AML are associ-ated with improved therapeutic sensitivity and a more favorable clinical out-come. These observations have prompted suggestions for a revision of the
current French-American-British leukemia classification (1) , utilizing geneti-cally defined principles (2) . Three recurrent chromosomal translocations are
identified in approx 25–30% of  de novo adult AMLs. These include the
t(15;17), associated with acute promyelocytic leukemia ([APL]; AML-M3);
the inv(16) and related t(16;16), associated with AML-M4Eo; and the t(8;21),
associated most commonly with AML-M2. Each of these abnormalities results
in the formation of a chimeric leukemia-specific fusion gene, which is tran-scribed and expressed as a fusion protein. The widespread genetic deregulation
caused by such fusion proteins is thought to interfere with proliferative control
and cell differentiation mechanisms, leading to the leukemic state. The presence of
these and other fusion gene events can be specifically and sensitively detected by
reverse transcriptase-polymerase chain reaction (RT-PCR) analysis.
The t(15;17) anomaly is associated with fusion of the  PML (15q24) and
RAR_(17q21) genes (3) . The PML gene product is localized to specific nuclear
body complexes and may function as a negative transcriptional regulator (4–6).
In addition,  PML expression is required for myeloid cell differentiation
induced via the retinoic acid pathway and further exhibits antiproliferative and
antitumor effects (7) . Recent evidence also suggests a role for PML in cellular
115
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
116 Viswanatha
apoptosis (8 ,9) and perhaps in immunomodulation by interactions with the
major histocompatibility system  (10) . The  RAR_ gene encodes a nuclear
receptor for retinoic acid; members of the retinoid receptor family are critically
involved in myeloid cell maturation. As part of the chimeric PML-RAR_pro-tein, the latter moiety is important for conferring sensitivity of the leukemic
blasts to all-trans retinoic acid ([atRA]; vitamin A), a now well-established
differentiative therapy in APL (11 ,12) . At the genomic level, breakpoints in
the PML gene occur in either introns 3 or 6, or rarely, within exon 6.
Breakpoints in the  RAR_ gene are distributed within intron 2. Thus, one of
three types of PML-RAR_mRNA fusion can occur and be detected in a given
case of APL (Fig. 1A): Short (S)-form, or BCR-3 (PML exon 3-RAR_exon 3,
approx 45–50%); Long (L)-form, or BCR–1 (PML exon 6-RAR_ exon 3,
approx 50–55%); and Variable (V)-form, or BCR-2 (PML exon 6 variable-RAR_ exon 3, approx 5–10%). The relative frequency of  PML-RAR_ types
differs in pediatric APL, in which V-form cases are more commonly encoun-tered than S-form cases (13) .
The inv(16)/t(16;16)(p13;q22) abnormality results in the fusion of the gene
encoding core binding factor-`subunit (CBF`), with the  MYH11 gene (14) .
The CBF protein complex is formed by heterodimerization of CBF`with its
heterodimeric DNA binding partner CBF_and acts transcriptionally at critical
genetic loci required for myeloid and lymphoid cell differentiation (reviewed
in ref. 15 ). The MYH11 gene encodes a type II smooth muscle myosin heavy
chain protein (SMMHC) and is not normally expressed in hematopoietic cells.
The CBF`-MYH11 gene fusion and resultant CBF`-SMMHC fusion protein is
thought to disrupt CBF transcriptional function in immature myeloid cells,
leading to interference with normal differentiation pathways in myelopoiesis
and, subsequently, AML (15) . Several different chimeric mRNA types can arise
from the CBF`-MYH11 gene fusion, owing mainly to breakpoint heterogene-ity in MYH11 (Fig. 1B). Fortunately, for diagnostic purposes, one fusion type
accounts for approx 90% of occurrences in inv(16)/t(16;16) AML and is easily
detected by RT-PCR analysis.
Interestingly, the t(8;21)(q22;q22) abnormality in AML also results in a
genetic fusion leading to disruption of CBF transcriptional regulation. In this
translocation, the AML1 gene encoding the CBF_subunit of CBF is fused to a
putative transcription factor gene, ETO (also designated MTG8), on the der(8)
chromosome (reviewed in ref. 16 ). The AML1 (CBF_) protein shares signifi-cant homology to the Drosophila pair rule gene runt (17 ,18) , and binds to DNA
“core” transcriptional sites in conjunction with CBF`. The AML1 gene is a
common target in leukemias, being involved in the t(3;21) associated with
myelodysplasia or therapy-induced AML (16) , and the t(12;21) anomaly present
in approx 25% of pediatric acute lymphoblastic leukemia (19,20). In the case of
Anomalies in Acute Myeloid Leukemias 117
t(8;21) AML, the resulting AML1(CBF_)-ETO fusion protein is similarly believed
to disrupt normal processes in myelopoietic differentiation and lead to leukemia.
Murine gene knock-out (for CBF`and AML1) and knock-in (for CBF`-MYH11
and AML1-ETO) models have identified a critical role for CBF in embryonic sur-vival and, most significantly, in definitive hematopoiesis (21–24). Although alter-native splice sites have been determined in both AML1 and ETO mRNAs (25), the
chimeric fusion transcript is highly uniform in the vast majority of cases of t(8;21)-positive AML (Fig. 1C).
Fig. 1. (A) Schematic of PML-RAR_Fusion in t(15;17) acute promyelocytic leuke-mia. (Top) Partial genomic map of PML (15q24) and RAR_(17q21) genes. Vertical
arrows indicate breakpoint regions. PML break sites can occur in intron 3, intron 6,
or rarely within exon 6. By contrast, RAR_breakpoints occur uniformly in intron 2.
(Bottom) Potential chimeric mRNAs produced by PML-RAR_break-fusion events:
S (short)-form/BCR-3, L (long)-form/BCR-1, and V (variable)-form/BCR-2. V-form
cases have a variable proportion of retained exon 6 nucleotides, usually with addi-tional nontemplated base additions or deletions. Alternative splicing out of  PML
exon 5 or exons 5 and 6 is also seen in L- and V-form fusion transcripts. Relative
locations of primers for first-round PCR (P3 and R4A) and second-round (nested)
PCR (P6 and R4B) are shown. Junction-specific oligoprobes for detection of S- and
L-form fusions are displayed as short bars.
118 Viswanatha
Fig. 1. (continued) (B) Schematic of major  CBF`-MYH11 fusions in inv(16)/
t(16;16) AML. The three most common CBF`-MYH11 fusion mRNA types are illus-trated. The type A chimeric mRNA is encountered in approx 90% of cases, whereas
types C and D are each detected in <5% of cases. Locations of C1 and M1 PCR prim-ers are shown. C2 and M7 primers can be utilized in allele-specific nested PCR confir-mation of the type A fusion transcript, as discussed in  Note 17. Detection of
CBF`-MYH11 fusion PCR products is accomplished by hybridization with an internal
CBF` sequence oligoprobe, indicated by the solid ovals.  (C) Schematic of  AML1-ETO fusion in t(8;21) AML. The diagram demonstrates the chimeric  AML1-ETO
mRNA, with relative locations of first-round PCR primers (AML1-A and ETO-A) and
second-round (nested) PCR primers (AML1-B and ETO-B) indicated. Confirmation
of the  AML1-ETO fusion is accomplished by hybridization to an internal  ETO
sequence oligoprobe, indicated by the black bar.
Anomalies in Acute Myeloid Leukemias 119
An enhanced understanding of the genetic basis of leukemogenesis will per-mit more precise correlation of molecular abnormalities with clinical disease
features and generate more-focused therapeutic efforts in AML. Such is the
case with the PML-RAR_fusion and the basis for the efficacy of atRA therapy
in APL. Data are accumulating that these “favorable outcome” fusion gene
leukemias (and perhaps others) may have a common molecular pathogenesis
involving interference with histone acetylation status and, correspondingly,
disruption of gene transcription processes required for cellular differentiation
(26–30) . This may provide yet another target for therapeutic intervention and
further underscores the importance of molecular diagnosis in AML. Finally, in
addition to detection of specific gene fusions for the identification of leukemic
subtypes, these abnormalities also serve as sensitive markers for monitoring of
minimal residual disease (MRD) in patients achieving remission status follow-ing therapy. An emerging paradigm in successful leukemia treatment is the
realization that clonal disease is not fully eradicated by conventional therapies,
but rather exists at submicroscopic levels (detectable by quantitative PCR
methods), with the propensity to fluctuate in quantity and activity over time.
Although beyond the scope of this chapter, the quantitative monitoring of MRD
will likely form a cornerstone of posttherapeutic disease evaluation in the leu-kemias; this is briefly mentioned at the end of the chapter.
2. Materials
2.1. RNA Isolation
1. RNAqueous small-scale RNA isolation kit (Ambion, Austin, TX) (see Note 1).
2. 3 M Sodium acetate (pH 5.2).
3. Cold 100% (absolute) ethanol.
4. Cold 75% ethanol.
5. 1 mM EDTA in diethylpyrocarbonate (DEPC) distilled water (see Note 2).
2.2. Reverse Transcription Reaction/cDNA Synthesis
1. 10X RT buffer: 100 mM Tris-HCl (pH 8.4), 500 mM KCl, 50 mM MgCl2 (see
Note 3).
2. 10 mM stock dNTPs solution (Boehringer Mannheim, Indianapolis IN).
3. Random hexamer solution (50 pmol/µL) (Perkin-Elmer, Foster City, CA).
4. RNasin RNase inhibitor (40 U/µL) (Promega, Madison, WI).
5. 50 mM Dithiothreitol (DTT).
6. MMLV-RT (200 U/µL) (Gibco-BRL/Life Technologies, Gaithersburg MD).
7. DEPC distilled H2O.
2.3. Polymerase Chain Reaction
1. 10X PCR buffer: 100 mM Tris-HCl (pH 8.4), 500 mM KCl, 15 mM MgCl2 (see
Note 3).
120 Viswanatha
2. 10 mM Stock dNTPs.
3. Sense and antisense primers (15–20 pmol/µL of each) (see Table 1).
4. AmpliTaq DNA polymerase (5 U/µL) (Perkin-Elmer/Roche, Foster City, CA).
5. DEPC distilled H2O.
2.4. Gel Analysis of PCR Product and Vacuum Blot Transfer
1. Standard powdered agarose for 1.5% agarose gel (3 g) (SeaKem ME, FMC Bio-products, ME).
2. Ethidium bromide (EtBr) solution (10 mg/mL) (Gibco-BRL/Life Technologies).
3. Stock 10X Tris-borate EDTA (TBE) buffer solution: 107.8 g of Tris-HCl, 55 g of
boric acid, 7.4 g of Na2EDTA per 1 L of sterile distilled H2O (0.5X TBE is used
for gel and 1X TBE for running buffer).
4. 10X Sample loading dye: 1 vol of 1% bromophenol blue, 1 vol of 1% xylene
cyanol, 2 vol of glycerol.
5. Horizontal gel box (40 cm) with 20-well combs (BRL Horizon 20.25; Life
Technologies).
Table 1
DNA Sequences for Primers and Probes (all 5v–3v)
t(15;17)/PML-RAR_
P3: ACCGATGGCTTCGACGAGTTC
R4A: AGCCCTTGCAGCCCTCACAG
P6: AATACAACGACAGCCCAGAAG
R4B: CTCACAGGCGCTGACCCCAT
L-form junction probea: GGTCTCAATGGCTGCCTCCCC
S-form junction probea: AATGGCTTTCCCCTGGGTGA
inv(16)/t(16;16)/CBFB-MYH11
C1: GCAGGCAAGGTATATTTGAAGG
M1: CTCTTCTCCTCATTCTGCTC
C2: ACACGCGAATTTGAAGATAGAG
M7: TTCTCCAGCTCATGGACCTCC
CBFB oligoprobea: ATAGAGACAGGTCTCATCGG
t(8;21)/AML1-ETO
AML1-A: AGCCATGAAGAACCAGG
ETO-A: AGGCTGTAGGAGAATGG
AML1-B: TACCACAGAGCCATCAAA
ETO-B: GTTGTCGGTGTAAATGAA
ETO oligoprobea: GTCTTCACATCCACAGGTGAGTCT
Beta2-microglobulin
B2 I sense GAAAAAGATGAGTATGCCTG
B2 II antisense: ATCTTCAAACCTCCATGATG
aOligonucleotide probes are 5v-biotinylated.
Anomalies in Acute Myeloid Leukemias 121
6. UV light box and instant camera setup with Polaroid 667 B+W film (Polaroid,
Cambridge, MA).
7. Nylon membrane (11 × 20 cm) (Biodyne PA; Pall-Biodyne, East Hills, NY) to
encompass area of gel being transferred.
8. Two clean plastic tubs.
9. Dilute (0.05 N) HCl (500 mL).
10. 0.4 M NaOH (2 L).
11. Bio-Rad Model 785 vacuum transfer apparatus with vacuum regulator (Bio-Rad,
Hercules, CA) (see Note 4).
12. UV crosslinking apparatus (UV Stratalinker; Stratagene, La Jolla, CA).
2.5. Nonisotopic Probe Hybridization
1. Stock 20X SSPE buffer: 210.4 g of NaCl, 27.6 g of NaH2PO4·H2O, 4.4 g of
NaOH, and 7.4 g of Na2EDTA per 1 L of sterile distilled H2O; buffer usually
requires gentle heat with constant stirring to solubilize completely. From this
buffer prepare the following:
a. Prehybridization solution (100 mL): 0.1X SSPE/0.5% sodium dodecyl sul-fate (SDS—from 10% SDS stock solution).
b. Blot wash solution (2 L): 2X SSPE/0.1% SDS.
c. 50-mL Aliquots of hybridization buffer: 5X SSPE/0.1% SDS (see Note 5).
2. 5v-Biotinylated specific oligonucleotide probe (5 pmol/µL) (see Table 1).
3. Heat-sealable plastic hybridization bags.
4. Several clean plastic tubs with lids.
5. Streptavidin-horseradish peroxidase (SA-HRP) (1 mg/mL) (Vector, Burling-ame, CA).
6. Enterochromaffin-like (ECL) chemiluminescence reagent solutions 1 and 2 (Amer-sham, Arlington Heights, IL).
7. Fluorescent marking pen.
8. Standard filter (blotting) paper.
9. Plastic wrap.
10. Radiographic film (Kodak XAR; Eastman-Kodak, Rochester, NY).
3. Methods
The PCR methods detailed herein are based on previously published proto-cols (14 ,31 ,32) , optimized for use in my laboratory (see Note 6). General labo-ratory principles pertaining to the performance of PCR and analysis of PCR
products should be followed (see Note 7).
3.1. Isolation of RNA
1. Obtain washed peripheral blood or bone marrow cell suspension following Ficoll
density gradient separation or RBC lysis (see Note 8). Gently pellet the cells and
decant all the supernatant.
2. Use the Ambion RNAqueous kit per the manufacturer’s directions.
122 Viswanatha
3. Following RNA elution from the column filter, collect in a 100  µL vol. Add
30 µL of 3 M Na acetate and 250 µL of cold absolute ethanol. Leave at –80°C for
15 min.
4. Centrifuge for 15 min at 12,000g and 4°C. Remove the supernatant carefully.
5. Gently wash the pellet with cold 75% ethanol. Air-dry the pellet for 5 min.
6. Resuspend the pellet in a volume of 1 mM EDTA to give an approximate concen-tration of 1 µg/µL. Calculate the concentration and yield by standard absorbance
spectrophotometry.
7. Store the RNA samples at –70°C.
3.2. Detection of t(15;17)-Associated PML-RAR_
Fusion by RT-PCR (see Note 9)
3.2.1. Reverse Transcription Reaction/cDNA Synthesis
1. Thaw RNA samples on ice. Aliquot 1  µg of each RNA sample into labeled
MicroAmp PCR tubes. Adjust the volume to 5 µL with sterile DEPC H2O. Keep
on ice or in a cold block.
2. Assemble RT master mix in a 1.5-mL microfuge tube, on ice, as follows: 7 µL
DEPC H2O, 2 µL of 10X RT buffer, 2 µL of 10 mM dNTPs, 1 µL (100 pmol) of
random hexamers, 0.5 µL of RNasin, 2 µL of DTT, 0.5 µL of MMLV-RT. These
amounts are per sample reaction and need to be multiplied by the number of
samples (n) being assayed. It is advisable to make enough RT master mix for one
extra reaction (i.e., n + 1). Gently vortex to mix.
3. Aliquot 15 µL of RT master mix to each sample tube proceeding in order from
the test samples to the positive control and finally the negative control (no RNA).
Mix by gentle, not vigorous, pipeting.
4. Place in a thermal cycler programmed as follows: 23°C for 10 min, 42°C for 30
min, 95°C for 5 min, then cooled to 4°C.
3.2.2. PCR for PML-RAR_ Fusion
1. Label a new set of MicroAmp tubes corresponding to the RT reaction set.
2. Transfer completed RT-reaction tubes to a PCR hood and aliquot 15 µL of syn-thesized cDNA to appropriately labeled new MicroAmp tubes proceeding in
order from the test samples to the positive control and finally the negative con-trol. This leaves 5 µL of cDNA remaining in the original tubes. Close the tube
caps and keep the samples on ice or in a cold block in the hood. Use 15 µL of the
cDNA product for the PML-RAR_PCR amplification and reserve 5 µL of prod-uct for the control `2-microglobulin amplification reaction.
3. Assemble the first-round  PML-RAR_ PCR master mix in a 1.5-mL microfuge
tube, on ice, as follows: 26.5 µL of DEPC H2O, 5 µL of 10X PCR buffer, 1 µL of
10 mM dNTPs, 1  µL each of primers P3 (sense) and R4A (antisense), 0.5  µL
of Taq DNA polymerase. This total of 35 µL of PCR master mix is per sample
and needs to be multiplied by the number of samples being tested, plus one extra
(n + 1). Gently vortex the tube with the lid closed.
Anomalies in Acute Myeloid Leukemias 123
4. Assemble the  `2-microglobulin control PCR master mix in a separate 1.5-mL
microfuge tube in exactly the same manner as in step 3; however, use 1 µL each
of primers `2M-sense and `2M-antisense instead. Gently vortex the tube to mix.
5. Start with the `2-microglobulin PCR tube set and add 35 µL of `2M master mix
to each of the respective MicroAmp tubes (resulting in a total volume of 40 µL
per reaction). Mix the contents of each tube gently and carefully. Similarly, add
35 µL of PML-RAR_master mix to each of the respective MicroAmp tubes in the
PML-RAR_PCR set, and mix the contents of each tube carefully (resulting in a
total volume of 50 µL per reaction). In either case, proceed in the order from the
test samples to the positive control and finally the negative (no RNA) control.
6. Place all the tubes in the thermal cycler, programmed as follows: 95°C for 5 min
(initial denaturation); 5 cycles of 95°C for 45 s, 62°C for 60 s, and 72°C for 30 s;
32 cycles of 95°C for 30 s, 60°C for 45 s, and 72°C for 60 s; then 72°C for 5 min
(final extension) and cool to 4°C. Following the PCR reaction, remove the tubes
from the thermal cycler and transfer the first-round PML-RAR_product tubes to
the PCR hood in a cold block. Put the completed `2-microglobulin PCR tubes on
ice or in a refrigerator.
7. Label one final set of fresh MicroAmp PCR tubes. Transfer 1 µL from the first-round PML-RAR_product tubes to the corresponding new PCR tubes. Close the
tube caps.
8. Assemble the second-round PML-RAR_master mix in a 1.5-mL microfuge tube,
on ice, as follows: 40.5 µL of DEPC H2O, 5 µL of 10X PCR buffer, 1 µL of 10
mM dNTPs, 1 µL each of second-round primers P6 (sense) and R4B (antisense),
0.5 µL of Taq DNA polymerase. This total of 49 µL of PCR master mix is per
sample and needs to be multiplied by the number of samples being tested, plus
one extra (n + 1). Gently vortex the tube with the lid closed.
9. Add 49  µL of second-round  PML-RAR_ master mix to each MicroAmp tube,
proceeding in the order from the test samples to the positive control and finally
the negative control. Mix the contents of each tube carefully (resulting in a total
volume of 50 µL per reaction).
10. Place the tubes in the thermal cycler, programmed as follows: 95°C for 3 min
(initial denaturation); 5 cycles of 95°C for 45 s, 62°C for 60 s, and 72°C for 30 s;
25 cycles of 95°C for 30 s, 62°C for 45 s, and 72°C for 60 s; then 72°C for 5 min
(final extension) and cool to 4°C.
11. Keep the first- and second-round  PML-RAR_ PCR products and  `2-micro-globulin products in a refrigerator or –20°C freezer until ready for gel
electrophoresis.
3.2.3. Gel Electrophoresis Analysis
and Vacuum Blot Transfer of PML-RAR_ PCR Products
1. Prepare a 1.5% agarose gel (with 0.5X TBE) (see Note 10).
2. In a microtiter plate, add 15 µL of PCR products to 2.5 µL of 10X sample loading
dye. A DNA size marker is also used, such as the 123-bp ladder (Gibco-BRL/
Life Technologies). Mix briefly and load the first-round P3-R4A PCR products
124 Viswanatha
in one set of lanes, followed by the second-round P6-R4B products, separating
the two groups of PCR products by one or two empty lanes. Load  `2-micro-globulin products in a separate set of lanes. Carry out electrophoresis at 180 V for
1.5–2 h.
3. Following electrophoresis, observe and photograph the gel under UV illumin-ation. Proper eye protection is mandatory when viewing gels by UV light.
Standard black-and-white Polaroid photographs are obtained with a photograph
stand and instant film (see Note 11). Expected PCR product sizes are as follows
(Fig. 2):
a. t(15;17)/PML-RAR_: L-form (BCR-1) first round—3 bands, range 280–700
bp; second round—380 bp; V-form (BCR-2) first round—3 bands, range
280 to approx 700 bp; second round—<380 bp; S-form (BCR-3) first round
only—221 bp.
b. `2-microglobulin: 120 bp.
4. Place the gel in a plastic tub and cover with 300–400 mL of dilute (0.05 N) HCl
(acid-nicking step). Place on a gyrating platform on the gentle setting for 10 min.
During this time, allow the precut nylon membrane to equilibrate in 300–400 mL
of 0.4 M NaOH.
5. Decant dilute HCl and add an equal amount of 0.4 M NaOH over the gel.
6. Assemble the vacuum transfer apparatus according to the manufacturer’s direc-tions. Align the gel over the nylon membrane, ensuring that the wells are outside
of the vacuum gasket seal. Slowly engage the vacuum to negative pressure of 4 to
5 mmHg. Pour approx 250 mL of 0.4 M NaOH over the gel to cover the area for
transfer. Transfer the gel products to the nylon membrane for 1 h.
7. Disassemble the vacuum transfer apparatus and remove the nylon membrane.
Place the membrane in a UV crosslinker and crosslink the DNA to the membrane
at a 500-µJ power setting. Label the well locations at the top of the gel using a
soft pencil and allow the membrane to dry.
8. Cut and separate a section containing P3-R4A PCR products from the section
containing P6-R4B PCR products, label appropriately, and trim off the excess
membrane. If `2-microglobulin products have been transferred on the same mem-brane, cut away the area containing `2-microglobulin product lanes and discard.
The P3-R4A reaction products will be hybridized with the S-form junction probe
whereas the P6-R4B products will be hybridized with the L-form junction probe
(see Subheading 3.2.4.). V-form PML-RAR_type fusions are not amenable to
junctional oligoprobe analysis; this is discussed in Subheading 3.2.5.
Fig. 2. (see facing page) t(15;17)/PML-RAR_PCR product analysis. (Top) Agar-ose gel electrophoresis of PML-RAR_PCR products. L, size marker (123-bp ladder).
Lanes A–E, first-round amplification products (with P3 and R4A primers): A, NB4
positive control for L-form fusion; B, APL patient positive for L-form fusion; C, APL
patient positive for V-form fusion; D, APL patient positive for S-form fusion; E, nega-tive control (no RNA). For L- and V-form fusions, a three-band pattern is observed
owing to alternative splicing out of PML exons 5, and 5 + 6. The full-length transcript
Anomalies in Acute Myeloid Leukemias 125
Fig. 2. (continued) PCR product  (uppermost band) is 700 bp for L-form fusions
and typically less than this for V-form fusions. The S-form fusion PCR product is
221 bp. Lanes F–I, second-round (nested) PCR of samples A, B, C, and E (with prim-ers P6 and R4B): F, NB4 cell line; G, L-form positive APL patient; H, V-form positive
APL patient; I, negative control (no RNA). L-form nested PCR products reveal a
prominent band at 380 bp, whereas the V-form product is slightly smaller. (Bottom)
Two blots of junction-specific oligoprobe hybridization to detect S- and L-form fusions.
Lanes A–E correspond to the PCR products from the gel above and represent first-round amplification products. These have been hybridized with the S-form oligoprobe.
Lanes F–I correspond to the PCR products from the gel above and represent second-round amplification products, hybridized with the L-form oligoprobe. Note the absence of
hybridization signal for the V-form PCR product (lane H), a useful distinguishing feature
when V-form PCR products are similar in size to L-form PML-RAR_ fusions.
126 Viswanatha
3.2.4. Nonisotopic (Chemiluminescent)
Probe Hybridization for PML-RAR_ Fusions
1. Warm to 60°C in a water bath 50 mL of prehybridization solution (0.1X SSPE/
0.5% SDS) and two 15-mL aliquots (one per separate hybridization reaction) of
hybridization buffer (5X SSPE/0.1% SDS).
2. Place nylon membranes of the (first round) P3-R4A and (second round) P6-R4B
PCR products in appropriately labeled, heat-sealable plastic bags and add 15 mL
of prehybridization buffer to each. Displace bubbles through the open end of the
bag, and then heat-seal the bag and place in a 60°C oven, on a rocking platform.
Allow 30 min for prehybridization.
3. Add 5  µL of the biotinylated junction-specific oligoprobes (S-form junction
and L-form junction) to respective 15-mL aliquots of warm hybridization
buffer and gently mix (final probe concentrations approx 1.5–2 pmol/mL of
buffer). Remove and cut the plastic bags open across the top. Drain the
prehybridization buffer and carefully add the 15 mL of the hybridization
buffer with the specific oligoprobe to the corresponding bags (i.e., S-form
junction to the membrane with P3-R4A products and L-form junction to the
membrane with P6-R4B products). Express excess air and bubbles, and then
heat-seal the bags just above the nylon membrane. Check for leaks and place in
a 60°C oven for 1 h. The hybridization reactions can be extended to overnight
for convenience.
4. Prepare 2 L of blot wash solution (2X SSPE/0.1% SDS). Aliquot 500 mL of wash
solution and heat to 60°C in a water bath. Be sure to verify the temperature of the
solution. Leave the remaining wash solution covered at room temperature. Warm
two clean empty plastic tubs in a 60°C oven.
5. Remove the hybridization bags, cut across to open, and drain the hybridization
solution. Remove the nylon membranes and place in separate plastic tubs, and
then add approx 125 mL of warm (60°C) blot wash buffer to each. Cover the tubs
and place in a 60°C oven on a rocking platform for 10 min. Repeat this step once
with the remaining warm blot wash solution (see Note 12).
6. Prepare SA-HRP solution in a clean plastic tub using 250 mL of room tempera-ture blot wash solution and 10 µL of SA-HRP (final concentration of 40 ng/mL).
7. Following the second warm (60°C) wash, briefly rinse the membranes with
approx 100 mL of room temperature wash solution, and transfer to the SA-HRP
solution. Place on a slowly gyrating platform for 15 min.
8. Drain the SA-HRP solution, briefly rinse with 100 mL of room temperature blot
wash solution and decant, and add another 400 mL of blot wash solution. Place
on a gyrating platform for 5–10 min. Repeat the wash with another 400 mL of
room temperature solution (see Note 13).
9. Place the wet membranes on clean dry filter paper. Quickly pipet 5 mL each of
ECL chemiluminescence reagents 1 and 2 into a clean glass tray and mix by
gentle agitation. Place each membrane in the tray and coat thoroughly with ECL
reagents for 1 min. Remove and affix the membranes to clean precut filter paper.
Mark well locations with a fluorescent pen, label blots as to probe type, and seal
Anomalies in Acute Myeloid Leukemias 127
the membranes over with plastic wrap. Develop chemiluminescent image on
X-ray film as usual (see Note 14).
3.2.5. Interpretation: t(15;17)/PML-RAR_ Results
As indicated in Subheading 1., the PML-RAR_gene fusion may produce three
distinct chimeric transcript types: Long (L)-form, or BCR-1 (PML exon 6-RAR_
exon 3); Short (S)-form, or BCR-3 (PML exon 3-RAR_ exon 3); Variable
(V)-form, or BCR-2 (PML exon 6 variable-RAR_ exon 3). Overall, the L- and
S-forms account for approx 90% of PML-RAR_fusions, with the V-form being
relatively rare (see Note 15). The V-form is unique in that the PML breakpoint
occurs within exon 6 and nontemplated nucleotides are often inserted or nucle-otides deleted to maintain an open reading frame in the fusion mRNA. For the first-round PCR products, the L- and V-form products give rise to three distinct bands
(Fig. 2, Top, lanes A–E; also see Note 16). The upper two bands in V-form cases
are typically smaller in size than for L-form cases, although rare V-form tumors
may have similar, or longer transcript lengths. This reflects the “variable” nature of
the PML exon 6 breakpoint in V-form cases. The S-form PCR product forms a
single band of much shorter length. Second-round (nested) PCR analysis confirms
the presence of the L- and V-form PML-RAR_fusions (Fig. 2, Top, lanes F–I) and,
in combination with L-form junction oligoprobe hybridization (Fig. 2, Bottom,
lanes F–I), allows distinction of these fusion types when they may be of similar
size. Junction-specific oligoprobe hybridization for the S-form PML-RAR_fusion
is also illustrated in  Fig. 2 (bottom panel, lane D). More important, for the
RT-PCR analysis described herein, gel electrophoresis will detect all three
fusion types. However, because of the variable nature of the junctional
sequence in V-form cases, a specific probe for this region cannot be developed as
for the L- and S-form PCR products. Hence, V-form cases are recognized on aga-rose gel alone. In some cases, we have resorted to direct DNA sequence analysis of
suspected V-form products to confirm the fusion type.
3.3. Detection of the inv(16)/t(16;16)-Associated
CBF`-MYH11 Fusion by RT-PCR
3.3.1. Reverse Transcription Reaction/cDNA Synthesis (see Note 9)
1. Thaw RNA samples on ice. Aliquot 1 to 2 µg of each RNA sample into labeled
MicroAmp PCR tubes. Adjust the volume to 5 µL with sterile DEPC H2O. Keep
on ice or in a cold block.
2. Assemble RT master mix in a 1.5-mL microfuge tube, on ice, as follows: 7 µL of
DEPC H2O, 2 µL of 10X RT buffer, 2 µL of 10 mM dNTPs, 1 µL of random
hexamers, 0.5 µL of RNasin, 2 µL of DTT, 0.5 µL of MMLV-RT. These amounts
are per sample reaction and need to be multiplied by the number of samples being
assayed. It is advisable to make enough RT master mix for one extra reaction
(n + 1). Gently vortex to mix.
128 Viswanatha
3. Aliquot 15 µL of RT master mix to each sample tube proceeding in order from
the test samples to the positive control and finally the negative control (no RNA).
Mix by gentle, not vigorous, pipeting.
4. Place in a thermal cycler programmed as follows: 23°C for 10 min, 42°C for 60
min, 95°C for 5 min, then cooled to 4°C.
3.3.2. PCR for CBF`-MYH11 Fusion
1. Label a new set of MicroAmp tubes corresponding to the RT reaction set.
2. Transfer completed RT-reaction tubes to a PCR hood and aliquot 15 µL of syn-thesized cDNA to appropriately labeled new MicroAmp tubes tube proceeding in
order from the test samples to the positive control and finally the negative control
(no RNA). This leaves 5 µL of cDNA remaining in the original tubes. Close the
tube caps and keep the samples on ice or in a cold block in the hood. Use 15 µL
of the cDNA product for the CBF`-MYH11 PCR amplification and reserve 5 µL
of product for the control `2-microglobulin amplification reaction.
3. Assemble the  CBF`-MYH11 PCR master mix in a 1.5-mL microfuge tube, on
ice, as follows: 26.5 µL of DEPC H2O, 5 µL of 10X PCR buffer, 1 µL of 10 mM
dNTPs, 1 mL each of primers C1 (sense) and M1 (antisense), 0.5 µL of Taq DNA
polymerase. This total of 35 µL PCR master mix is per sample and needs to be
multiplied by the number of samples being tested, plus one extra (n + 1). Gently
vortex the tube with the lid closed.
4. Assemble the  `2-microglobulin control PCR master mix in a separate 1.5-mL
microfuge tube in exactly the same manner as in step 3; however, use 1 µL each
of primers `2M-sense and `2M-antisense instead. Gently vortex the tube to mix.
5. Start with the `2-microglobulin PCR tube set and add 35 µL of `2M master mix
to each of the respective MicroAmp tubes (resulting in a total volume of 40 µL
per reaction). Mix the contents of each tube gently and carefully. Similarly, add
35 µL of CBF`-MYH11 master mix to each of the respective MicroAmp tubes in
the CBF`-MYH11 PCR set, and mix the contents of each tube carefully (resulting
in a total volume of 50 µL per reaction). In either case, proceed in the order from
the test samples to the positive control and finally the negative control.
6. Place all the tubes in the thermal cycler, programmed as follows: 95°C for 5 min (ini-tial denaturation); 3 cycles of 95°C for 30 s, 59°C for 45 s, and 72°C for 90 s; 30 cycles
of 95°C for 30 s, 57°C for 30 s, and 72°C for 60 s; then 72°C for 8 min (final exten-sion) and cool to 4°C. Keep the CBF`-MYH11 PCR products and `2-microglobulin
products in a refrigerator or –20°C freezer until ready for gel electrophoresis.
3.3.3. Gel Electrophoresis Analysis
and Vacuum Blot Transfer of CBF`-MYH11 PCR Products
1. Prepare a 1.5% agarose gel (with 0.5X TBE) (see Note 10).
2. In a microtiter plate, add 15 µL of PCR products to 2.5 µL of 10X sample loading
dye. A DNA size marker is also used, such as the 123-bp ladder (Gibco-BRL/
Life Technologies). Mix briefly and load C1-M1 PCR products in one set of
lanes, followed by  `2-microglobulin products, separating the two groups of PCR
products by one or two empty lanes. Carry out electrophoresis at 180 V for 1.5–2 h.
Anomalies in Acute Myeloid Leukemias 129
3. Following electrophoresis, observe and photograph the gel under UV illumina-tion. Proper eye protection is mandatory when viewing gels by UV light. Stan-dard black-and-white Polaroid photographs are obtained with a photograph stand
and instant film (see Note 11). Expected PCR product sizes are as follows
(Fig. 3A):
a. inv(16)/t(16;16)  CBF`-MYH11: Type A, 415 bp; Type C, 1.2 kb; Type D,
1.4 kb; other types rare. (Types A, C, and D collectively account for approx
95% of CBF`-MYH11 fusions [see Subheading 3.3.5.])
b. `2-Microglobulin: 120 bp.
4. Place the gel in a plastic tub and cover with 300–400 mL of dilute (0.05 N) HCl
(acid-nicking step). Place on a gyrating platform on gentle setting for 10 min.
During this time, allow the precut nylon membrane to equilibrate in 300–400 mL
of 0.4 M NaOH.
5. Decant the dilute HCl and add an equal amount of 0.4 M NaOH over the gel.
6. Assemble the vacuum transfer apparatus according to the manufacturer’s direc-tions. Align the gel over the nylon membrane, ensuring that the wells are outside
of the vacuum gasket seal. Slowly engage the vacuum to negative pressure of 4 to
5 mmHg. Pour approx 250 mL of 0.4 M NaOH over the gel to cover the area for
transfer. Transfer the gel products to the nylon membrane for 1 h.
7. Disassemble the vacuum transfer apparatus and remove the nylon membrane.
Place the membrane in a UV crosslinker and crosslink the DNA to the membrane
at a 500-µJ power setting. Label the well locations at the top of the gel using a
soft pencil and allow the membrane to dry.
8. Cut and separate the section containing C1-M1 PCR products, label appropri-ately, and then trim off the excess membrane. If `2-microglobulin products have
been transferred on the same membrane, cut away the area containing the `2-micro-globulin product lanes and discard.
3.3.4. Nonisotopic (Chemiluminescent) Probe
Hybridization for CBF`-MYH11 Fusions
1. Warm to 60°C in a water bath 50 mL of prehybridization solution (0.1X SSPE/
0.5% SDS) and a 15-mL aliquot of hybridization buffer (5X SSPE/0.1% SDS).
2. Place a nylon membrane of C1-M1 PCR products in an appropriately labeled,
heat-sealable plastic bag and add 15 mL of prehybridization buffer. Displace
bubbles through the open end of the bag, and then heat-seal the bag and place in
a 60°C oven, on a rocking platform. Allow 30 min for prehybridization.
3. Add 5  µL of the biotinylated  CBF` internal sequence oligoprobe (CBF`
oligoprobe) to a 15-mL aliquot of warm hybridization buffer and gently mix
(final probe concentration of approx 1.5–2 pmol/mL of buffer). Remove and cut
the plastic bag open across the top. Drain the prehybridization buffer and care-fully add 15 mL of the hybridization buffer with the specific oligoprobe to the
bag. Express excess air and bubbles, and then heat-seal the bag just above the
nylon membrane. Check for leaks and place in a 60°C oven for 1 h. The hybrid-ization reaction can be extended to overnight for convenience.
130 Viswanatha
Fig. 3A.  inv(16)/t(16;16)CBF`-MYH11 PCR product analysis. (Top) Agarose gel
electrophoresis of C1-M1 PCR products. L, size marker (123-bp ladder). Lanes A, B,
and E: type A fusion product (415 bp). Lane D: type D fusion product (1.4 kb). Lanes
F and G: type C fusion product (1.2 kb). Lane marked (–) is a negative control (no
RNA). Lane C is a novel CBF`-MYH11 fusion product that closely mimics the type A
fusion in size. (Bottom) Internal  CBF` oligoprobe hybridization for detection of
CBF`-MYH11 PCR products. Lanes are exactly as depicted at top. Novel  CBF`-MYH11 fusion (lane C) does not hybridize with this probe and required sequencing
for confirmation (see Note 18). Figures reprinted with permission of publisher, with
minor modifications (52) .
Anomalies in Acute Myeloid Leukemias 131
4. Prepare 2 L of blot wash solution (2X SSPE/0.1% SDS). Aliquot 500 mL of wash
solution and heat to 60°C in a water bath. Be sure to verify the temperature of the
solution. Leave the remaining wash solution covered at room temperature. Warm
a clean empty plastic tub in the 60°C oven.
5. Remove the hybridization bag, cut across to open, and drain the hybridization
solution. Remove the nylon membrane and place in a plastic tub, and then add
250 mL of warm (60°C) blot wash buffer. Cover the tub and place in a 60°C oven
on a rocking platform for 10 min. Repeat this step once with the remaining warm
blot wash solution (see Note 12).
6. Prepare SA-HRP solution in a clean plastic tub using 250 mL of room tempera-ture blot wash solution and 10 µL of SA-HRP (final concentration of 40 ng/mL).
7. Following the second warm (60°C) wash, briefly rinse the membranes with
approx 100 mL of room temperature wash solution, and transfer to the SA-HRP
solution. Place on a slowly gyrating platform for 15 min.
Fig. 3B.  Application of type A CBF`-MYH11 junctional oligoprobe. Detection of
type A  CBF`-MYH11 fusion in a cell dilution experiment using a junction-specific
oligonucleotide probe. Patient cells positive for type A  CBF`-MYH11 fusion tran-script were serially diluted in normal bone marrow cells and analyzed by C1-M1
RT-PCR. Lane A, undiluted patient cells; lane B, 1 10 dilution; lane C, 1 20 dilution;
lane D, 1 100 dilution; lane E, 1 103 dilution; lane F, 1 104 dilution; lane G, pure
normal bone marrow. Lane marked (–) is a negative control (no RNA). Figure
reprinted with permission of publisher, with minor modifications (52) .
132 Viswanatha
8. Drain the SA-HRP solution, briefly rinse with 100 mL of room temperature blot
wash solution and decant, and then add another 400 mL of blot wash solution.
Place on a gyrating platform for 5–10 min. Repeat the wash with another 400 mL
of room temperature solution (see Note 13).
9. Place the wet membranes on clean dry filter paper. Quickly pipet 5 mL each of
ECL chemiluminescence reagents 1 and 2 into a clean glass tray and mix by
gentle agitation. Place each membrane in the tray and coat thoroughly with the
ECL reagents for 1 min. Remove and affix the membranes to clean precut filter
paper. Mark the well locations with a fluorescent pen, label the blots as to probe
type, and seal over the membranes with plastic wrap. Develop chemiluminescent
image on X-ray film as usual (see Note 14).
3.3.5. Interpretation: inv(16)/t(16;16) CBF`-MYH11 Results
The inv(16)/t(16;16)-associated CBF`-MYH11 fusion results in formation
of a type A chimeric transcript in approx 90% of positive cases. The longer
type C and type D fusion PCR products account for approx 5% of CBF`-MYH11
fusions overall; thus, the PCR assay and hybridization reaction described will
detect nearly all the commonly encountered CBF`-MYH11 fusions occurring
in AML. Although several other CBF`-MYH11 fusion types have been described
(15) , these are relatively rare and arise mainly owing to significant heterogene-ity in MYH11 gene break point–fusion sites. By contrast, the CBF`fusion site
in chimeric CBF`-MYH11 mRNAs is constant with very rare exceptions (33) .
Therefore, an internal  CBF` oligoprobe can be utilized to identify  CBF`-MYH11 PCR products following Southern blot transfer (Fig. 3A). The type A
fusion can also be specifically confirmed by using a junctional sequence
oligoprobe (inv(16) junction A, 5v-3v:GCTCATGGACCTCCATTTCC)
applied to blots of C1-M1 PCR products, although this may be a more useful
reagent in RT-PCR assessment of minimal residual disease (Fig. 3B).
Confirmation of the common type A CBF`-MYH11 fusion product (415 bp)
can also be obtained following C1-M1 primer PCR, by allele-specific nested
PCR (34) , usually obviating the need to perform a probe hybridization (see
Note 17). However, this nested PCR method will not detect other (non–type A)
CBF`-MYH11 types (see Note 18).
3.4. Detection of the t(8;21)-Associated
AML1-ETO Fusion by RT-PCR
3.4.1. Reverse Transcription Reaction/cDNA Synthesis (see Note 9)
1. Thaw RNA samples on ice. Aliquot 1  µg of each RNA sample into labeled
MicroAmp PCR tubes. Adjust the volume to 5 µL with sterile DEPC H2O. Keep
on ice or in a cold block.
2. Assemble RT master mix in a 1.5-mL microfuge tube, on ice, as follows: 7 µL of
DEPC H2O, 2 µL of 10X RT buffer, 2 µL of 10 mM dNTPs, 1 µL of random
Anomalies in Acute Myeloid Leukemias 133
hexamers, 0.5 µL of RNasin, 2 µL of DTT, 0.5 µL of MMLV-RT. These amounts
are per sample reaction and need to be multiplied by the number of samples being
assayed. It is advisable to make enough RT master mix for one extra reaction
(n + 1). Gently vortex to mix.
3. Aliquot 15 µL of RT master mix to each sample tube proceeding in order from
the test samples to the positive control and finally the negative control (no RNA).
Mix by gentle, not vigorous, pipetting.
4. Place in a thermal cycler programmed as follows: 23°C for 10 min, 37°C for 60
min, 95°C for 5 min, then cooled to 4°C.
3.4.2. PCR for AML1-ETO Fusion
1. Label a new set of MicroAmp tubes corresponding to the RT reaction set.
2. Transfer the completed RT-reaction tubes to a PCR hood and aliquot 15 µL of
synthesized cDNA to appropriately labeled new MicroAmp tubes tube proceed-ing in order from the test samples to the positive control and finally the negative
control (no RNA). This leaves 5  µL of cDNA remaining in the original tubes.
Close the tube caps and keep the samples on ice or in a cold block in a hood. Use
15 µL of cDNA product for the AML1-ETO PCR amplification and reserve 5 µL
of product for the control `2-microglobulin amplification reaction.
3. Assemble the first-round  AML1-ETO PCR master mix in a 1.5-mL microfuge
tube, on ice, as follows: 26.5 µL of DEPC H2O, 5 µL of 10X PCR buffer, 1 µL of
10 mM dNTPs, 1 µL each of primers AML1-A (sense) and ETO-A (antisense),
0.5 µL of Taq DNA polymerase. This total of 35 µL of PCR master mix is per
sample and needs to be multiplied by the number of samples being tested, plus
one extra (n + 1). Gently vortex the tube with the lid closed.
4. Assemble the  `2-microglobulin control PCR master mix in a separate 1.5-mL
microfuge tube in exactly the same manner as in step 3; however, use 1 µL each
of primers `2M-sense and `2M-antisense instead. Gently vortex the tube to mix.
5. Start with the `2-microglobulin PCR tube set and add 35 µL of `2M master mix
to each of the respective MicroAmp tubes (resulting in a total volume of 40 µL
per reaction). Mix the contents of each tube gently and carefully. Similarly, add
35 µL of AML1-ETO master mix to each of the respective MicroAmp tubes in the
AML1-ETO PCR set and mix the contents of each tube carefully (resulting in a
total volume of 50 µL per reaction). In either case, proceed in the order from the
test samples to the positive control and finally the negative control.
6. Place all the tubes in the thermal cycler, programmed as follows: 95°C for 3 min
(initial denaturation); 30 cycles of 95°C for 30 s; 55°C for 30 s; 72°C for 60 s;
then 72°C for 5 min (final extension) and cool to 4°C. Following the PCR reac-tion, remove the tubes from the thermal cycler and transfer the first-round AML1-ETO product tubes to the PCR hood in a cold block. Put the completed
`2-microglobulin PCR tubes on ice or in a refrigerator.
7. Label one final set of fresh MicroAmp PCR tubes. Transfer 2 µL from the first-round AML1-ETO product tubes to the corresponding new PCR tubes. Close the
tube caps.
134 Viswanatha
8. Assemble the second-round (nested)  AML1-ETO master mix in a 1.5-mL
microfuge tube, on ice, as follows: 39.5 µL of DEPC H2O, 5  µL of 10X PCR
buffer, 1  µL of 10 mM dNTPs, 1  µL each of second-round internal primers
AML1-B (sense) and ETO-B (antisense), 0.5 µL of Taq DNA polymerase. This
total of 48 µL of PCR master mix is per sample and needs to be multiplied by the
number of samples being tested, plus one extra (n + 1). Gently vortex the tube
with the lid closed.
9. Add 48  µL of second-round  AML1-ETO master mix to each MicroAmp tube,
proceeding in the order from test samples to the positive control and finally the
negative control. Mix the contents of each tube carefully (resulting in a total
volume of 50 µL per reaction).
10. Place the tubes in the thermal cycler, using the same program parameters as for
the first-round AML1-ETO PCR.
11. Keep the first- and second-round AML1-ETO PCR products and `2-microglobulin
products in a refrigerator or –20°C freezer until ready for gel electrophoresis.
3.4.3. Gel Electrophoresis Analysis and Vacuum Blot
Transfer of AML1-ETO PCR Products
1. Prepare a 1.5% agarose gel (with 0.5X TBE) (see Note 10).
2. In a microtiter plate, add 15 µL of PCR products to 2.5 µL of 10X sample loading
dye. A DNA size marker is also used, such as the 123-bp ladder (Gibco-BRL/
Life Technologies). Mix briefly and load first-round AML1-A/ETO-A PCR prod-ucts in one set of lanes, followed by second-round AML1-B/ETO-B products,
separating the two groups of PCR products by one or two empty lanes. Load
`2-microglobulin products in a separate set of lanes. Carry out electrophoresis at
180 V for 1.5–2 h.
3. Following electrophoresis, observe and photograph the gel under UV illumina-tion. Proper eye protection is mandatory when viewing gels by UV light.
Standard black-and-white Polaroid photographs are rendered with a photograph
stand and instant film (see Note 11). Expected PCR product sizes are as follows
(Fig. 4):
a. t(8;21)/AML1-ETO: first round, 338 bp (May not be observed after first-round
amplification [see Subheading 3.4.5.]); second round, 185 bp.
b. `2-microglobulin: 120 bp.
4. Place the gel in a plastic tub and cover with 300–400 mL of dilute (0.05 N) HCl
(acid-nicking step). Place on a gyratory platform on gentle setting for 10 min.
During this time, allow the precut nylon membrane to equilibrate in 300–400 mL
of 0.4 M NaOH.
5. Decant the dilute HCl and add an equal amount of 0.4 M NaOH over the gel.
6. Assemble the vacuum transfer apparatus according to the manufacturer’s direc-tions. Align the gel over the nylon membrane, ensuring that wells are outside of
the vacuum gasket seal. Slowly engage the vacuum to negative pressure of 4 to
5 mmHg. Pour approx 250 mL of 0.4 M NaOH over the gel to cover the area for
transfer. Transfer the gel products to the nylon membrane for 1 h.
Anomalies in Acute Myeloid Leukemias 135
7. Disassemble the vacuum transfer apparatus and remove the nylon membrane.
Place the membrane in a UV crosslinker and crosslink the DNA to the membrane
at a 500-µJ power setting. Label the well locations at the top of the gel using a
soft pencil and allow the membrane to dry.
8. Cut and separate the sections containing first-round AML1-A/ETO-A and sec-ond-round AML1-B/ETO-B PCR products, label appropriately, and then trim off
the excess membrane from each. If `2-microglobulin products have been trans-ferred on the same membrane, cut away the area containing  `2-microglobulin
product lanes and discard.
Fig. 4. t(8;21)/AML1-ETO PCR product analysis. Gel blot illustrating detection of
AML1-ETO fusion PCR products with internal  ETO sequence oligoprobe. Figure
shows hybridized blot of second-round amplification products (with AML1-B and
ETO-B primers). Lane marked (+) is the positive control (Kasumi-1 cell line). Lanes
A and B are patient samples, with B demonstrating the presence of the AML1-ETO
fusion. Lane marked (–) is a negative control (no RNA).
136 Viswanatha
3.4.4. Nonisotopic (Chemiluminescent) Probe
Hybridization for AML1-ETO Fusion
1. Warm to 60°C in a water bath 50 mL of prehybridization solution (0.1X SSPE/
0.5% SDS) and a 15-mL aliquot of hybridization buffer (5X SSPE/0.1% SDS).
2. Place nylon membranes of first- and second-round  AML1-ETO PCR products
back-to-back in an appropriately labeled, heat-sealable plastic bag and add
15 mL of prehybridization buffer. Displace bubbles through the open end of the
bag, and heat-seal the bag, and place in a 60°C oven, on a rocking platform.
Allow 30 min for prehybridization.
3. Add 5 µL of the biotinylated ETO internal sequence oligoprobe (ETO oligoprobe)
to a 15-mL aliquot of warm hybridization buffer and gently mix (final probe
concentration of approx 1.5–2 pmol/mL of buffer). Remove and cut the plastic
bag open across the top. Drain the prehybridization buffer and carefully add
15 mL of the hybridization buffer with the specific oligoprobe to the bag.
Express excess air and bubbles, and then heat-seal the bag just above the nylon
membranes. Check for leaks and place in a 60°C oven for 1 h. The hybridization
reaction can be extended to overnight for convenience.
4. Prepare 2 L of blot wash solution (2X SSPE/0.1% SDS). Aliquot 500 mL of wash
solution and heat to 60°C in a water bath. Be sure to verify the temperature of the
solution. Leave the remaining wash solution covered at room temperature. Warm
a clean empty plastic tub in a 60°C oven.
5. Remove the hybridization bag, cut across to open, and drain the hybridization
solution. Remove the nylon membranes, place both in a plastic tub, and add
250 mL of warm (60°C) blot wash buffer. Cover and place in a 60°C oven on a
rocking platform for 10 min. Repeat this step once with the remaining warm blot
wash solution (see Note 12).
6. Prepare SA-HRP solution in a clean plastic tub using 250 mL of room tem-perature blot wash solution and 10  µL of SA-HRP (final concentration of
40 ng/mL).
7. Following the second warm (60°C) wash, briefly rinse the membranes with
approx 100 mL of room temperature wash solution and transfer to the SA-HRP
solution. Place on a slowly gyrating platform for 15 min.
8. Drain the SA-HRP solution, briefly rinse with 100 mL of room temperature blot
wash solution, and decant. Then add another 400 mL of blot wash solution. Place
on a gyrating platform for 5–10 min. Repeat the wash with another 400 mL of
room temperature solution (see Note 13).
9. Place the wet membranes on clean dry filter paper. Quickly pipet 5 mL each of
ECL chemiluminescence reagents 1 and 2 into a clean glass tray and mix by
gentle agitation. Place each membrane in the tray and coat thoroughly with ECL
reagents for 1 min. Remove and affix the membranes to clean precut filter paper.
Mark the well locations with a fluorescent pen, label the blots as to probe type,
and seal over the membranes with plastic wrap. Develop chemiluminescent
image on X-ray film as usual (see Note 14).
Anomalies in Acute Myeloid Leukemias 137
3.4.5. Interpretation: t(8;21)/AML1-ETO Results
In contrast to the t(15;17)/PML-RAR_ and inv(16) or t(16;16)/  CBF`-MYH11 chimeric fusions, the  AML1-ETO abnormality is characterized by a
constant and predictable fusion type (Fig. 4). In a minority of cases, detection
of a slightly larger transcript has been described in addition to the usual AML1-ETO transcript, owing to an additional 68 bp of ETO sequence (25) . However,
significant heterogeneity in AML1-ETO breakpoint-fusion sites is not a feature
of this genetic abnormality. Nested PCR is performed in this assay to
detect the presence of the AML1-ETO mRNA, because first-round PCR alone
may not achieve sufficient sensitivity. Some protocols have been described
that appear to obtain good results with a single round of PCR (35) .
3.5. Conclusion
The results of gel electrophoresis and blot hybridization analyses are used to
determine whether the specific PCR assay is positive (i.e., whether the pres-ence of the translocation fusion mRNA in question has been detected). The
clarity and strength of specific oligoprobe hybridization, as well as the
molecular size of the PCR product, are considered to establish a result as posi-tive. The absence of appropriately sized PCR products and lack of specific
probe hybridization denote negative results. Concomitant demonstration of
intact and amplifiable control RNA (cDNA) is always required in order to
exclude a sample or technical failure, particularly in the case of a negative
PCR analysis. In the latter case, amplification of a segment of `2-microglobulin
RNA (cDNA) is commonly utilized.
Although an in-depth discussion is beyond the scope of this chapter, the
choice of RT-PCR control(s) is important. I have illustrated `2-microglobulin
herein, because it is relatively well expressed in hematopoietic cells and has
proven to be informative in most cases. However, more substantial guidelines
are emerging regarding this issue (36) . In particular, care should be taken to
exclude amplification of genomic DNA (e.g., genes with short intron segments
spanning the area amplified), control gene products should be relatively unique
and lack pseudogenes, and the expression level of the chosen control gene
should not be greater than that of the target gene. Notably, high-purity isola-tion and proper storage (at least –80°C) of high-quality RNA from viable, fresh,
or well-preserved cells is essential; sensitivity of the RT-PCR assay can be
significantly compromised in poorly preserved material. Although this may be
somewhat less critical in examples with abundant target mRNA and short
amplicon length (e.g., BCR-ABL fusion in chronic myeloid leukemia), detec-tion of other labile or low-abundance fusion mRNAs may be adversely
affected (e.g., PML-RAR_ fusion in acute promyelocytic leukemia).
138 Viswanatha
Finally, brief mention is made here regarding the sensitivity of RT-PCR
assays of leukemic fusion genes, in the context of monitoring MRD. In my
experience, the sensitivity of the inv(16)/CBF`-MYH11 and t(8;21)/AML1-ETO RT-PCR assays is typically 1 in 104–105 cells. Further enhancements can
be gained by the use of radiolabeled PCR primers or probes, although the clini-cal utility of this goal is uncertain. Although the reported data are not yet clear
concerning the presence of MRD and risk of relapse in inv(16)/CBF`-MYH11
leukemia (37) , low-level t(8;21)/AML1-ETO fusion transcript is commonly
detected in patients with persistent long-term clinical remissions (38–40) . By
contrast, RT-PCR for the t(15;17)/PML-RAR_has a sensitivity of one or two
orders of magnitude less than for the other described RT-PCR assays. This
likely reflects a combination of factors, of which mRNA instability and rela-tively low transcript abundance are important. However, this relatively lower
level of sensitivity in detection of PML-RAR_MRD is of substantial clinical
prognostic significance and can be utilized for prediction of impending disease
relapse (41 ,42) . An emerging paradigm for the management of acute leukemic
patients is the realization that despite attainment of clinical remission, residual
disease persists and can be detected by sensitive molecular methods. The
development of novel technologies such as the fluorescence-based automated
quantitative real-time PCR (e.g., ABI 7700 TaqMan system [Applied Bio-systems, Foster City, CA] and LightCycler system [Roche Diagnostics,
Indianapolis, IN]) now permits extremely accurate and reproducible measure-ments of specific molecular disease markers over a dynamic range of 105–106.
Application of this technology in our laboratory has revealed dramatic fluctua-tions in the level of residual PML-RAR_positive clonal leukemic cells over
time in patients treated for acute promyelocytic leukemia, with increasing lev-els of fusion transcript heralding impending overt relapse. Similar data are also
emerging for other leukemia-associated fusion translocations (43–45) . Thus, it
appears that both the quantitative (“threshold”) level of residual leukemia and
the temporal behavior of the clonal population are determinants of disease
activity and potential for relapse. Automated quantitative PCR monitoring of
patients will likely play an increasing role in this context.
4. Notes
1. RNA quality and recovery are vital to the success of RT-PCR reactions, particu-larly if sensitive MRD studies are also being performed. The integrity of RNA is
also especially important for transcripts such as the t(15;17)-associated  PML-RAR_ fusion, which are relatively labile and in low abundance. We have had
excellent results with proprietary complete RNA isolation kits, such as those from
Ambion or Qiagen (Santa Clarita, CA) and have opted to utilize these in favor of
standard guanidinium-based isolation methods. Despite an increase in cost over
Anomalies in Acute Myeloid Leukemias 139
more traditional methods, the consistency in RNA quality and yield is better
suited to meet the stringent quality assurance issues required in a clinical labora-tory and to provide RNA sufficient for routine monitoring of MRD.
2. All stock solutions utilized in reverse transcription and PCR reactions are pre-pared with DEPC-treated double-distilled water and are stored or manipulated in
similarly treated, autoclaved, and dedicated glassware.
3. Stock solution tubes of 10X RT buffer, 10X PCR buffer, and 10 mM dNTPs are
made in quantity and labeled with a specific lot number. New reagent tubes are
tested prior to use in actual assays. The stock buffer and dNTP tubes are kept at
–20°C.
4. The Bio-Rad model 785 vacuum transfer system (Bio-Rad) is used in my labora-tory with reliable and excellent results. The setup parameters described in
Subheading 3. refer to this system and will vary slightly for different types of
apparatus.
5. Hybridization buffer can be made in quantity and aliquots stored at –20°C, then
removed, and warmed in a 60°C water bath when performing hybridization.
6. For positive controls, these sources can be obtained with appropriate permission
as required: the NB4 cell line for the t(15;17)/PML-RAR_ L-form (BCR-1)
fusion (46) ; the UF-1 cell line for the t(15;17)/PML-RAR_ S-form (BCR-3)
fusion (47) ; the Kasumi-1 cell line for the t(8;21)/AML1-ETO fusion (48) ; and a
cytogenetically confirmed anonymous patient sample for the inv(16)/t(16;16)/
CBF`-MYH11 anomaly (type A fusion). Finally, in addition to positive control
reactions, negative controls (blanks) containing everything except nucleic acid
template should be routinely utilized to monitor for contamination.
7. Because PCR techniques are generally quite robust and capable of detecting small
amounts of initial nucleic acid target, control of potential contamination during
every step of the assay is paramount. My laboratory utilizes separate areas for
sample preparation (RNA and DNA isolation), PCR set-up and gel analysis/
hybridization. Analyzed PCR products (post-PCR) should never be returned to
or circulated through PCR setup, and nucleic acid isolation areas. Nucleic acid
isolation and PCR stock and working solutions are kept separate from all other
laboratory reagents. Positive displacement micropipets (e.g., Rainin Microman,
Rainin, Emeryville, CA) and sterile, RNase-free, barrier-type, disposable pipet
tips are used throughout. Separate dedicated pipets are utilized in the nucleic acid
isolation and PCR areas.
8. The red blood cell lysis method is less favored owing to the presence of residual
heme, which can inhibit PCR efficiency.
9. The reverse transcription and PCR steps are performed in thin-walled 100-µL
MicroAmp tubes (Perkin-Elmer), for use in a Perkin-Elmer model 9600 or 9700
thermal cycler, or equivalent instrument. Mineral oil overlay is avoided with this
approach. Other instruments may require different PCR sample tubes and possi-bly addition of an oil layer before PCR.
10. For ease of analysis, I add 15  µL of 10 mg/mL of EtBr directly to the warm,
prepoured gel.
140 Viswanatha
11. If `2-microglobulin PCR products are on a separate gel, or are clearly separated
from the fusion gene PCR products (e.g., on the other half of a double-comb gel),
these are not transferred to the nylon membrane. If they are on the same part of
the gel, the `2-microglobulin PCR products are transferred along with the fusion
gene PCR products; the section of membrane containing `2-microglobulin PCR
products is then separated and discarded prior to probe hybridization.
12. Following the specific probe hybridization and warm blot wash steps, several
blots can be combined and processed together in the SA-HRP and subsequent
room temperature wash steps.
13. During the SA-HRP and room temperature wash steps, the plastic tubs can be
kept loosely covered to prevent possible precipitation of the SSPE/SDS solution.
Precipitation can occasionally occur in laboratories with lower ambient temperature.
14. Typical exposure times are between 5 and 15 min; however, strong signals are
often present on film within 1–3 min. The chemiluminescent output decays rap-idly after a few hours.
15. The type of  PML-RAR_ fusion in APL does not appear to be associated with
clinical outcome. Some previous studies have suggested a more aggressive course
for S-form (BCR-3) positive tumors  (49 ,50) . However, the results of a large
Intergroup Cooperative trial (51) did not confirm a significant independent asso-ciation of this fusion type. Thus, specific chimeric PML-RAR_mRNA types do
not appear to have prognostic value in APL. However, identification of the tran-script type in individual patients is important to serve as a marker in MRD stud-ies. We have also encountered rare situations in which different  PML-RAR_
transcript types may coexist at diagnosis (e.g., both L- and S-forms), or change
from one type to another over time in follow-up samples (personal observations).
16. The three-band pattern observed in first-round L- and V-form PML-RAR_amplifica-tion products arises from alternative splicing. The largest (uppermost) PCR fragment
represents the full-length fusion of  PML exon 6 to  RAR_ exon 3. The two lower
molecular weight bands occur owing to alternative splicing out of PML exon 5 and
exons 5 and 6, respectively. The upper two bands of V-form (BCR-2) cases typically
are of smaller size compared to the L-form fusion product, owing to the variable
break point–fusion events situated within PML exon 6; however, rare V-form cases
may demonstrate larger or similarly sized PCR bands. For any of these V-form cases,
no hybridization signal is expected with the L-form fusion junction oligoprobe.
17. Nested allele-specific PCR  (34) to confirm the presence of the type A  CBF`-MYH11 fusion mRNA is usually performed in our laboratory. Following first-round PCR, transfer 5 µL from each of the C1-M1 product tubes to a new set of
labeled MicroAmp PCR tubes. Assemble the second-round CBF`-MYH11 mas-ter mix in a 1.5-mL microfuge tube, on ice, as follows: 36.5 µL of DEPC H2O,
5 µL of 10X PCR buffer, 1  µL of 10 mM dNTPs, 1  µL each of second-round
primers C2 (sense) and M7 (allele-specific antisense), 0.5 µL of Taq DNA poly-merase (see Fig. 1B for location of nested primers). This total of 45 µL of PCR
master mix is per sample and needs to be multiplied by the number of samples
being tested, plus one extra (n + 1). Add 45 µL of second-round CBF`-MYH11
Anomalies in Acute Myeloid Leukemias 141
master mix to each MicroAmp tube and mix the contents of each tube carefully
(resulting in a total volume of 50 µL per reaction). Place the tubes in the thermal
cycler, programmed as follows: 95°C for 3 min (initial denaturation); 25 cycles
of 95°C for 30 s, 57°C for 30 s, and 72°C for 60 s; then 72°C for 5 min (final
extension) and cool to 4°C. The C2-M7 PCR products are 65 bp in size on stan-dard gel electrophoresis. This method will specifically confirm the presence of
the type A CBF`-MYH11 transcript and extends the sensitivity of detection of
MRD for this particular fusion by one or two orders of magnitude over first-round (C1-M1) PCR alone in cell dilution experiments (personal observations).
18. As a cautionary note, a rare  CBF`-MYH11 fusion PCR product with a size
almost indistinguishable from the type A product has been identified (52) , result-ing from novel fusion sites in both CBF`and MYH11 genes. This PCR product
did not amplify with the allele-specific PCR (Note 17) and did not hybridize with
the internal CBF`or the inv(16) junction A oligoprobes. Thus, although the use
of probe hybridization is generally recommended, DNA sequencing of the PCR
product is reserved for rare cases with unusual break points in the CBF`gene.
References
1. Bennett, J. M., Catovsky, D., Daniel, M. T., Flandrin, G., Galton, D. A., Gralnick,
H. R., and Sultan, C. (1985) Proposed revised criteria for the classification of
acute myeloid leukemia: a report of the French-American-British Cooperative
Group. Ann. Intern. Med. 103, 620–625.
2. Head, D. R. (1996) Revised classification of acute myeloid leukemia. Leukemia
10, 1826–1831.
3. Grignani, F., Fagioli, M., Alcalay, M., Longo, L., Pandolfi, P. P., Bonti, E., Biondi,
A., LoCoco, F., Grignani, F., and Pelicci, P. G. (1994) Acute promyelocytic leu-kemia: from genetics to treatment. Blood 83, 10–25.
4. Dyck, J. A., Maul, G. G., Miller, W. H. Jr., Chen, J. D., Kakizuka, A., and Evans,
R. M. (1994) A novel macromolecular structure is a target of the promyelocyte-retinoic acid receptor oncoprotein. Cell 76, 333–343.
5. Weism K., Rambaud, S., Lavau, C., Jansen, J., Carvalho, T., Carmo-Fonseca, M.,
Lamond, A., and Dejean, A. (1994) Retinoic acid regulates aberrant nuclear local-ization of PML-RAR_ in acute promyelocytic leukemia cells. Cell 76, 345–356.
6. Lamond, A. I. and Earnshaw, W. C. (1998) Structure and function in the nucleus.
Science 280, 547–553.
7. Wang, Z. G., Delva, L., Gaboli, M., Rivi, R., Giorgio, M., Cordon-Cardo, C.,
Grosveld, F., and Pandolfi, P. P. (1998) Role of PML in cell growth and the
retinoic acid pathway. Science 279, 1547–1551.
8. Quignon, F., De Bels, F., Koken, M., Feunteun, J., Ameisen, J. C., and de The, H.
(1998) PML induces a novel caspase-independent death process. Nat. Genet. 20,
259–265.
9. Wang, Z. G., Ruggero, D., Ronchetti, S., Zhong, S., Gaboli, M., Rivi, R., and
Pandolfi, P. P. (1998) PML is essential for mulitple apoptotic pathways.  Nat.
Genet. 20, 266–272.
142 Viswanatha
10. Zheng, P., Guo, Y., Niu, Q., Levy, D. E., Dyck, J. A., Lu, S., Sheiman, L. A., and
Liu, Y. (1998) Proto-oncogene PML controls genes devoted to MHC class I anti-gen presentation. Nature 396, 373–376.
11. Degos, L., Dombret, H., Chomienne, C., Daniel, M. T., Micléa, J. M., Chasting,
C., Castaigne, S., and Fenaux, P. (1995) All-trans-retinoic acid as a differenti-ating agent in the treatment of acute promyelocytic leukemia.  Blood 85,
2643–2653.
12. Tallman, M. S., Andersen, J. W., Sciffer, C. A., Appelbaum, F. R., Feusner, J. H.,
Ogden, A., Shepherd, L., Willman, C., Bloomfield, C. D., Rowe, J. M., and
Wiernik, P. H. (1997) All-trans-retinoic acid in acute promyelocytic leukemia.
N. Engl. J. Med. 337, 1021–1028.
13. Kane, J. R., Head, D. R., Balazs, L., Hulshof, M. G., Motroni, T. A., Raimondi,
S. C., Carroll, A. J., Behm, F. G., Krance, R. A., Shurtleff, S. A., and Downing,
J. R. (1996) Molecular analysis of the PML/RAR alpha chimeric gene in pediatric
acute promyelocytic leukemia. Leukemia 10, 1296–1302.
14. Liu, P., Tarlé, S. A., Hajra, A., Claxton, D. F., Marlton, P., Freedman, M.,
Siciliano, M. J., and Collins, F. S. (1993) Fusion between transcription factor
CBF` and a myosin heavy chain in acute myeloid leukemia.  Science 261,
1041–1044.
15. Liu, P. P., Hajra, A., Wijmenga, C., and Collins, F. S. (1995) Molecular pathogen-esis of the chromosome 16 inversion in the M4Eo subtype of acute myeloid leuke-mia. Blood 85, 2289–2302.
16. Nucifora, G. and Rowley, J. D. (1995) AML1 and the 8;21 and 3;21 translocations
in acute and chronic myeloid leukemia. Blood 86, 1–14.
17. Daga, A., Tighe, J. E., and Calabi, F. (1992) Leukemia/Drosophila homology.
Nature 356, 484.
18. Erickson, P., Gao, J., Chang, K. S., Look, T., Whisenant, E., Raimondi, S., Lasher,
R., Trujillo, J., Rowley, J. D., and Drabkin, H. (1992) Identification of breakpoints
in t(8;21) AML and isolation of a fusion transcript with similarity to Drosophila
segmentation gene runt. Blood 80, 1825.
19. Shurtleff, S. A., Buijs, A., Behm, F. G., Rubnitz, J. E., Raimondi, S. C., Hancock,
M. L., Chan, CG. F., Pui, C. H., Grosveld, G., and Downing, J. R. (1995) TEL/
AML1 fusion resulting from a cryptic t(12;21) is the most common genetic lesion
in pediatric ALL and defines a subgroup of patients with excellent prognosis.
Leukemia 9, 1985–1989.
20. Borkhardt, A., Cazzaniga, G., Viehmann, S., Valsecchi, M. G., Ludwig, W. D.,
Burci, L., Mangioni, S., Schrappe, M., Riehm, H., Lampert, F., Basso, G., Masera,
G., Harbott, J., and Biondi, A. (1997) Incidence and clinical relevance of TEL/
AML1 fusion genes in children with acute lymphoblastic leukemia enrolled in the
German and Italian multicenter therapy trials. Blood 90, 571–577.
21. Wang, Q., Stacy, T., Miller, J. D., Lewis, A. F., Gu, T. L., Huang, X., Bushweller,
J. H., Bories, J. C., Alt, F. W., Ryan, G., Liu, P. P., Wynshaw-Boris, A., Binder,
M., Marín-Padilla, M., Sharpe, A. H., and Speck, N. A. (1996) The CBF`subunit
is essential for CBF_2 (AML1) function in vivo. Cell 87, 697–708.
Anomalies in Acute Myeloid Leukemias 143
22. Wang, Q., Stacy, T., Binder, M., Marín-Padilla, M., Sharpe, A. H., and Speck,
N. A. (1996) Disruption of the Cbfa2 gene causes necrosis and hemorrhaging in
the central nervous system and blocks definitive hematopoiesis. Proc. Natl. Acad.
Sci. USA 93, 3444–3449.
23. Castilla, L. H., Wijmenga, C., Wang, Q., Stacy, T., Speck, N. A., Eckhaus, M.,
Marín-Padilla, M., Collins, F. S., Wynshaw-Boris, A., and Liu, P. P. (1996) Fail-ure of embryonic hematopoiesis and lethal hemorrhages in mouse embryos het-erozygous for a knocked-in leukemia gene CBFB-MYH11. Cell 87, 687–696.
24. Okuda, T., Cai, Z., Yang, S., Lenny, N., Lyu, C. J., van Deursen, J. M., Harada,
H., and Downing, J. R. (1998) Expression of a knocked-in AML1-ETO leukemia
gene inhibits the establishment of normal differentiative hematopoiesis and
directly generates dysplastic hematopoietic progenitors. Blood 91, 3134–3143.
25. Van de Locht, L. T., Smetsers, T. F., Wittebol, S., Raymakers, R. A., and Mensink,
E. J. (1994) Molecular diversity in AML1/ETO fusion transcripts in patients with
t(8;21) positive acute myeloid leukaemia. Leukemia 8, 1780–1784.
26. Lin, R. J., Nagy, L., Inoue, S., Shao, W., Miller, W. H. Jr., and Evans, R. M.
(1998) Role of the histone deacetylase complex in acute promyelocytic leukaemia.
Nature 391, 811–814.
27. Grignani, F., De Matteis, S., Nervi, C., Tomassoni, L., Gelmetti, V., Cioce, M., Fanelli,
M., Ruthardt, M., Ferrara, F. F., Zamir, I., Seiser, C., Grignani, F., Lazar, M. A.,
Minucci, S., and Pelicci, P. G. (1998) Fusion proteins of the retinoic acid receptor-_
recruit histone deacetylase in promyelocytic leukaemia. Nature 391, 815–818.
28. Guidez, F., Ivins, S., Zhu, J., Söderström, M., Waxman, S., and Zelent, A. (1998)
Reduced retinoic acid-sensitivities of nuclear receptor corepressor binding to
PML- and PLZ-RAR_ underlie molecular pathogenesis and treatment of acute
promyelocytic leukemia. Blood 91, 2634–2642.
29. Wang, J., Hoshino, T., Redner, R. L., Kajigaya, S., and Liu, J. M. (1998) ETO,
fusion partner in t(8;21) acute myeloid leukemia, represses transcription by inter-action with the human N-COR/mSIN3/HDAC1 complex. Proc. Natl. Acad. Sci.
USA 95, 10,860–10,865.
30. Lutterbach, B., Westendorf, J. J., Linggi, B., and Hiebert, S. W. (1998) AML-1, a
target of multiple chromosomal translocations in acute leukemia, interacts with
mSIN3 to repress transcription from the  p21WAF1/CIP1 promoter. Blood
92(Suppl.), 508a.
31. Gallagher, R. E., Li, Y. P., Rao, S., Paietta, E., Andersen, J., Etkind, P., Bennett,
J. M., Tallman, M. S., and Wiernik, P. H. (1995) Characterization of acute
promyelocytic leukemia cases with PML-RAR_break/fusion sites in PML exon
6: identification of a subgroup with decreased in vitro responsiveness to all-trans
retinoic acid. Blood 86, 1540–1547.
32. Downing, J. R., Head, D. R., Curcio-Brint, A. M., Hulshof, M. G., Motroni, T. A.,
Raimondi, S. C., Carroll, A. J., Drabkin, H. A., Willman, C., Theil, K. S., Civin,
C. I., and Erickson, P. (1993) An  AML1/ETO fusion transcript is consistently
detected by RNA-based polymerase chain reaction in acute myelogenous leuke-mia containing the (8;21)(q22;q22) translocation. Blood 81, 2860–2865.
144 Viswanatha
33. Shurtleff, S. A., Meyers, S., Hiebert, S. W., Raimondi, S. C., Head, D. R.,
Willman, C. L., Wolman, S., Slovak, M. L., Carroll, A. J., Behm, F., Hulshof,
M. G., Motroni, T. A., Okuda, T., Liu, P., Collins, F. S, and Downing, J. R. (1995)
Heterogeneity in CBF`/MYH11 fusion messages encoded by the inv(16)(p13q22)
and the t(16;16)(p13;q22) in acute myelogenous leukemia.  Blood 85,
3695–3703.
34. Hébert, J., Cayuela, J. M., Daniel, M. T., Berger, R., and Sigaux, F. (1994) Detec-tion of minimal residual disease in acute myelomonocytic leukemia with abnor-mal marrow eosinophils by nested polymerase chain reaction with allele specific
amplification. Blood 84, 2291–2296.
35. Barragan, E., Bonanad, S., Lopez, J. A., Bolufer, P., and Sanz, M. A. (1998) Com-parison of two reverse transcription-polymerase chain reaction methods for
detection of AML1/ETO rearrangement in the M2 subtype of acute myeloid leu-kaemia. Clin. Chem. Lab. Med. 36, 137–142.
36. Watzinger, F. and Lion, T. (1998) Multiplex PCR for quality control of template
RNA/cDNA in RT-PCR assays. Leukemia 12, 1930–1936.
37. Marcucci, G., Caligiuri, M. A., and Bloomfield, C. D. (1997) Defining the
“absence” of the CBF`/MYH11 fusion transcript in patients with acute myeloid
leukemia and inversion of chromosome 16 to predict long-term complete remis-sion: a call for definitions. Blood 90, 5022–5025 (letter).
38. Kusec, R., Laczika, K., Knöbl, P., Friedl, J., Greinix, H., Kahls, P., Linkesh, W.,
Schwarzinger, I., Mitterbauer, G., Purtscher, B., Hass, O. A., Lechner, K., and
Jaeger, U. (1994) AML1/ETO fusion mRNA can be detected in remission blood
samples of all patients with t(8;21) acute myeloid leukemia after chemotherapy or
autologous bone marrow transplantation. Leukemia 8, 735–739.
39. Jurlander, J., Caligiuri, M. A., Ruutu, T., Baer, M. R., Strout, M. P., Oberkircher,
A. R., Hoffmann, L., Ball, E. D., Frei-Lahr, D. A., Christiansen, N. P., Block, A.
W., Knuutila, S., Herzig, G. P., and Bloomfield, C. D. (1996) Persistence of the
AML1/ETO fusion transcript in patients treated with allogeneic bone marrow
transplantation for t(8;21) leukemia. Blood 88, 2183–2191.
40. Tobal, K. and Liu Yin, J. A. (1996) Monitoring of minimal residual disease by
quantitative reverse transcriptase-polymerase chain reaction for AML1-MTG8
transcripts in AML-M2 with t(8;21). Blood 88, 3704–3709.
41. Diverio, D., Rossi, V., Avvisati, G., De Santis, S., Pistilli, A., Pane, F., Saglio, G.,
Martinelli, G., Petti, M. C., Santoro, A., Pelicci, P. G., Mandelli, F., Biondi, A.,
and Lo Coco, F. (1998) Early detection of relapse by prospective reverse tran-scriptase-polymerase chain reaction analysis of the PML/RAR_ fusion gene in
patients with acute promyelocytic leukemia enrolled in the GIMEMA-AIEOP
multicenter “AIDA” trial. Blood 92, 784–789.
42. Diverio, D., Pandolfi, P. P., Rossi, V., Biondi, A., Pelicci, P. G., and Lo Coco, F.
(1994) Monitoring of treatment outcome in acute promyelocytic leukemia by
RT-PCR. Leukemia 8(Suppl. 2), S63–S65.
43. Marcucci, G., Livak, K. J., Bi, W., Strout, M. P., Bloomfield, C. D., and Caligiuri,
M. A. (1998) Detection of minimal residual disease in patients with AML1/ETO-
Anomalies in Acute Myeloid Leukemias 145
associated acute myeloid leukemia using a novel quantitative reverse transcrip-tion polymerase chain reaction assay. Leukemia 12, 1482–1489.
44. Zhao, S., Xu, H., Xie, Z., Chang, S., Konopleva, M., Kavka, K., Calvert, L., Guo,
J. Q., Arlinghaus, R., Siciliano, M., and Andreef, M. (1998) Preliminary compari-son of real-time QPCR with cytogenetic karyotype, hypermetaphase FISH and
western blot analysis in CML patients in clinical remission.  Blood 92(Suppl.),
72a.
45. Mensink, E., van de Locht, A., Schattenberg, A., Linders, E., Schaap, N., Geurts
van Kessel, A., De Witte, T. (1998) Quantitation of minimal residual disease in
Philadelphia chromosome positive chronic myeloid leukaemia patients using real-time quantitative RT-PCR. Br. J. Haematol. 102, 768–774.
46. Lanotte, M., Martin-Thouvenin, V., Najman, S., Balerini, P., Valensi, F., and
Berger, R. (1991) NB4, a maturation inducible cell line with t(15;17) marker iso-lated from a human acute promyelocytic leukemia (M3). Blood 77, 1080–1086.
47. Kizaki, M., Matsushita, H., Takayama, N., Muto, A., Ueno, H., Awaya, N., Kawai,
Y., Asou, H., Kamada, N., and Ikeda, Y. (1996) Establishment and characteriza-tion of a novel acute promyelocytic leukemia cell line (UF-1) with retinoic acid-resistant features. Blood 88, 1824–1833.
48. Asou, H., Tashiro, S., Hamamoto, K., Otsuji, A., Kita, K., and Kamada, N. (1991)
Establishment of a human acute myeloid leukemia cell line (Kasumi-1) with
t(8;21) chromosome translocation. Blood 77, 2031–2036.
49. Huang, W., Sun, G. L., Li, X. S., Cao, Q., Lu, Y., Jang, G. S., Zhang, F. Q., Chai,
J. R., Wang, Z. Y., Waxman, S., Chen, Z., and Chen, S. J. (1993) Acute
promyelocytic leukemia: clinical relevance of two major PML-RAR_isoforms
and detection of minimal residual disease by retro-transcriptase/polymerase chain
reaction to predict relapse. Blood 82, 1264.
50. Vahdat, L., Maslak, P., Miller, H. W., Eardley, A., Heller, G., Scheinberg, D. A.,
and Warrell, R. P. Jr. (1994) Early mortality and the retinoic acid syndrome in
acute promyelocytic leukemia: impact of leukocytosis, low-dose chemotherapy,
PMN/RAR-_ isoform, and CD13 expression in patients treated with all-trans
retinoic acid. Blood 84, 3843–3849.
51. Gallagher, R. E., Willman, C. L., Slack, J. L., Andersen, J. W., Li, Y. P.,
Viswanatha, D., Bloomfield, C. D., Appelbaum, F. R., Schiffer, C. A., Tallman,
M. S., and Wiernik, P. H. (1997) Association of PML-RAR_fusion mRNA type
with pretreatment hematologic characteristics but not treatment outcome in acute
promyelocytic leukemia: an intergroup molecular study. Blood 90, 1656–1663.
52. Viswanatha, D. S., Chen, I. M., Liu, P. P., Slovak, M. L., Rankin, C., Head, D. R.,
and Willman, C. L. (1998) Characterization and use of an antibody detecting the
CBF`-SMMHC fusion protein in inv(16)/t(16;16)-associated acute myeloid leu-kemias. Blood 91, 1882–1890.
Gene Fusion in Non-Hodgkin Lymphoma 147
13
Detection of t(14;18)(q32;q21)-Associated
BCL-2/JH Gene Fusion in Non-Hodgkin Lymphoma
David S. Viswanatha
1. Introduction
The identification and study of nonrandom recurrent chromosomal translo-cations has substantially increased our understanding of the non-Hodgkin lym-phomas. Cytogenetic and molecular genetic data now form an integral part of
current lymphoma classifications  (1) and provide important information for
diagnosis, tumor biology, and in some cases prognosis. The t(14;18)(q32;q21)
abnormality is the most common translocation detected in B-lineage lymphoma
and results in juxtaposition of the BCL-2 gene (18q21) and the JH locus of the
immunoglobulin (Ig) heavy chain gene (14q32)  (2–5) . More specifically, in
the North American population, alterations of the BCL-2 gene are detected in
approx 75 to 85% of low-grade follicular lymphomas, 20–30% of aggressive
large B-cell lymphomas, and rarely in other B-cell tumors (e.g., chronic lym-phocytic leukemia (CLL), acute lymphoblastic leukemia) (2 ,6–9) . As a conse-quence of the BCL-2/JH fusion, deregulated overexpression of the antiapoptotic
bcl-2 protein occurs owing to constitutive transcriptional activation of the BCL-2
gene by the Ig heavy chain gene enhancer. The unbridled expression of bcl-2
protein in lymphoid tumors confers resistance to programmed cell death (10 ,11)
and is implicated in primary therapeutic failure and a less favorable prognosis
(12–14). Although karyotypic detection of lymphoma-associated translocations
such as the t(14;18) has proved to be useful in disease diagnosis and
subcategorization, molecular genetic approaches including polymerase chain
reaction (PCR) and fluorescence in situ hybridization (FISH) have gained sub-stantial popularity owing to their rapidity, relatively low cost, and increased
sensitivity (6,15–22) .
147
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
148 Viswanatha
The partial genomic structures of the BCL-2 and Ig heavy chain genes are
illustrated schematically in Fig. 1. In the case of the Ig heavy chain gene locus,
the vast majority of breakpoints arising from the t(14;18) are distributed within
the joining region (JH ) introns. The largest proportion (60%) of break-fusion
sites in the BCL-2 gene occurs in a tightly clustered area in the 3vnoncoding
segment of exon 3, known as the major breakpoint region (MBR) (3) , whereas
a small number (15–20%) involves a second clustered region called the minor
cluster region (mcr) (23 ,24) . Rarely, a 5v region of  BCL-2 called the variant
Fig. 1. Schematic diagrams of BCL-2 and immunoglobulin JH gene regions. (Top)
BCL-2 gene. Exons are rectangular boxes (I–III). Gray shading indicates noncoding
exon regions. Relative locations of principal breakpoint regions are indicated by verti-cal arrows: VCR, variant cluster region; MBR, major breakpoint region; mcr, minor
cluster region. The MBR is situated in the 3 noncoding area of  BCL-2 exon 3 and
encompasses a tightly clustered segment of approx 150 bp. Positions of MBR and mcr
locus primers for PCR are indicated by short horizontal arrows. Oligonucleotide probes
for MBR and mcr rearrangement detection are situated internal to the respective prim-ers (not shown). (Bottom) Immunoglobulin heavy chain gene  JH locus. The six  JH
exons are shown as rectangular boxes. Breakpoints can be distributed in any of the
JH intron areas. Consensus JH primer for PCR is illustrated by short horizontal arrows
(one primer, complementary to conserved JH sequence).
Gene Fusion in Non-Hodgkin Lymphoma 149
cluster region (VCR) is rearranged in B-cell CLL; typically, the VCR locus is
involved in translocations with the Ig light chain genes (25 ,26) . As a result of
the relatively tight clustering of genomic breakpoints in the MBR and mcr loci
of the BCL-2 gene, PCR techniques can be employed to detect the majority of
t(14;18)-associated BCL-2/JH fusions. This strategy, depicted in Fig. 1, can be
accomplished using MBR and mcr oligonucleotide primers placed upstream of
the clustered regions, in combination with a consensus JH primer.
The identification of BCL-2 gene rearrangements in lymphoid proliferations
is quite useful in several clinical contexts including the diagnosis and subclas-sification of non-Hodgkin lymphoma. Demonstration of clonality represents
an important means of confirming the diagnosis of lymphoma in cases in which
morphologic and phenotypic methods have been inconclusive. Clonal Ig gene
rearrangements are often not detected by standard PCR techniques in follicular
center cell lymphomas  (27) . However, the  BCL-2/JH abnormality is readily
detected by PCR, providing an alternative clonal marker for distinguishing fol-licular lymphoma from an atypical or florid but benign follicular hyperplasia.
As a corollary, PCR analysis for the BCL-2/JH fusion can be used to differenti-ate subtypes of non-Hodgkin lymphoma with similar morphologic features,
such as follicular lymphoma versus lymphomas of mantle or marginal zone
type. As knowledge of lymphoma biology continues to broaden, the recogni-tion of specific entities based on molecular genetic criteria will likely form an
integral aspect of “tumor-directed” therapies. Finally, sensitive PCR detection
of the BCL-2/JH fusion can serve as a patient-specific marker for monitoring of
minimal residual disease (MRD) in individuals undergoing intensive chemo-therapy or bone marrow transplantation for follicular lymphoma; this aspect is
discussed briefly in Subheading 3.6.
2. Materials
2.1. Isolation of DNA
2.1.1. Isolation of DNA from Cell Pellets and Fresh or Frozen Tissues
1. DNA isolation kit (Puregene; Gentra Systems, Minneapolis MN) (see Note 1).
2. Proteinase K solution (20 µg/µL) (Puregene; Gentra Systems).
3. RNase A solution (4 µg/µL) (Puregene; Gentra Systems).
4. Isopropanol, reagent grade (EM Science, Gibbstown, NJ).
5. Cold 70% ethanol.
2.1.2. Isolation of DNA from Fixed, Paraffin-Embedded Tissues
1. Xylene (Fisher, Fair Lawn, NJ).
2. Absolute ethanol (Midwest Grain Products, Pekin, IL).
3. QIAamp DNA Mini kit (Qiagen, Santa Clarita, CA) (see Note 1).
4. Proteinase K solution (approx 18 µg/µL) (Qiagen).
150 Viswanatha
2.2. PCR for BCL-2/JH Fusion
1. 10 mM stock dNTPs (Perkin-Elmer, Foster City, CA) (see Note 2).
2. 10X PCR buffer (Perkin-Elmer): 100 mM Tris-HCl (pH 8.4), 500 mM KCl,
15 mM MgCl2.
3. Sense and antisense primers (15–20 pmol/µL of each) (see Table 1).
4. AmpliTaq DNA polymerase (5 U/µL) (Perkin-Elmer/Roche, Foster City, CA).
5. Diethylpyrocarbonate (DEPC) distilled H2O.
2.3. Gel Analysis of PCR Product and Vacuum Blot Transfer
1. Standard powdered agarose for 1.5% agarose gel (3g) (SeaKem ME, FMC
Bioproducts, ME).
2. Ethidium bromide (EtBr) solution (10 mg/mL) (Gibco-BRL/Life Technologies,
Gaithersburg, MD).
3. Stock 10X Tris-borate EDTA (TBE) buffer solution: 107.8 g of Tris-HCl, 55 g of
boric acid, 7.4 g of Na2EDTA per 1 L of sterile distilled H2O (0.5X TBE is used
for gel and 1X TBE for running buffer).
4. 10X Sample loading dye: 1 vol of 1% bromophenol blue, 1 vol of 1% xylene
cyanol, 2 vol of glycerol.
5. Horizontal gel box (40 cm) with 20-well combs (BRL Horizon 20.25; Gibco-BRL/Life Technologies).
6. UV light box and instant camera setup with Polaroid 667 B+W film (Polaroid,
Cambridge, MA).
7. Biodyne B nylon membrane (11  × 20 cm) (Pall-Biodyne, East Hills, NY) to
encompass area of gel being transferred.
8. Two clean plastic tubs.
9. Dilute (0.05 N ) HCl (500 mL).
10. 0.4 M NaOH (1 L).
Table 1
Primer and Probe Sequences for BCL-2/JH PCR Analysis
Primer or Probe Sequence (5vA 3v)a
MBR CCAAGTCATGTGCATTTCCACGTC
mcr ACAGCGTGGTTAGGGTTAGGTCGTA
JH ACCTGAGGAGACGGTGACC
`-Globin I GGTTGGCCAATCTACTCCCAGG
`-Globin II GCTCACTCAGTGTGGCAAAG
MBR probe *TAGAGAGTTGCTTTACGTGGCCTG
mcr probe *AGTGCCTGGCATAGAGCAAG
aAn asterisk denotes 5v-biotin label for use in chemiluminescent probe hybridization and
detection.
Gene Fusion in Non-Hodgkin Lymphoma 151
11. Bio-Rad Model 785 vacuum transfer apparatus with vacuum regulator (Bio-Rad,
Hercules, CA) (see Note 3).
12. UV crosslinking apparatus (UV Stratalinker; Stratagene, La Jolla, CA).
2.4. Nonisotopic Probe Hybridization
1. Stock 20X SSPE buffer: 210.4 g of NaCl, 27.6 g of NaH2PO4 · H2O, 4.4 g of
NaOH, 7.4 g of Na2EDTA per 1 L of sterile distilled H2O; the buffer usually
requires gentle heat with constant stirring to completely solubilize. From this
prepare the following:
a. Prehybridization solution (100 mL): 0.1X SSPE/0.5% sodium dodecyl sul-fate (SDS) (from 10% SDS stock solution).
b. Blot wash solution (2 L): 2X SSPE/0.1% SDS.
c. 50-mL Aliquots of hybridization buffer: 5X SSPE/0.1% SDS (see Note 4).
2. 5v-Biotinylated oligonucleotide probes (10–15 pmol/µL) (see Table 1).
3. Heat-sealable plastic hybridization bags.
4. Several clean plastic tubs with lids.
5. Streptavidin-horseradish peroxidase (SA-HRP) (1mg/mL) (Vector, Burling-ame, CA).
6. ECL chemiluminescence detection reagent solutions 1 and 2 (Amersham
Pharmacia Biotech, Piscataway, NJ).
7. Fluorescent marking pen.
8. Standard filter (blotting) paper.
9. Plastic wrap.
10. Radiographic film (Kodak XAR; Eastman-Kodak, Rochester, NY).
3. Methods
The PCR method detailed is based on a previously described protocol (6) ,
optimized for use in my laboratory. General laboratory principles pertaining to
the performance of PCR and analysis of PCR products should be followed (see
Note 5).
3.1. Isolation of DNA
3.1.1. Isolation of DNA from Cell Pellets and Fresh or Frozen Tissues
1. Obtain a cell pellet (of blood or bone marrow) or minced tissue sample in a
15-mL conical centrifuge tube.
2. Follow manufacturer’s general guidelines for the Puregene kit, for cell pellets
(10–20 × 106 cells) or tissue samples (10–20 mg) (see Note 6).
3. Resuspend the DNA pellet in a volume of DNA hydration buffer (e.g., 1X TE
or equivalent) to give an approximate concentration of 1 µg/µL. Hydration may
be accomplished overnight at ambient temperature for large sample isolates.
Calculate the concentration and yield by standard 260-nm UV absorbance
spectrophotometry.
4. Store the DNA samples at –20°C.
152 Viswanatha
3.1.2. Isolation of DNA from Fixed, Paraffin-Embedded Tissues
1. Obtain three to four 10-µm thick sections of paraffin-embedded tissue. Gener-ally, only two of these are required for isolation and the remaining material can
be kept for reisolation if the initial yield or PCR is inadequate. Place the tissue
sections in a 1.5-mL microfuge tube (see Note 7).
2. Add 1 mL of xylene to the tube and vortex vigorously to mix well. Centrifuge in
a tabletop microcentrifuge on the highest setting for 5 min (see Note 8). Care-fully draw off the xylene and discard in an appropriate organic waste container.
3. Add 1 mL of absolute ethanol, vortex, and centrifuge on highest setting for
5 min. Draw off and discard the ethanol into an appropriate waste container.
Repeat this step once more.
4. Follow the manufacturer’s directions for the Qiagen QiaAmp DNA Mini kit, for
fixed, paraffin-embedded tissues.
5. Calculate the concentration and yield by standard 260-nm UV absorbance
spectrophotometry.
6. Store the DNA samples at –20°C.
3.2. PCR for MBR and mcr Region BCL-2/JH Rearrangements
1. Label three sets of MicroAmp tubes for the samples being tested (see Note 9),
positive controls (see Note 10), and negative control (blank—no DNA). One set
of tubes is for the MBR-JH PCR, one for the mcr-JH PCR, and one for the control
amplification of the `-globin gene. Similarly, label three 1.5-mL microfuge tubes
for each of the PCR master mix solutions.
2. Thaw the DNA tubes and keep on ice. Aliquot 1  µg of DNA into respective
MicroAmp tubes, proceeding in order of the sample tubes and then the positive
control. Do not add DNA template to the negative control (blank) tubes. Adjust the
volume of each tube to 5 µL with sterile DEPC H2O. Keep on ice or in a cold block.
3. Assemble the PCR master mixes in the corresponding 1.5-mL microfuge tubes,
on ice, as follows:
a. For MBR-JH: 36.5 µL of DEPC H2O, 5 µL of 10X PCR buffer, 1 µL of 10 mM
dNTPs, 1 µL each of primers MBR (sense) and JH (antisense), 0.5 µL of Taq
DNA polymerase.
b. For mcr-JH: 36.5 µL of DEPC H2O, 5 µL of 10X PCR buffer, 1 µL of 10 mM
dNTPs, 1 µL each of primers mcr (sense) and JH (antisense), 0.5 µL of Taq
DNA polymerase.
c. For  `-globin: 36.5  µL of DEPC H2O, 5  µL of 10X PCR buffer, 1  µL of
10 mM dNTPs, 1 µL each of primers `I (sense) and `II (antisense), 0.5 µL of
Taq DNA polymerase. (Note that this total 45-µL vol of each specific PCR
master mix is per sample and needs to be multiplied by the number of samples
being tested, plus one extra (n + 1). Gently vortex the tubes with lids closed to
mix the contents.)
4. Begin with the `-globin PCR tube set and add 45 µL of master mix to each of the
respective MicroAmp tubes (resulting in a total volume of 50 µL/tube). Mix the
contents of each tube gently and carefully. Proceed similarly, adding 45 µL of
Gene Fusion in Non-Hodgkin Lymphoma 153
MBR-JH and mcr-JH master mixes to each of the MicroAmp tubes in the respec-tive PCR sets, mixing the contents of tubes carefully (resulting in a total volume
of 50  µL/tube). In each PCR reaction set, proceed in the order from the test
samples to the positive control and finally the negative control.
5. Place all the tubes in a thermal cycler, programmed as follows: 95°C for 5 min
(initial denaturation); 2 cycles of 95°C for 30 s, 61°C for 15 s, and 72°C for 30 s;
then 30 cycles of 95°C for 15 s, 58°C for 15 s, and 72°C for 30 s; then 72°C for
5 min (final extension) and cool to 4°C. Following the PCR reaction, remove the
tubes from the thermal cycler and store the PCR products in a refrigerator or
–20°C freezer until ready for gel electrophoresis.
3.3. Gel Electrophoresis Analysis and Vacuum Blot
Transfer of BCL-2/JH PCR Products
1. Prepare a 1.5% agarose gel (with 0.5X TBE) (see Note 11).
2. In a microtiter plate, add 15 µL of PCR products to 2.5 µL of 10X sample loading
dye. A DNA size marker is also used, such as the 100-bp ladder (Gibco-BRL/
Life Technologies). Mix briefly and load the MBR-JH PCR products in one set of
lanes, followed by mcr-JH products, separating the two groups of PCR products
by two empty lanes. Load `-globin products in a separate set of lanes. Carry out
electrophoresis at 180 V for 1.5–2 h.
3. Following electrophoresis, observe and photograph the gel under UV illumina-tion. Proper eye protection is required when viewing gels by UV light. Standard
black-and-white Polaroid photographs are obtained using a photograph stand
(see Note 12).
4. Place the gel in a plastic tub and cover with 300–400 mL of dilute (0.05 M) HCl
(acid-nicking step). Place on a gyratory platform on gentle setting for 10 min.
During this time, allow the precut nylon membrane to equilibrate in 300–400 mL
of 0.4 M NaOH.
5. Decant the dilute HCl and add an equal amount of 0.4 M NaOH over the gel.
6. Assemble the vacuum transfer apparatus according to the manufacturer’s direc-tions. Align the gel over the nylon membrane, ensuring that the wells are outside
of the vacuum gasket seal. Slowly engage the vacuum to negative pressure of 4 to
5 mmHg. Pour approx 250 mL of 0.4 M NaOH over the gel to cover the area for
transfer. Transfer the gel products to the nylon membrane for 1 h.
7. Disassemble the vacuum transfer apparatus and remove the nylon membrane.
Place the membrane in a UV crosslinker and crosslink the DNA to the membrane
at a 500-µJ power setting. Label the well locations at the top of the gel using a
soft pencil and allow the membrane to dry.
8. Cut and separate sections containing MBR-JH and mcr-JH PCR products, label
each appropriately, and trim off the excess membrane. If `-globin products have
been transferred on the same membrane, cut away the area containing `-globin
product lanes and discard. The MBR-JH reaction products will be hybridized
with the internal sequence MBR oligoprobe whereas the mcr-JH products will be
hybridized with the internal sequence mcr oligoprobe.
154 Viswanatha
3.4. Nonisotopic (Chemiluminescent) Probe
Hybridization for BCL-2/JH Fusion
1. Warm to 60°C in a water bath 50 mL of prehybridization solution (0.1X SSPE/
0.5% SDS) and two 15-mL aliquots (one per separate hybridization reaction) of
hybridization buffer (5X SSPE/0.1% SDS).
2. Place nylon membranes of MBR-JH and mcr-JH PCR products in appropri-ately labeled, heat-sealable plastic bags and add 15 mL of prehybridization
buffer to each. Displace bubbles through the open end of the bag, heat-seal
the bag, and place in a 60°C oven, on a rocking platform. Allow 30 min for
prehybridization.
3. Add 5 µL of the biotinylated oligoprobes (MBR and mcr) to respective 15-mL
aliquots of warm hybridization buffer and gently mix (final concentration of
approx 1 to 2 pmol/mL of buffer). Remove and cut open the plastic bags across
the top. Drain the prehybridization buffer and carefully add 15 mL of the hybrid-ization buffer with the specific oligoprobe to the corresponding bags. Express
excess air and bubbles and heat-seal the bags just above the nylon membrane.
Check for leaks and place in a 60°C oven for 1 h. The hybridization reactions can
be extended to overnight for convenience.
4. Prepare 2 L of blot wash solution (2X SSPE/0.1% SDS). Aliquot 500 mL of wash
solution and heat to 60°C in a water bath. Be sure to verify the temperature of the
solution. Leave the remaining wash solution covered at room temperature. Warm
two clean empty plastic tubs in a 60°C oven.
5. Remove the hybridization bags, cut across to open, and drain the hybridization
solution. Remove the nylon membranes and place in separate plastic tubs. Then
add approx 125 mL of warm (60°C) blot wash buffer to each. Cover the tubs and
place in 60°C oven on a rocking platform for 10 min. Repeat this step once with
the remaining warm blot wash solution (see Note 13).
6. Prepare SA-HRP solution in a clean plastic tub using 250 mL of room tempera-ture blot wash solution and 10 µL of SA-HRP (final concentration of 40 ng/mL).
7. Following the second warm (60°C) wash, briefly rinse the membranes with
approx 100 mL of room temperature wash solution and transfer to the SA-HRP
solution. Place on a slowly gyrating platform for 15 min.
8. Drain the SA-HRP solution, briefly rinse with 100 mL of room temperature blot
wash solution, and decant. Then add another 400 mL of blot wash solution. Place
on a gyrating platform for 5–10 min. Repeat the wash with another 400 mL of
room temperature solution (see Note 14).
9. Place the wet membranes on clean dry filter paper. Quickly pipet 5 mL each of
ECL chemiluminescence reagents 1 and 2 into a clean glass tray and mix by
gentle agitation. Place each membrane in the tray and coat thoroughly with ECL
reagents for 1 min. Remove and affix the membranes to clean precut filter paper.
Mark the well locations with a fluorescent pen, label the blots as to probe type,
and seal over the membranes with plastic wrap. Develop chemiluminescent
image on X-ray film as usual (see Note 15).
Gene Fusion in Non-Hodgkin Lymphoma 155
3.5. Interpretation: t(14;18)/BCL-2/JH Results
Both agarose gel and chemiluminscent blot results are examined to deter-mine the presence or absence of a  BCL-2/JH fusion. A prominent ethidium-stained gel band with corresponding positivity on the blot denotes a positive
result (Figs. 2 and 3). Conversely, the lack of gel and hybridization signals is
Fig. 2. Agarose gel electrophoresis of BCL-2/JH PCR (MBR and mcr). Lanes A–C
and MBR/JH PCR products: A, patient sample; B, positive control; C, negative control
(no DNA). Note the difference in PCR amplicon fragment length between patient and
positive control, illustrating the range of size that can be generated by the BCL-2/JH
fusion. Lanes D–F are mcr/JH PCR products: D, patient sample; E, positive control;
F, negative control (no DNA). Lane L is a 100-bp molecular weight marker.
156 Viswanatha
consistent with the absence of the BCL-2/JH rearrangement. In any case, dem-onstration of a positive `-globin amplification is also required to exclude the
possibility of poor-quality DNA or PCR failure, prior to concluding that a PCR
test is negative. Occasionally one or more low-intensity bands may be observed
on agarose gels with diagnostic specimens; in my experience, these low-intensity (low-quantity) PCR products are nearly always artifactual in nature
and do not hybridize with the oligoprobes.
PCR product fragments will be of unique size in individual cases (Figs. 2
and 3), owing to the variability of breakpoint sites in the MBR or mcr loci, as
well as the possibility of fusion to any of six JH Ig heavy chain gene segments.
This feature can help to distinguish between true  BCL-2/JH rearrangements
and false-positive bands resulting from contamination, in assays with multiple
samples. For the MBR locus, PCR product sizes may range between 300 and
>1000 bp  (6) . The typical size range for most MBR BCL-2/JH amplicons is
approx 400–500 bp. PCR analysis of the mcr locus as presented herein can be
expected to identify amplified fragments in the 400-bp size range, although,
again, larger products up to 1000 bp can be encountered (6 ,23) .
Fig. 3. Chemiluminescent oligoprobe hybridization to detect MBR/JH and mcr/JH
fusions. BCL-2/JH PCR products hybridized with biotinylated internal MBR or mcr
oligoprobes. Lane (sample) designations are the same as for Fig. 2.
Gene Fusion in Non-Hodgkin Lymphoma 157
3.6. Conclusion
With the use of the single-round PCR technique described, one can expect to
identify slightly more than two thirds of all BCL-2/JH gene fusions arising from the
t(14;18)(q32;q21) in non-Hodgkin lymphoma (6). PCR therefore represents a pow-erful tool to detect this abnormality in a rapid and cost-effective manner. The
majority of fusions involving the MBR locus will be detected by this PCR method;
however, mcr locus fusions have a lower overall detection rate (approx 50%). The
latter situation arises in part because of a looser and less well-defined clustering of
break-fusion sites in the mcr such that breakpoints upstream or far downstream of
the mcr primer will not be detected. Notably, large amplicon PCR targets created
by both MBR and mcr locus  BCL-2/JH fusions may be inconsistently detected
owing to PCR inefficiency and may require nested PCR techniques. The presence
of large PCR amplicons is also of considerable significance when using archival
(fixed) tissues, because DNA extracted from paraffin-embedded sources is fre-quently compromised owing to crosslinking and degradation of high molecular
weight strands. As a result, the detection rate of BCL-2/JH for both MBR and mcr
locus fusions is usually lower in paraffin tissue specimens compared to fresh or
frozen tissue samples.
Other investigators have recently described the application of long distance PCR
(LD-PCR) to identify and further characterize the molecular anatomy
of BCL-2/JH fusions (28) . LD-PCR and sequencing analyses have revealed
extended breakpoint sites occurring in the far 3vMBR region and also 5vof the mcr
locus, largely accounting for the remainder of BCL-2/JH fusions not identified by
standard PCR techniques (28). To achieve a more comprehensive detection rate for
BCL-2 gene rearrangements, Southern blot hybridization or FISH methods have
been described (6,16,21,22,29). However, these approaches can be laborious and
require the availability of genomic or cloned probes. Nonetheless, Southern blot
hybridization or FISH remain important modalities for the detection of rare BCL-2
rearrangements, such as those involving the VCR.
Brief mention is made here regarding the distinction of detection rate versus
sensitivity of this PCR assay. Detection rate refers to the number of truly posi-tive BCL-2 gene–rearranged cases that can be identified by the assay (against
an established standard method). PCR sensitivity is defined in a dilutional
sense: that is, how capable is the assay of identifying low levels of the specific
gene fusion in question? Although  BCL-2/JH PCR is used in my laboratory
primarily to assist in the diagnosis and classification of lymphoid tumors, the
sensitivity of this assay can be optimized for detection of BCL-2/JH fusions at
a level of 1 in 104–105 cells. The high sensitivity of PCR and unique BCL-2/JH
amplicon size (and sequence) for a given patient sample creates a clonal “fin-gerprint” for assessment of MRD during or following therapy for lymphoma.
158 Viswanatha
This concept has been applied to patients with follicular lymphoma undergo-ing intensive chemotherapy with or without autologous bone marrow trans-plantation, in which the detection of residual BCL-2/JH fusion DNA can serve
as a prognostic marker of posttherapeutic outcome. In general, the molecular
clearance of residual BCL-2/JH DNA, or its absence in preinfusion peripheral
blood or bone marrow progenitor cell autografts, has been correlated with sig-nificantly prolonged disease-free and relapse-free survival  (30–34) . Finally,
the advent of highly sensitive and automated real-time fluorescent PCR tech-niques promises to revolutionize the measurement of MRD in these clinical
settings, by greatly increasing the accuracy and reproducibility of detection
(35 ,36) . However, as a cautionary note, BCL-2/JH fusion events have been re-ported at very low levels in normal individuals or in patients without
lymphoproliferative disease (37–42) , implying that errant BCL-2/JH rearrange-ments may occur in lymphoid cells in the absence of frank neoplasia. Although
standard PCR methods have not been shown to detect false-positive BCL-2/JH
fusions in histologically benign tissues (43) , the finding of rare event positivity
is of potential concern in more sensitive MRD analyses, unless sequencing and
design of patient tumor-specific primers or probes is employed (44) .
In conclusion, PCR evaluation of the  BCL-2/JH fusion abnormality can
be a valuable adjunct to the diagnosis and assessment of MRD of B-cell
lymphoproliferative disorders. As with any ancillary investigation, the results
of BCL-2/JH PCR analyses should be correlated with clinical, histologic, and
laboratory data to be of the greatest use in diagnostic medicine.
4. Notes
1. The use of proprietary DNA isolation kits has been adopted in my laboratory,
both for fresh or frozen tissue and paraffin-embedded tissue extractions. Many
“homemade” reagents are used for DNA isolation, some of which may be as
effective, or less costly. However, the consistent reagent quality of the former
products has been an added benefit to quality control issues in my clinical
molecular diagnostic setting.
2. All stock solutions utilized in PCR reactions are prepared with DEPC-treated
double-distilled water and stored or manipulated in similarly treated, autoclaved,
and dedicated glassware. Stock solution tubes of 10 mM dNTPs are made in quan-tity and labeled with a specific lot number. New reagent tubes are tested prior to
use in actual assays. The dNTPs, buffer, primers, and enzyme tubes are stored at
–20°C.
3. The Bio-Rad model 785 vacuum transfer system (Bio-Rad) is used in my labora-tory with reliable and excellent results. The setup parameters described in
Subheading 3. refer to this system and will vary slightly for different types of
apparatus.
Gene Fusion in Non-Hodgkin Lymphoma 159
4. Hybridization buffer can be made in quantity and aliquots stored at –20°C, then
removed, and warmed in a 60°C water bath when performing hybridization.
5. Because PCR techniques are generally very robust and capable of detecting small
amounts of initial nucleic acid target, control of potential contamination during
every step of the assay is paramount. My laboratory utilizes separate areas for
sample preparation (RNA and DNA isolation), PCR setup, and gel analysis/
hybridization. Analyzed PCR products (post-PCR) should never be returned to
or circulated through PCR setup and nucleic acid isolation areas. Nucleic acid
isolation and PCR stock and working solutions are kept separate from all other
laboratory reagents. Positive displacement micropipets (e.g., Rainin Microman;
Rainin, Emeryville, CA) and sterile, RNase-free, barrier-type, disposable pipet
tips are used throughout. Separate dedicated pipets are utilized in the nucleic acid
isolation and PCR areas.
6. For cell pellets (e.g., from blood, bone marrow, or body fluid samples), cells can
be obtained by the density gradient method (Ficoll Hypaque; Pharmacia) or red
cell lysis. The latter can result in the presence of residual heme, which may
inhibit PCR efficiency. Cell pellets are easily lysed and DNA can generally be
extracted without a proteinase K digestion. For tissue samples, the specimen is
quickly diced as finely as possible using a sterile disposable blade, and then
placed in lysis buffer with proteinase K. Larger tissue volumes or fibrotic speci-mens may require prolonged (i.e., overnight) digestion at 37 or 55°C.
7. Procurement of paraffin tissue sections must be done with considerable attention
to detail. The microtome area must be extremely clean, and preferably a new
disposable blade should be used between samples. Tissue curls can be carefully
placed in a 1.5-mL microfuge tube; however, tissue “flaking” is common and
may increase the risk of contamination. Generally, unknown samples are cut first,
then positive control(s), then negative control. As an alternative, unstained paraf-fin sections can be provided on glass slides and either scraped into the sample
tubes or carefully dissolved in xylene before transfer to the tube. The issue of
section thickness is controversial; although some anecdotal reports suggest that
thicker sections increase the concentration of PCR inhibitors, I have had good
results from sections between 5 and 20  µm thick. If adequate DNA has been
obtained and PCR inhibition is suspected, 1 10 and 1 50 dilutions of the DNA
extract can be made for PCR, to dilute out the effect of a potential inhibitor.
8. Xylene is a biohazardous organic chemical and must be used cautiously, prefer-ably in a ventilated hood area. Following dissolution of the paraffin and centrifu-gation, the xylene must be removed slowly, because the tissue pellet tends to be
poorly compacted and may be disrupted, leading to lower tissue (and thus DNA)
yield.
9. PCR steps are performed in thin-walled 100-µL MicroAmp tubes (Perkin-Elmer)
for use in a Perkin-Elmer model 9600 or 9700 thermal cycler, or equivalent
instrument. Mineral oil overlay is avoided with this approach. Other instruments
may require different PCR sample tubes and possibly the addition of an oil layer
before PCR.
160 Viswanatha
10. For the BCL-2 MBR positive control, DNA from an anonymous patient lymphoma
with a documented t(14;18) and MBR-JH fusion was used. Several cell lines contain-ing BCL-2/JH MBR rearrangements have been reported, including the DOHH-2 and
SU-DHL-4 lines. For the BCL-2 mcr positive control, the DHL-16 cell line was used
(kindly provided by Dr. J. Gribben, Dana Farber Cancer Institute).
11. For ease of analysis, I add 15  µL of 10 mg/mL EtBr directly to the warm,
prepoured gel.
12. If `-globin PCR products are on a separate gel, or are clearly separated from the
BCL-2/JH fusion gene PCR products (e.g., on the other half of a double-comb
gel), these are not transferred to the nylon membrane. If they are on the same part
of the gel, the `-globin PCR products are transferred along with the fusion gene
PCR products. The section of membrane containing  `-globin PCR products is
then separated and discarded prior to probe hybridization.
13. Following the specific probe hybridization and warm blot wash steps, several
blots can be combined and processed together in the SA-HRP and subsequent
room temperature wash steps.
14. During the SA-HRP and room temperature wash steps, the plastic tubs can be kept
loosely covered to prevent possible precipitation of the SSPE/SDS solution. Precipi-tation occasionally can occur in laboratories with lower ambient temperature.
15. Typical exposure times are between 5 and 15 min; however, strong signals are
often present on film within 1–3 min. The chemiluminescent output decays rap-idly after a few hours.
References
1. Harris, N. L., Jaffe, E. S., Stein, H., et al. (1994) A revised European-American
classification of lymphoid neoplasms: a proposal from the International Lym-phoma Study Group. Blood 84, 1361–1392.
2. Weiss, L. M., Warnke, R. A., Sklar, J., and Cleary, M. L. (1987) Molecular analy-sis of the t(14;18) chromosomal translocation in malignant lymphomas. N. Engl.
J. Med. 317, 1185.
3. Cleary, M. L. and Sklar, J. (1985) Nucleotide sequence of t(14;18) chromosomal
translocation breakpoint in follicular lymphoma and demonstration of a breakpoint
cluster region near a transcriptionally active locus on chromosome 18. Proc. Natl.
Acad. Sci. USA 82, 7439.
4. Bakhshi, A., Jensen, J. P., Goldman, P., Wright, J. J., McBride, O. W., Epstein,
A. L., and Korsmeyer, S. J. (1985) Cloning the chromosomal breakpoint of
t(14;18) human lymphomas: clustering around JH on chromosome 14 and near a
transcriptional unit on 18. Cell 41, 899.
5. Tsujimoto, Y., Cossman, J., Jaffe, E., and Croce, C. M. (1985) Involvement of the
bcl-2 gene in human follicular lymphomas. Science 228, 1440.
6. Horsman, D. E., Gascoyne, R. D., Coupland, R. W., Coldman, A. J., and Adomat,
S. A. (1995) Comparison of cytogenetic analysis, Southern analysis, and poly-merase chain reaction for the detection of t(14;18) in follicular lymphoma. Am.
J. Clin. Pathol. 103, 472–478.
Gene Fusion in Non-Hodgkin Lymphoma 161
7. Yabumoto, K., Akasaka, T., Muramatsu, M., Kadowaki, N., Hayashi, T., Ohno,
H., Fukuhara, S., and Okuma, M. (1996) Rearrangement of the 5vcluster region of
the bcl-2 gene in lymphoid neoplasm: a summary of nine cases.  Leukemia 10,
970–977.
8. Adachi, M., Cossman, J., Longo, D., Croce, C. M., and Tsujimoto, Y. (1989)
Varian translocation of the bcl-2 gene to immunoglobulin light chain gene in
chronic lymphocytic leukemia. Proc. Natl. Acad. Sci. USA 86, 2771.
9. Kouides, P. A., Phatak, P. D., Wang, N., and Bennett, J. M. (1994) B-cell acute
lymphoblastic leukemia with L1 morphology and coexistence of t(1;19) and
t(14;18) chromosome translocations. Cancer Genet. Cytogenet. 78, 23–27.
10. Yang, E. and Korsmeyer, S. J. (1996) Molecular thanatopsis: a discourse on the
BCL2 family and cell death. Blood 88, 386–401.
11. Hockenberry, D., Nunez, G., Milliman, C., Schreiber, R. D., and Korsmeyer, S. J.
(1990) Bcl-2 is an inner mitochondrial membrane protein that blocks programmed
cell death. Nature 348, 334.
12. Gascoyne, R. D., Adomat, S. A., Krajewski, S., Krajewska, M., Horsman, D. E.,
Tolcher, A. W., O’Reilly, S. E., Hoskins, P., Coldman, A. J., Reed, J. C., and
Connors, J. M.(1997) Prognostic significance of Bcl-2 protein expression and
Bcl-2 gene rearrangement in diffuse aggressive non-Hodgkin’s lymphoma. Blood
90(1), 44–51.
13. Hill, M. E., MacLennan, K. A., Cunningham, D. C., et al. (1996) Prognostic sig-nificance of BCL-2 expression and bcl-2 major breakpoint region rearrangement
in diffuse large cell non-Hodgkin’s lymphoma: a British National Lymphoma
Investigation Study. Blood 88(3), 1046–1051.
14. Hermine, O., Haioun, C., Lepage, E., et al. (1996) Prognostic significance of
bcl-2 protein expression in aggressive non-Hodgkin’s lymphoma. Groupe d’Etude
des Lymphomes de l’Adulte (GELA). Blood 87(1), 265–272.
15. Liu, J., Johnson, R. M., and Traweek, S. T. (1993) Rearrangement of the BCL-2
gene in follicular lymphoma: detection by PCR in both fresh and fixed tissue
samples. Diagn. Mol. Pathol. 2, 241–247.
16. Ladanyi, M. and Wang, S. (1992) Detection of rearrangements of the BCL2 major
breakpoint region in follicular lymphomas: correlation of polymerase chain reac-tion results with southern blot analysis. Diagn. Mol. Pathol. 1, 31–35.
17. Shibata, D., Hu, E., Weiss, L. M., Brynes, R. K., and Nathwani, B. N. (1990)
Detection of specific t(14;18) chromosomal translocations in fixed tissues. Hum.
Pathol. 21, 199–203.
18. Pezella, F., Gatter, K. C., and Mason, D. Y. (1989) Detection of 14;18 chromo-somal translocation in paraffin-embedded lymphoma tissue. Lancet 1, 779,780.
19. Crescenzi, M., Seto, M., Herzig, G. P., et al. (1988) Thermostable DNA poly-merase chain amplification of t(14;18) chromosome breakpoints and detection of
minimal residual disease. Proc. Natl. Acad. Sci. USA 85, 4869–4873.
20. Lee, M., Chang, K., Cabanillas, F., et al. (1987) Detection of minimal residual
cells carrying the t(14;18) by DNA sequence amplification.  Science 230,
1350–1354.
162 Viswanatha
21. Poetsch, M., Weber-Matthieson, K., Plendl, J. J., Grote, W., and Schlegelberger,
B. (1996) Detection of the t(14;18) chromosomal translocation by interphase
cytogenetics with yeast-artificial-chromosome probes in follicular lymphoma and
nonneoplastic lymphoproliferation. J. Clin. Oncol. 14, 963–969.
22. Taniwaki, M., Silverman, G. A., Nishida, K., Horike, S., Misawa, S., Shimazaki,
C., Miura, I., Nagai, M., Abe, M., Fukuhara, S., and Kashima, K. (1995) Translo-cations and amplification of the BCL2 gene are detected in interphase nuclei of
non-Hodgkin’s lymphoma by in situ hybridization with yeast artificial chromo-some clones. Blood 86, 1481–1486.
23. Ngan, B., Nourse J., and Cleary, M. L. (1989) Detection of chromosomal translo-cation t(14;18) within the minor cluster region of bcl-2 by polymerase chain reac-tion and direct genomic sequencing of the enzymatically amplified DNA in
follicular lymphomas. Blood 73, 1759–1762.
24. Cleary, M. L., Galili, N., and Sklar, J. (1986) Detection of a second t(14;18)
breakpoint cluster region in follicular lymphomas. J. Exp. Med. 164, 315.
25. Dyer, M. J. S., Zani, V. J, Lu, W. Z., et al. (1994) BCL2 Translocations in leuke-mias of mature B cells. Blood 83, 3682–3688.
26. Adachi, M., Tefferi, A., Greipp, P. R., Kipps, T. J., and Tsujimoto, Y. (1990)
Preferential linkage of BCL-2 to immunoglobulin light chain gene in chronic lym-phocytic leukemia. J. Exp. Med. 171, 559.
27. Segal, G. H., Jorgensen, T., Scott, M., and Braylan, R. C. (1994) Optimal primer
selection for clonality assessment by polymerase chain reaction analysis: II. Fol-licular lymphomas. Hum. Pathol. 25, 1276–1282.
28. Akasaka, T., Akasaka, H., Yonetani, N., Ohno, H., Yamabe, H., Fukuhara, S., and
Okuma, M. (1998) Refinement of the BCL2/immunoglobulin heavy chain fusion
gene in t(14;18)(q32;q21) by polymerase chain reaction amplification for long
targets. Genes Chromosomes Cancer 21, 17–29.
29. Pezzella, F., Ralfkiaer, E., Gatter, K. C., and Mason, D. Y. (1990) The 14;18
translocation in European cases of follicular lymphoma: comparison of Southern
blotting and the polymerase chain reaction. Br. J. Haematol. 76, 58.
30. Lopez-Guillermo, A., Cabanillas, F., McLaughlin, P., et al. (1998) The clinical
significance of molecular response in indolent follicular lymphomas. Blood 91,
2955–2960.
31. Corradini, P., Astolfi, M., Cherasco, C., Ladetto, M., Voena, C., Caracciolo, D.,
Pileri, A., and Tarella, C. (1997) Molecular monitoring of minimal residual disease
in follicular and mantle cell non-Hodgkin’s lymphomas treated with high-dose
chemotherapy and peripheral blood progenitor cell autografting. Blood 89, 724–731.
32. Freedman, A. S., Gribben, J. G., Neuberg, D., et al. (1996) High-dose therapy and
autologous bone marrow transplantation in patients with follicular lymphoma
during first remission. Blood 88, 2780–2786.
33. Gribben, J. G., Neuberg, D., Freedman, A. S., et al. (1993) Detection by poly-merase chain reaction of residual cells with the bcl-2 translocation is associated
with increased risk of relapse after autologous bone marrow transplantion for
B-cell lymphoma. Blood 81, 3449–3457.
Gene Fusion in Non-Hodgkin Lymphoma 163
34. Gribben, J. G., Saprito, L., Barber, M., Blake, K. W., Edwards, R. M., Griffin,
J. D., Freedman, A. S., and Nadler, L. M. (1992) Bone marrows of non-Hodgkin’s
lymphoma patients with a bcl-2 translocation can be purged of polymerase chain
reaction-detectable lymphoma cells using monoclonal antibodies and
immunomagnetic bead depletion. Blood 80, 1083–1089.
35. Luthra, R., McBride, J. A., Cabanillas, F., and Sarris, A. (1998) Novel 5v exonu-clease-based real-time PCR assay for the detection of t(14;18)(q32;q21) in
patients with follicular lymphoma. Am. J. Pathol. 153, 63–68.
36. Heid, C. A., Stevens, J., Kivak, K. J., and Williams, P. M. (1996) Real time quan-titative PCR. Genome Res. 6, 986–994.
37. Rauzy, O., Galoin, S., Chale, J. J., Adoue, D., Albarede, J. L., Delsol, G., and
al Saati, T. (1998) Detection of t(14;18) carrying cells in bone marrow and
peripheral blood from patients affected by non-lymphoid diseases. Mol. Pathol.
51, 333–338.
38. Dolken, G., Illerhaus, G., Hirt, C., and Mertelsmann, R. (1996) BCL-2/JH rear-rangements in circulating B cells of healthy blood donors and patients with non-malignant diseases. J. Clin. Oncol. 14, 1333–1344.
39. Limpens, J., Stad, R., Vos, C., de Vlaam, C., de Jong, D., van Ommen, G. B.,
Schuuring, E., and Kluin, P. M. (1995) Lymphoma-associated translocation
t(14;18) in blood B cells of normal individuals. Blood 85, 2528–2536.
40. Corbally, N., Grogan, L., Keane, M. M., Devaney, D. M., Dervan, P. A., and
Carney, D. N. (1994) Bcl-2 rearrangement in Hodgkin’s disease and reactive
lymph nodes. Am. J. Clin. Pathol. 101, 756–760.
41. Aster, J. C., Kobayashi, Y., Shiota, M., Mori, S., and Sklar, J. (1992) Detection of
the t(14;18) at similar frequencies in hyperplastic lymphoid tissues from Ameri-can and Japanese patients. Am. J. Pathol. 141, 291–299.
42. Limpens, J., de Jong, D., van Krieken, J. H., Price, C. G., Young, B. D., van
Ommen, G. J., and Kluin, P. M. (1991) Bcl-2/JH rearrangements in benign lym-phoid tissues with follicular hyperplasia. Oncogene 6, 2271–2276.
43. Segal, G. H., Scott, M., Jorgensen, T., and Braylan, R. C. (1994) Standard poly-merase chain reaction analysis does not detect t(14;18) in reactive lymphoid
hyperplasia. Arch. Pathol. Lab. Med. 118, 791–794.
44. Berinstein, N. L, Reis, M. D., Ngan, B. Y., Sawka, C. A., Jamal, H. H., and
Kuzniar, B. (1993) Detection of occult lymphoma in the peripheral blood and
bone marrow of patients with untreated early-stage and advanced-stage follicular
lymphoma. J. Clin. Oncol. 11, 1344–1352.
Detection of Breast Cancer Cells 165
14
Detection of Breast Cancer Cells
Using Immunomagnetic Beads and Reverse
Transcriptase Polymerase Chain Reaction
Scott Luke and Karen L. Kaul
1. Introduction
Molecular methods permit the detection of cells too few in number to be
detected by light microscopy, immunohistochemistry, or flow cytometry
(1–5) . Numerous investigators are therefore developing sensitive and specific
reverse transcriptase polymerase chain reaction (RT-PCR) assays for tumor
cell detection. The detection of small numbers of tumor cells in blood, lymph
node, and stem cell harvests may have a significant impact on our understand-ing of the spread of breast cancer, and eventually may impact the management
of breast cancer patients as well.
These assays are based on the detection of an amplified product from an
mRNA target uniquely expressed in a particular cell type. In the case of carci-noma, however, tumor-specific targets are not available. RT-PCR assays there-fore demonstrate the presence of cells of a particular type such as epithelial
cells, rather than tumor cells per se. It is important to keep in mind that normal
cells may also be detectable by RT-PCR assays, as can cells showing nonspe-cific expression of the mRNA of interest. Assay specificity must be rigorously
investigated.
Several markers for breast epithelial cells have been reported, including
cytokeratins, gross cystic disease fluid protein, and mammaglobin  (2 ,6–10) .
However, lack of specificity has been a commonly encountered problem (11) .
The use of immunomagnetic beads for positive selection of epithelial cells per-mits the elimination of false-positive signal from background expression of the
165
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
166 Luke and Kaul
chosen marker. Additionally, enhancements in sensitivity owing to enrichment
of tumor cells in the fraction used for RNA isolation have been observed
(12 ,13) . The beads used in this method are magnetic, polystyrene beads coated
with mouse monoclonal antibody (Ber-EP4) directed against human epithelial
antigen. The bound antiepithelial beads will target two membrane glycopro-teins found in both normal and neoplastic epithelial cells (14) .
This chapter outlines two RT-PCR assays that have been used successfully for
the detection of breast cancer cells in a variety of sample types. The first target
is cytokeratin-19 (CK-19), a widely expressed gene in cells of epithelial origin.
CK-19, a member of the family of cytokeratins, is a component of the mammalian
cell cytoskeleton (2,6,7). Detection of CK-19 expression is therefore potentially
useful for the study of many carcinomas. By contrast, mammaglobin, a member
of the uteroglobin gene family, appears to be specifically expressed in mammary
epithelium and present in the majority of breast carcinomas (10). The clinical util-ity of these assays is currently being investigated.
The general format of these assays for the detection of breast cancer cells
via RT-PCR includes the immunoselection and enrichment of epithelial cells,
followed by RNA preparation. cDNA is then generated via a reverse transcrip-tion protocol and amplified by PCR using primers specific for the chosen tar-get. The amplified products are first assessed on an agarose gel counterstained
with ethidium bromide (EtBr), followed by probe hybridization to increase
both the specificity and sensitivity of the assay. Additionally, when studying
human samples, an internal control such as `2-microglobulin (`2M) must be
RT-PCR amplified to demonstrate the intactness and ability to amplify iso-lated mRNA.
To ensure that amplification products truly reflect the presence of epithelial
cell RNA, primers must be carefully chosen to avoid complementarity to pro-cessed pseudogenes that may be present, and primers must be located across
exon-intron boundaries. Failure on either of these requirements may result in
production of amplicons from traces of genomic DNA that are indistinguish-able from the desired mRNA of the tumor cells.
2. Materials
2.1. Positive Control Cells
As a positive control for the CK-19 RT-PCR, the T47D cell line can be used
(see Fig. 1). T47D is an epithelial breast cancer cell line available from the
American Type Culture Collection (ATCC). Another epithelial breast cancer
cell line, BT-474 (also available from ATCC), is used as a positive control for
Detection of Breast Cancer Cells 167
the mammaglobin assay. Normal blood serves as a control for the presence of
the ubiquitous `2M mRNA target. Immunoselected normal blood is used as the
negative control in the CK-19 and mammaglobin assays.
2.2. Preparation of Sample
Care must be taken to avoid mRNA degradation from RNases. Gloves
should be worn at all times and changed often to avoid the introduction of
RNases. Diethylpyrocarbonate (DEPC)-treated water should be used in all
reagents. DEPC-treated water is made in-house (see Note 1). Samples should
be processed as quickly as possible (within 2 h of collection) to minimize loss
of mRNA from the presence of RNases owing to cellular destruction.
1. Patient blood, 5–10 mL collected in EDTA-treated Vacutainer tubes.
2. RNase-free microcentrifuge tubes and pipet tips.
3. Wash buffer consisting of 1X phosphate-buffered saline, pH 7.4, 0.1% bovine
serum albumin, and 1 mM EDTA.
4. Dynabeads® Epithelial Enrich immunobeads (Dynal). The beads are 4.5 µm in
diameter.
5. Dynal MPC® magnetic separator (Dynal).
Fig. 1. CK-19 RT-PCR of immunoselected blood using Dynabeads. A 1.5% agar-ose gel stained with EtBr showing the sensitivity of the CK-19 RT-PCR assay. Lanes
1, 5, and 9 are mRNA prepared from blood of healthy volunteers (all negative by
RT-PCR). Lanes 2–4, 6–8, and 10–12 correspond to the addition of 1000, 100, and 10
T47D cells to the volunteer blood, respectively. Lane 13 is a positive T47D control
(751 bp). Lane 14 is an mRNA blank and lane 15 is a 100-bp ladder.
168 Luke and Kaul
2.3. Isolation of RNA
Gloves should be worn and changed often to reduce the risk of contamina-tion with RNases present in the oils of the skin and to minimize the risk of
carryover between samples.
1. Trizol reagent (Life Technologies).
2. Chloroform (see Note 2).
3. RNase-free tubes and plasticware.
4. DEPC-treated deionized H2O.
5. Glycogen (Boehringer Mannheim).
6. Isopropanol.
7. 75% Ethanol diluted with DEPC-treated water.
2.4. Reverse Transcription
Master mix for the RT reaction should be made in large batches to minimize
run-to-run variability. Reagents may be susceptible to degradation via repeated
freezing and thawing, and all reagents should be aliquoted appropriately to
minimize this occurrence. Reverse transcription and amplification mixes uti-lize dUTP in place of dTTP so that amplicons can be degraded by uracil
N-glycosylase if desired to ablate contamination by amplicon carryover.
1. 25 mM MgCl2 (Perkin-Elmer), 10X PCR Buffer II (Perkin-Elmer), and DEPC-treated water.
2. Stock dNTP mixtures consisting of 10 mM for A, G, and C nucleotides (Gene
Amp RNA PCR kit; Perkin-Elmer); 20 mM stock of dUTP (Perkin-Elmer).
3. MuLV reverse transcriptase (25 U/µL), RNase Inhibitor (10 U/µL), and 25 µM
oligo d(T)16 (Perkin-Elmer).
2.5. Polymerase Chain Reaction
1. 25 mM MgCl2, 10X PCR Buffer II, DEPC-treated water.
2. AmpliTaq DNA polymerase (5 U/µL) (Perkin-Elmer).
3. 40 µM Stocks of each primer target.
CK-19 (15)  gives a 751-bp product:
Sense primer CK-19: GAC TAC AGC CAC TAC TAC ACG ACC
Antisense primer CK-19: AGC CGC GAC TTG ATG TCC ATG AGC C
(see Note 3).
Mammaglobin (10)  gives a 430-bp product:
Sense primer mammaglobin: CAG CGG CTT CCT TGA TCC TTG
Antisense primer mammaglobin: CAT AAG AAA GAG AAG GTG TGG
`2M (16) gives a 158-bp product:
Sense primer `2M: CTT GTC TTT CAG CAA GGA CTG G
Antisense primer `2M: CCT CCA TGA TGC TGC TTA CAT GTC
Detection of Breast Cancer Cells 169
2.6. Agarose Gel Electrophoresis
1. 1.5% Agarose in 1X TAE buffer. A 50X stock solution consists of the following
in a 1L of solution: 242.2 g of Trizma base (Sigma), 57.1 mL of glacial acetic
acid, 100 mL of 0.5 M EDTA (pH 8.0).
2. 10X Loading dye: 0.25% bromophenol blue, 0.25% xylene cyanol, 15% Ficoll
(Type 400); store at room temperature.
3. Staining box containing 1X TAE and 0.7 µg/mL of EtBr.
2.7. Solution Hybridization
1. 5% Acrylamide gel (acrylamide/bis-acrylamide 19:1, 40% stock solution
(Sigma).
2. Loading dye: 0.25% bromophenol blue, 0.25% xylene cyanol, 15% Ficoll (Type
400); store at room temperature.
3. Adenosine 5v-[a-32P] triphosphate (9.25 MBq), triethylammonium salt (250 µCi)
(Pharmacia).
4. T4 Polynucleotide kinase, cloned, pure (1000 U) (Pharmacia).
5. 10X One-Phor-All Buffer PLUS 10X solution (Pharmacia).
6. NAP-5 columns (Pharmacia).
7. 1X TE buffer consisting of 10 mM Tris-Cl (pH 7.4) and 1 mM EDTA (pH 8.0).
8. 1X TBE buffer: A 5X stock solution consisting of the following in a 1-L solution:
54 g of Trizma base, 27.5 g of boric acid, 20 mL of 0.5 M EDTA (pH 8.0).
9. DNA probe for CK-19 (100 µM; sense strand): GCG GGA CAA GAT TCT TGG
TG or mammaglobin (100 µM; sense strand): CTT TCT GCA AGA CCT TTG
GCT CAC. No probe is necessary for the `2M product.
10. Beta counter (Bioscan/QC 2000).
3. Methods
3.1. Preparation of Sample (see Note 4)
1. Collect 5–10 mL of patient blood in EDTA-treated Vacutainer collection tubes.
2. Chill the blood at 4°C for 20 min.
3. Transfer the blood to a 15-mL conical polypropylene tube (Becton Dickinson).
4. Add 25 × 106 washed Dynabeads Epithelial Enrich immunobeads (Dynal) to each
sample and rock gently at 4°C for 30 min to allow for immunomagnetic capture
of any present epithelial cells.
5. Following the incubation, place the conical tube containing the blood into a Dynal
Magnetic Particle Concentrator (Dynal MPC; Dynal) and rock gently for 5 min allowing
the immunomagnetic beads to migrate to the side of the tube abutting the magnets.
6. Using a vacuum aspirator and a clean, sterile pipet, slowly aspirate the blood
being careful to avoid the aspiration of any attached beads.
7. Remove the magnet and wash the beads by resuspending in 10 mL of cold buffer.
8. Replace the slide-out magnet and rock for 2 to 3 min allowing the beads to
migrate to the magnet.
9. Aspirate the supernatant.
10. Repeat steps 7–9 two more times.
170 Luke and Kaul
3.2. Isolation of RNA
RNA is isolated using the Trizol (Life Technologies) kit according to the
manufacturer’s protocol. Aerosol-resistant pipet tips are always used along
with pipettors dedicated to RNA preparation. RNA is isolated on a laboratory
bench kept separate from other activities.
1. Following the last wash and aspiration, add a volume of 0.8 mL of Trizol reagent
to lyse the captured cells adherent to the beads. Incubate the cells in the Trizol for
at least 3 min to allow complete detachment of nucleoprotein complexes. Trans-fer the volume to a clean 1.5-mL tube (see Note 5).
2. Add 160 µL of chloroform to each Trizol-containing tube and briefly vortex.
3. Spin the tubes in a refrigerated centrifuge at 4°C for 15 min at 12,000g.
4. An upper aqueous layer, an interphase, and a lower organic layer will be evident
following the centrifugation. Using a clean plastic transfer pipet (Scientific Prod-ucts), gently aspirate the upper aqueous layer containing the RNA into a clean
1.5-mL microcentrifuge tube.
5. Add 1 µL of glycogen (20 mg/mL) (Boehringer Mannheim) to the RNA-containing
solution and mix thoroughly.
6. Add 400 µL of 100% isopropanol and invert the tube several times. Let the solu-tion sit at room temperature for 10 min to allow the RNA to precipitate.
7. Centrifuge the sample for 10 min at 4°C and 12,000g.
8. A glycogen pellet (which will contain the desired RNA) will be visible at the
bottom of the tube. Discard the isopropanol supernatant and add 1 mL of 75%
ethanol in DEPC water to the pellet. Dislodge the pellet and invert the tube sev-eral times to wash.
9. Spin for 5 min at 4°C and 7500g.
10. Discard the supernatant and place the tube with its top open in a 55°C water bath
for 5 min to allow evaporation of excess ethanol. Do not overdry the pellet
because this will make it difficult to resolubilize the RNA.
11. Add 7.5 µL of DEPC-treated water and place in a 55°C water bath for 10 min
(see Note 6).
12. Spin the tubes down and place on ice in preparation for cDNA synthesis and
subsequent PCR.
3.3. Reverse Transcription
It must be stressed that when working with RNA, gloves must be worn and
changed often. Mixes to be used for cDNA synthesis and PCR reactions are
prepared in a UV-equipped hood separate from the sample preparation and the
amplification areas.
1. Add 1 µL of RNA sample (see Note 7) to 9 µL of the RT mix. Each RT reaction
vial contains the following with final concentrations listed: 5 mM MgCl2; 1X
Detection of Breast Cancer Cells 171
PCR Buffer II; 1 mM each of dATP, dCTP, and dGTP and 2 mM dUTP; 2.5 U/µL
of MuLV-RT; 1.0 U/µL of RNase Inhibitor; 2.5  µM oligo d(T)16; and DEPC-treated water to achieve a final volume of 10 µL.
2. The RT cycle parameters (GeneAmp 2400 thermalcycler; Perkin-Elmer) are as
follows: 10 min at 24°C, 15 min at 42°C, and 5 min at 99°C.
3. Store the cDNA at 4°C and proceed with the PCR reaction. If the PCR portion of
the assay is not to be carried out immediately (within the same day), store the
cDNA at –20°C until ready to proceed.
3.4. PCR Amplification
PCR amplification is performed in a total volume of 50 µL, which includes
the 10 µL from the RT reaction.
1. The PCR mix (40 µL/sample) contains the following with final concentrations
listed: 1X PCR Buffer II, 1.25 U of AmpliTaq DNA polymerase, 0.20  µM of
each primer, 2.0 mM MgCl2, and DEPC-treated water.
2. Add 40 µL of the PCR mix to each vial containing 10 µL of the cDNA solution
from the RT reaction.
3. The cycling parameters for CK-19 are as follows: cycle 1, 5 min at 94°C; cycles
2–36, 30 s at 94°C, 30 s at 68.5°C, and 30 s at 72°C; cycle 37, 7-min final exten-sion period at 72°C. The cycling parameters for  `2M are as follows: cycle 1,
105 s at 95°C; cycles 2–36, 15 s at 95°C and 30 s at 60°C; cycle 37, 7-min final
extension period at 72°C. The cycling parameters for mammaglobin are as fol-lows: cycle 1, 4 min at 94°C and 2 min at 80°C (see Note 8); cycles 2–36, 30 s at
94°C, 20 s at 57°C, and 30 s at 72°C; cycle 37, 7-min final extension period at
4°C. After amplification store the RT-PCR solution at 4°C.
3.5. Agarose Gel Electrophoresis
Tubes containing amplified products are opened and analyzed by electro-phoresis in a room separate from the sample reagent preparation area to avoid
amplicon contamination. Aerosol-resistant tips are used.
1. Mix 10 µL of each RT-PCR reaction thoroughly with 1 µL of loading dye and
load the wells.
2. Load one well with a 100-bp ladder (Pharmacia) mixed with dye. This will be
used as a reference for amplicon size.
3. Run the gel for 30 min to 1 h at 110 V, depending on the size of the desired
amplicon. CK-19 has a 751-bp product, mammaglobin displays a 430-bp prod-uct, and `2M has a 158-bp product.
4. Stain the gel in 1X TAE containing EtBr (0.7  µg/mL) for 20 min at room
temperature.
5. Visualize the presence or absence of bands in control and sample lanes with a UV
transilluminator.
172 Luke and Kaul
3.6. Labeling Probe for Solution Hybridization
In our assays, the antisense strand of the amplified product is hybridized
with a radioactively labeled probe (32P). The solution hybridization is employed
to increase both the sensitivity and specificity of the CK-19 and mammaglobin
assays. Because of the ubiquitous nature of `2M, a solution hybridization is
unwarranted for this primer set.
1. To label the probe, add the following to a 1.5-mL tube: 17 µL of H2O, 2.0 µL of
the DNA probe, 2.5 µL of 10X One-Phor-All buffer, 1.5 µL of a-32P ATP, and
2.0 µL of T4 polynucleotide kinase.
2. Incubate the tube in a 37°C water bath for 30 min.
3. Following the 30-min incubation, add 5 µL of 0.25 M EDTA (pH 7.0) to stop the
reaction.
4. Add 470 µL of 1X TE buffer to the labeled probe.
5. Prepare the NAP-5 column during the 30-min incubation (see Note 9).
6. Remove the top and bottom caps of the NAP-5 column and discard the liquid.
7. Wash the column with 10 mL of 1X TE buffer.
8. Following the 30-min incubation, add the 500-µL sample to the column, let the
liquid run through, and discard.
9. Add 500 µL of 1X TE buffer, allow the liquid to run through, and discard.
10. Add 750 µL of 1X TE buffer and collect this portion in a 1.5-µL tube.
11. On a beta counter obtain a radioactive count on the labeled probe.
3.6.1. Solution Hybridization
1. Put 10 µL of the amplified product into a 1.5-µL tube.
2. Add 10 µL of the radioactive probe to the amplified product.
3. Incubate the probe/amplicon mixture for 10 min at 95°C.
4. Following the 95°C incubation, allow the mixture to cool at room temperature. It
is during this step that the radioactive probe will hybridize to the antisense region
of interest.
5. Add 5 µL of loading buffer to each sample and mix well.
6. Load the 5% acrylamide gel and run for 1.5 h at 110 V in 1X TBE buffer (the
acrylamide gels are made with 1X TBE buffer).
7. Enclose the gel in plastic wrap and develop an autoradiogram. An overnight film
should be sufficient for analysis.
4. Notes
1. DEPC is used to purge reagents of RNase activity for use in RNA recovery pro-cedures. DEPC is added to stock solutions at a final concentration of 0.1% and
allowed to incubate overnight. DEPC must be completely destroyed by autoclav-ing the solution, where it degrades into carbon dioxide and ethanol. The degrada-
Detection of Breast Cancer Cells 173
tion of DEPC is absolutely necessary because even trace amounts will result in
modification of adenine residues. Note also that DEPC cannot be added to any
solution containing Tris or mercaptans. DEPC-treated water that has been auto-claved should be used to make solutions containing Tris or mercaptans and then
autoclaved again.
2. Aged chloroform can degenerate, producing a variety of undesirable byproducts,
including phosgene. Chloroform should be stabilized, ideally with alcohol, to
minimize this process. Chloroform should also be stored properly in a dark glass
bottle and used within a few months of opening.
3. We generally prefer amplicon sizes smaller than 300 bp, or even smaller than
200 bp if fixed tissues are to be analyzed. To avoid amplification of the CK-19
pseudogene, a published primer set amplifying a much larger amplicon was used.
The CK-19 primers span five introns and were carefully chosen to possess bases
that differ from the known processed pseudogene (4,6 ,15) . A high annealing tem-perature is used to increase the stringency of the assay, ensuring specific binding
of the primers to the desired sequence.
4. The protocol described here has been optimized for detection of breast cancer
cells in peripheral whole blood samples but can also be used for other sample
types. Fresh lymph nodes can be disaggregated to produce a single cell suspen-sion suitable for immunoselection. Dimethylsulfoxide-preserved stem cell har-vests can also be washed and immunoselected (unpublished observations). For
samples in which immunoselection cannot be performed (fixed or frozen lymph
nodes, some stem cell harvests) targets such as mammaglobin may be preferen-tial to cytokeratin for detection of breast cancer cells because of the absence of
low-level background expression.
5. There are several convenient points at which to stop during this procedure, per-mitting storage of the reactants.
6. RNA solutions are stored at –70°C. RNA is susceptible to degradation via freeze-thaw; therefore, we recommend that the RNA be aliquoted in single reaction
quantities. However, RNA can also be stored in 100% formamide with RT
formamide concentrations not to exceed 7% (http://www.nwfsc.noaa.gov/proto-cols/rnasoluble.html).
7. The RNA collected from patient blood is not quantified spectrophotometrically
because so few cells are gathered from the immunobead selection technique.
We nonetheless hydrate our RNA with 7.5 µL because that volume affords good
sensitivity.
8. The mammaglobin assay goes through a “hot start.” The DNA polymerase is not
added to the PCR mixture until the solution is at 80°C in cycle 1. Adding the
DNA polymerase at 80°C will minimize the amplification of nonspecific targets
that may occur if the DNA polymerase is added at room temperature.
9. Pharmacia NAP-5 columns are prepared disposable columns containing
Sephadex® G-25 medium of DNA grade for rapid and convenient desalting and
buffer exchange of oligonucleotides *10 mers.
174 Luke and Kaul
References
1. Brown, D. C., Purushotham, A. D., Birnie, G. D., and George, W. D. (1995)
Detection of intraoperative tumor cell dissemination in patients with breast can-cer by use of reverse transcription and polymerase chain reaction. Surgery 117,
96–101.
2. Burchill, S. A., Bradbury, M. F., Pittman, K., Southgate, J., Smith, B., and Selby,
P. (1995) Detection of epithelial cancer cells in peripheral blood by reverse tran-scriptase polymerase chain reaction. Br. J. Cancer 71, 278–281.
3. Choy, A. and McCulloch, P. (1996) Induction of tumour cell shedding into efflu-ent venous blood breast cancer surgery. Br. J. Cancer 73, 79–82.
4. Datta, Y. H., Adams, P. T., Drobyski, W. R., Ethier, S. P., Terry, V. H., and Roth,
M. S. (1994) Sensitive detection of occult breast cancer by the reverse-transcriptase polymerase chain reaction. J. Clin. Oncol. 12(3), 475–482.
5. Gross, H. J., Verwer, B., Houck, D., Hoffman, R. A., and Recktenwald, D. (1995)
Model study detecting breast cancer cells in peripheral blood mononuclear cells
at frequencies as low as 107. Proc. Natl. Acad. Sci. USA 92, 537–541.
6. Savtchenko, E. S., Schiff, T. A., Jiang, C. K., Freedburg, I. M., and Blumenburg,
M. (1988) Embryonic expression of the human 40-kD keratin: evidence from a
processed pseudogene sequence. Am. J. Hum. Genet. 43, 630–637.
7. Traweek, S. T., Liu, J., and Battifora, H. (1993) Keratin gene expression in non-epithelial tissues. Am. J. Pathol. 142, 1111–1118.
8. Mazoujian, G., Bodian, C., Haagensen, D. E., and Haagensen, C. D. (1989)
Expression of GCDFP-15 in breast carcinomas. Cancer 63, 2156–2161.
9. Mazoujian, G., Pinkus, G. S., Davis, S., and Haagensen, D. E. (1983) Immunohis-tochemistry of a gross cystic disease fluid protein (GCDFP-15) of the breast.
Am. J. Pathol. 110(2), 105–112.
10. Watson, M. A. and Fleming, T. P. (1996) Mammaglobin, a mammary-specific
member of the uteroglobin gene family, is overexpressed in human breast cancer.
Cancer Res. 56, 860–865.
11. Luke, S. and Kaul, K. (1998) Detection of breast cancer cells in blood using
immunomagnetic bead selection and reverse transcription-polymerase chain
reaction. Mol. Diagn. 3(3), 149–155.
12. Gomm, J. J., Browne, P. J., Coope, R. C., Liu, Q. Y., Bulawela, L., and Coombes,
R. C. (1995) Isolation of pure populations of epithelial and myoepithelial cells
from the normal human mammary gland using immunomagnetic separation with
Dynabeads. Analyt. Biochem. 226, 91–99.
13. Yaremko, M. L., Kelemen, P. R., Kutza, C., Barker, D., and Westbrook, C. A.
(1996) Immunomagnetic separation can enrich fixed solid tumors for epithelial
cells. Am. J. Pathol. 148, 95–104.
Detection of Breast Cancer Cells 175
14. Latza, U., et al. (1990) Ber-EP4; new monoclonal Ab which distinguishes epithe-lia from mesothelia. J. Clin. Pathol. 43, 213–219.
15. Eaton, M. C., Hardingham, J. E., Kotasek, D., and Dobrovic, D. (1997)
Immunobead RT-PCR: a sensitive method for detection of circulating tumor
cells. Biotechniques 21, 100–105.
16. Krafft, A. E., Duncan, B. W., Bijwaard, K. E., Taubenberger, J. K., and Lichy,
J. H. (1997) Optimization of the isolation and amplification of RNA from forma-lin-embedded tissue: the Armed Forces Institute of Pathology experience and lit-erature review. Mol. Diagn. 2(3), 217–230.
Detection of Prostate Cancer Cells 177
15
Molecular Detection of Circulating
Prostate Cancer Cells
Karen L. Kaul
1. Introduction
Molecular methods have proven extremely useful for the detection of occult
tumor cells and can yield valuable clinical information as well as a better
understanding of the mechanisms of metastasis and relapse of cancer (1) . In
the case of many hematopoietic malignancies, the presence of a unique
molecular marker such as a chromosomal translocation has made this task rela-tively straightforward. Carcinomas generally lack such markers, however. Cer-tain oncogene mutations have been targeted, but the lack of consistent and
specific markers has remained problematic.
An alternative approach has been increasingly utilized in the past decade
(2–4) . Reverse transcriptase polymerase chain reaction (RT-PCR) assays have
been designed to amplify the mRNA of a gene specifically expressed in the
cells of interest. Generally, such markers are not truly tumor specific, but are
expressed in both benign and malignant cells of the same origin. The mRNA of
prostate-specific antigen (PSA), e.g., is one such marker that has been success-fully used to detect prostate epithelial cells. The finding of such cells in the
circulation, lymph nodes, or bone marrow of a patient with prostate cancer
most likely reflects the spread of disease, although benign prostate epithelium
will also generate a positive RT-PCR result. Several clinical studies have been
performed to investigate the potential utility of RT-PCR assays in the manage-ment of patients with prostate cancer (4–26) . At this time, however, the role of
RT-PCR has not been clearly established. Further long-term follow-up studies
are needed and are under way. This chapter details one successful method for
the detection of prostate epithelial cells in blood and lymph node samples.
177
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
178 Kaul
A number of issues must be considered in the design of an RT-PCR assay
for the detection of occult tumor cells. The choice of a target is perhaps the
most critical. The target gene expression must be generally limited to the cells
of interest, with minimal or no expression in other cells within the sample,
such as leukocytes. As mentioned, target expression is generally seen in both
benign and malignant cells, and thus the potential for positivity owing to the
presence of benign cells must always be considered. Low-level expression of
the gene by other cells in the sample can also be problematic. Once a suitable
target is chosen, primer design becomes the next consideration. Primers ide-ally should bridge one or more intron/exon boundaries so that the amplicons
generated from mRNA in the RT-PCR process can be differentiated by size
from those originating from any genomic DNA that might be contaminating
the RNA sample. Additionally, primers must be chosen such that amplification
of genomic pseudogenes is avoided. During assay validation, it is worth inves-tigating these possibilities.
Assay validation takes place in two phases. Analytic validation utilizes
known negative and positive samples, often cultured cells, to demonstrate that
positive results truly correlate with the presence of tumor cells in the sample of
interest, and to further investigate the number of cells detectable. Clinical vali-dation encompasses analysis of a sufficient number of patient samples to dem-onstrate that the assay detects real tumor cells in actual samples without
false-positive signals in this heterogeneous sample group. This phase leads
into the actual RT-PCR study, in which long-term patient follow-up may be
needed to determine the clinical significance of a positive result.
2. Materials
As a positive control, the PSA-producing prostate cancer cell line LNCaP,
obtained from the American Type Culture Collection (Rockville, MD) is uti-lized. Cultured LNCaP cells are trypsinized and mixed in known numbers into
samples of prostate cell–free whole blood and used as a positive/sensitivity
control for the assay. Whole blood from normal volunteers, both male and
female, serves as negative control samples.
2.1. Collection and Preparation of Samples
Samples must be processed as quickly as possible to minimize loss of
mRNA; a maximum of 2 h is recommended.
1. Patient blood, 5–10 mL collected in EDTA-anticoagulant (Vacutainer; Becton-Dickenson) tubes (see Note 1).
2. RNase-free microcentrifuge tubes and pipet tips.
Detection of Prostate Cancer Cells 179
3. Polymorphprep (Robbins Scientific) or Ficoll-Hypaque gradient density media
(see Note 2).
4. Wash buffer consisting of 1X phosphate-buffered saline (PBS), pH 7.4, 0.1%
bovine serum albumin, and 1mM EDTA.
2.2. Isolation of RNA
Care must be taken to avoid mRNA degradation. Diethylpyrocarbonate
(DEPC)-treated water (see Note 3) should be used in all reagents. Gloves
should be worn and changed often to lessen the risk of contamination with
RNases and to minimize the risk of carryover between samples.
1. Trizol reagent (Life Technologies).
2. Chloroform (see Note 4).
3. RNase-free tubes and plasticware.
4. DEPC-treated deionized H2O.
5. Glycogen, Rnase free.
6. Isopropanol.
7. 75% Ethanol diluted with DEPC-treated water.
2.3. Reverse Transcription
Master mix for the RT reaction should be made in large batches to mini-mize run-to-run variability. Reagents are susceptible to degradation via
repeated freezing and thawing, and all reagents should be aliquoted appro-priately to minimize this exposure. The DEPC-treated water is made in-house (see Note 3).
1. 25 mM MgCl2, 10X PCR Buffer II (Perkin-Elmer), DEPC-treated water.
2. Stock dNTP mixtures of 10 mM for A, G, and C nucleotides (GeneAmp RNA
PCR kit; Perkin-Elmer); 20 mM stock of dUTP (Pharmacia).
3. 25 U/µL of Moloney murine leukemia virus (MuLV) RT, 10 U/µL of
RNase inhibitor, and 25 µM of oligo d(T)16 (all in the GeneAMP RNA PCR kit;
Perkin-Elmer).
2.4. Polymerase Chain Reaction
1. 10X PCR Buffer II (Perkin-Elmer).
2. 25 mM MgCl2.
3. DEPC-treated water.
4. AmpliTaq DNA polymerase (2.5 U) (Perkin-Elmer).
5. 40 µM Stocks of each primer target (PSA, prostate-specific membrane antigen
[PSMA] [see Note 5], or `2-microglobulin).
180 Kaul
The following primer pair, derived from Katz, et al.  (5) generates a 710-bp
amplicon (see Note 6):
Sense primer PSA710: CAC AGA CAC CCC ATC CTA TC
Antisense primer PSA710: GAT GAC TCC AGC CAC GAC CT
The following primer pair, derived from the PSA sequence (Genbank acces-sion no. #X14810) generates a 144-bp amplicon:
Sense primer PSA 144: AGG CTG GGG CAG CAT TGA ACC AGA GGA
Antisense primer PSA144: GTC CAG CGT CCA GCA CAC AGC ATG
AAC T
The following primers target the PSMA gene (Genbank accession no. M99487)
and yield a 165-bp amplicon:
Sense primer PSMA: AAA AGT CCT TCC CCA GAG TTC AGT
Antisense primer PSMA: ACT GTG ATA CAG TGG ATA GCC GCT
The primers for `2-microglobulin, used to amplify mRNA as a control for RNA
integrity, are taken from Krafft et al. (27)  and amplify a 158-bp fragment:
Sense primer `2M: CTT GTC TTT CAG CAA GGA CTG G
Antisense primer `2M: CCT CCA TGA TGC TGC TTA CAT GTC
2.5. Agarose Gel Electrophoresis
1. Tris-acetate EDTA (TAE) buffer: 40 mM Tris acetate, pH 7.4, 1 mM EDTA.
2. 1.5% Agarose gel in 1X TAE buffer.
3. 10X sample-loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol,
25% Ficoll.
4. Staining box containing 1X TAE and 0.7 µg/mL of ethidium bromide (EtBr).
2.6. Probe Labeling and Hybridization
1. Oligoprobes (stock solution of 100 µM):
a. PSA 710: Use a cocktail of Pr1: 5vCAA GTT CAC CCT CAG AAG GTG
ACC AAG TTC AT 3v; Pr2: 5vAGG CTG GGG CAG CAT TGA ACC AGA
GGA GT3v; Pr3: 5vTTC AGT GTG TGG ACC TCC ATG TTA TTT CCA
ATG ACT TGT GT 3v.
b. PSA 144 probe: TTC AGT GTG TGG ACC TCC ATG TTA TTT CCA ATG
ACG TGT GT.
c. PSMA 165 probe: GAG GTG TTC TTC CAA CGA CTT GGA ATT GCT.
2. a32P ATP (50 µCi), 3000 Ci/mmol (Amersham, Arlington Heights, IL).
3. T4 polynucleotide kinase (20 U) (Pharmacia).
4. One-Phor-All buffer (Pharmacia).
5. 0.25 M EDTA (pH 7.0).
6. NAP-5 Column (Pharmacia).
7. Beta counter (Bioscan QC 2000 or equivalent).
8. Boiling water bath.
Detection of Prostate Cancer Cells 181
2.7. Acrylamide Gel Electrophoresis
1. Vertical minigel apparatus and power supply.
2. Acrylamide/bis-acrylamide 19 1, 40% stock solution (Sigma, St. Louis, MO).
3. Ammonium persulfate, 10% solution, prepared fresh in distilled water.
4. TEMED (Sigma).
5. TBE buffer: 10 mM Tris-borate, pH 8.0, 2 mM EDTA.
6. 10X Sample-loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol,
25% Ficoll.
2.8. Autoradiography
1. X-ray film (Kodak, Rochester, NY).
2. Photographic developing tanks or equipment.
3. Methods
3.1. Collection and Preparation of Samples
Peripheral blood must be collected in a lavender top (EDTA) tube with a
minimum volume of 5 mL. Tubes must be delivered to the laboratory immedi-ately or within 2 h. Specimens that are clotted, drawn with incorrect anticoagu-lant, or frozen cannot be accepted for testing.
1. Prepare the leukocyte fraction, which includes circulating tumor cells, using Poly-morphprep according to the manufacturer’s suggested protocol. Ficoll-Hypaque
may also be used.
2. Wash the cells with cold PBS buffer and place on ice.
3.2. Isolation of RNA
RNA is isolated using the Trizol (Life Technologies) kit according to the
manufacturer’s protocol. Aerosol-resistant pipet tips are always used along
with pipettors dedicated to RNA preparation. RNA is isolated on a laboratory
bench kept separate from other activities.
1. Following the wash, add a volume of 0.8 mL of Trizol reagent to lyse the cells.
Pipet up and down several times to ensure cell lysis. Transfer to a 1.5-mL
microcentrifuge tube, and incubate for 3 min at RT. If necessary, samples can be
stored at –20°C at this point.
2. Add 160 µL of chloroform to each Trizol-containing tube and briefly vortex.
3. Spin the tubes in a refrigerated centrifuge at 4°C for 15 min at 12,000g.
4. An upper aqueous layer, an interphase, and a lower organic layer will be evident
following centrifugation. Using a clean plastic transfer pipet (Scientific Prod-ucts), gently aspirate the upper RNA-containing aqueous layer into a clean
1.5-mL microcentrifuge tube.
5. Add 1 µL of glycogen (20 mg/mL) (Boehringer Mannheim) to the RNA-containing
solution and mix thoroughly.
182 Kaul
6. Add 400 µL of 100% isopropanol and invert the tube several times. Let the solu-tion sit at room temperature for 10 min to allow the RNA to precipitate. If
desired, samples may be stored in alcohol at –80°C and will remain stable.
7. Centrifuge the sample for 10 min at 4°C and 12,000g.
8. A glycogen pellet will be visible at the bottom of the tube. Discard the isopro-panol supernatant and add at least 1 mL of 75% ethanol in DEPC-treated water to
the pellet; vortex to wash (see Note 7).
9. Spin for 5 min at 4°C and 7500g.
10. Discard the supernatant and place the tube with its top open in a 55°C water bath
for 5 min to allow evaporation of excess ethanol.
11. Add 7.5 µL of DEPC-treated water and place in a 55°C water bath for 10 min.
12. Spin the tubes down and place on ice in preparation for cDNA synthesis and
subsequent PCR.
3.3. Reverse Transcription
When working with RNA, gloves must be worn and changed often. Mixes
to be used for cDNA synthesis and PCR reactions are prepared in a hood sepa-rate from the sample preparation and amplification areas.
1. Aliquot 2.5 mM oligo d(T)16 and the desired DEPC-treated water volume to all
tubes. Add RNA to a final volume of 4 µL.
2. Anneal the primers at 65°C for 5 min. Place on ice.
3. Add 16 µL of RT master mix: 5 mM MgCl2, 1X PCR Buffer II, 1 mM each dNTP,
1 U of RNase inhibitor, 2.5 U of MuLV RT.
4. Incubate for 15 min at 42°C, 5 min at 99°C, and then 5 min at 5°C.
3.4. PCR Amplification
PCR amplification is performed in a total volume of 50 µL, including 10 µL
from the RT reaction.
1. Aliquot 40  µL of appropriate PCR Master Mix (containing  `2-microglobulin,
PSA, or PSMA primers) to the correct sets of tubes. PCR Master mix contains
2 mM MgCl2, 1X PCR Buffer II, 2.5 U of Taq polymerase, 0.2 µM of each primer,
and DEPC water. Keep the mixture on ice until completion of RT.
2. Add 10 µL of RT reaction.
3. Perform PCR amplification in a Perkin-Elmer 9600 or 2400 thermalcycler.
a. For PSA 710, cycle according to Katz et al. (5) : 1 cycle of 4 min at 95°C; 15
cycles of 1 min at 95°C, 1 min at 60°C and 30 s at 72°C; 11 cycles of 1 min at
95°C, 1 min at 60°C, and 1 min at 72°C; 7 cycles of 1 min at 95°C, 1 min
at 60°C, and 2 min at 72°C; 1 cycle of 15 min at 72°C; indefinite hold/storage
at 4°C.
b. For all other amplifications cycle as follows: 1 cycle of 4 min at 95°C; 35
cycles of 15 s at 95°C, 30 s at 60°C, and 30 s at 72°C; 1 cycle of 5 min at
72°C; indefinite hold/storage at 4°C.
Detection of Prostate Cancer Cells 183
3.5. Agarose Gel Electrophoresis
Tubes containing amplified products are opened and analyzed by electro-phoresis in a room separate from where the samples were prepared, and sepa-rate from where the mixes for the RT and PCR reactions were made.
Aerosol-resistant tips are used.
1. Mix 10 µL of each RT-PCR reaction thoroughly with 1 µL of loading dye, and
load the wells.
2. Load one well with a 100-bp ladder (Pharmacia Biotech) mixed with dye. This
will be used as a reference for amplicon size.
3. Run the gel for about 1 h at 110 V.
4. Stain the gel in 1X TAE containing EtBr (0.7 µg/mL) for 20 min at RT.
5. Visualize the absence or presence of bands in the control and sample lanes under
UV light. A 158-bp product should be visible in all specimens amplified using
`2-microglobulin primers.
6. Further analyze negative or inconclusive specimens by probe hybridization.
3.6. Probe Labeling and Hybridization
1. End label oligoprobe: Mix 1 µL of oligoprobe stock with 2.5 µL of buffer, 2 µL
of polynucleotide kinase, and 5  µL of radioactive ATP. Add sterile deionized
water to 25 µL. Incubate at 37°C for 30 min. Stop the reaction by adding 5 µL of
0.25 M EDTA (pH 7.0).
2. Separate labeled probe from unincorporated radioactive ATP using the NAP col-umn according to the manufacturers’ directions.
3. Mix 10 µL of amplicons and 10 µL of labeled probe.
4. Incubate at 95°C for 10 min, then at room temperature for 20 min.
5. Add 5 µL of loading dye.
3.7. Acrylamide Gel Electrophoresis
1. Prepare 5% acrylamide gel using acrylamide/bis-acrylamide 19 1, 40% stock
solution.
2. Electrophorese at 75–150 V until bromophenol blue dye reaches the end of
the gel.
3.8. Autoradiography
1. Wrap the gel in plastic and place next to a sheet of film in an autoradiography
cassette.
2. Expose for 4–16 h at –70°C, or as desired.
3. Develop the film using conventional methods.
184 Kaul
4. Notes
1. CPT tubes (Vacutainer; Becton Dickenson) contain a polymeric gel that sepa-rates mononuclear cells from neutrophils and red blood cells after a short cen-trifugation step. Our preliminary studies indicated that these could be used as
alternative collection tubes and may be particularly useful in the collection of
samples from other institutions or after hours, where complete gradient separa-tion is not possible.
2. Ficoll-Hypaque can be used in place of Polymorphprep with no apparent alter-ation in recovery of tumor cells from whole blood samples.
3. DEPC-treated water is prepared by adding DEPC to a final concentration of 0.1%
in distilled water. After an overnight incubation, autoclave to destroy the DEPC.
4. Aged chloroform can degenerate, producing a variety of undesirable byproducts,
including phosgene. Chloroform should stabilized, ideally with alcohol, to mini-mize this process. Chloroform should also be stored properly in a dark glass
bottle, and used within a few months of opening,
5. Although PSMA has been used for the detection of prostate epithelial cells, sev-eral laboratories have reported background expression of this marker in leuko-cytes and nonprostate cell lines, yielding nonspecific results and making PSMA
of questionable value in the detection of prostate cancer cells (28–30) .
6. Although our clinical studies have generally utilized the primers developed by
Katz et al. (5) , which amplify a 710-bp portion of the PSA gene, we generally
prefer amplicon sizes smaller than 300 bp, or even smaller than 200 bp if fixed
tissues are to be analyzed.
7. RNA is most stable when stored under ethanol at –80°C. Alternatively, RNA can
be reconstituted in formamide and can be used directly in reaction mixtures pro-vided that the final concentration of formamide is less than 8–10%.
8. The sensitivity of the assay should be carefully determined. In our hands, detec-tion of a single LNCaP cell in 5 mL of blood is possible; tenfold dilution or more
of this mRNA with that of unspiked blood can be done without losing the posi-tive signal, as shown in Fig. 1, because several hundred copies of PSA mRNA
are estimated to be present in each LNCaP cell (6) . Actual tumor cells may vary
in the copy number of the target message.
References
1. Glaves, D. (1986) Detection of circulating metastatic cells, in Cancer Metastasis:
Experimental and Clinical Strategies (Welch, D. R., Bhuyan, B. K., and Liotta,
L. A., eds.), Alan R. Liss, New York, pp. 151–165.
2. Moreno, J. G., Croce, C. M., Fisher, R., et al. (1992) Detection of hematogenous
micrometastasis in patients with prostate cancer. Cancer Res. 52, 6110–6112.
3. Ghossein, R. A. and Rosai, J. (1996) Polymerase chain reaction in the detection of
micrometastases and circulating tumor cells. Cancer 78, 10–16.
4. Raj, G. V., Moreno, J. G., and Gomella, L. G. (1998) Utilization of polymerase
chain reaction technology in the detection of solid tumors. Cancer 82, 1419–1442.
Detection of Prostate Cancer Cells 185
5. Katz, A. E., Olsson, C. A., Raffo, A. J., et al. (1994) Molecular staging of prostate
cancer with the use of an enhanced reverse transcriptase-PCR assay. Urology 43,
765–775.
6. Oefelein, M. G., Kaul, K., Herz, B., Blum, M. D., Holland, J. M., Keeler, T. C.,
Cook, W. A., and Ignatoff, J. M. (1996) Molecular Detection of prostate epithelial
cells from the surgical field and peripheral circulation during radical prostatec-tomy. J. Urol. 155, 238–242.
7. Oefelein, M. G., Ignatoff, J. M., Clemens, Q., Watkin, W., and Kaul, K. L. (1999)
Clinical and molecular followup after radical retropubic prostatectomy. J. Urol.
162, 307–310.
8. Ignatoff, J. M., Oefelein, M. G., Watkin, W., et al. (1997) Prostate specific anti-gen-reverse transcriptase-polymerase chain reaction assay in preoperative staging
of prostate cancer. J. Urol. 158, 1870–1875.
9. Israeli, R. S., Miller, W. H., Su, S. L., et al. (1994) Sensitive nested reverse tran-scriptase polymerase chain reaction detection of circulating prostate tumor cells:
comparison of prostate specific membrane antigen and prostate specific antigen
based assays. Cancer Res. 54, 6306–6310.
10. Seiden, M. V., Kantoff, P. W., Krithivas, K., et al. (1994) Detection of circulating
tumor cells in men with localized prostate cancer. J. Clin. Oncol. 12, 2634–2639.
Fig. 1. Agarose gel stained with EtBr showing PSA RT-PCR assay sensitivity for
the detection of LNCaP cells mixed with normal whole blood. Lane A, 100-bp
molecular weight marker; lanes B–F, dilutions of LNCaP mRNA increasing from 0.01
to 100 cell equivalents/2.5 mL of whole female blood; lane G, RNA blank; lane H,
LNCaP mRNA (see Note 8). (Reprinted from ref. 6  with permission from Lippincott
Williams & Wilkins.)
186 Kaul
11. Jaakkola, S., Vornanen, T., Leinonen, J., et al. (1995) Detection of prostatic cells
in peripheral blood: correlation with serum concentrations of prostate specific
antigen. Clin. Chem. 41, 182–186.
12. Loric, S., Dumas, F., Eschwege, P., et al. (1995) Enhanced detection of hematog-enous circulating prostatic cells in patients with prostate adenocarcinoma by
using nested reverse transcription polymerase chain reaction assay based on pros-tate-specific membrane antigen. Clin. Chem. 41, 1698–1704.
13. Sokoloff, M. H., Tso, C. L., Kaboo, R., et al. (1996) Quantitative polymerase
chain reaction does not improve preoperative cancer staging: a clinico-pathological
molecular analysis of 121 patients. J. Urol. 156, 1560–1566.
14. Ghossein, R. A., Rosai, J., Scher, H. I., et al. (1997) Prognostic significance of
detection of prostate-specific antigen transcripts in the peripheral blood of
patients with metastatic androgen-independent prostatic carcinoma. Urology 50,
100–105.
15. Kawakami, M., Okaneya, T., Furihata, K., et al. (1997) Detection of prostate can-cer cells circulating in peripheral blood by reverse transcription-PCR for hKLK2.
Cancer Res. 57, 4167–4170.
16. Corey, E., Arfman, E. W., Oswin, M. M., et al. (1997) Detection of circulating
prostate cells by reverse transcriptase-polymerase chain reaction of human glan-dular kallikrein (hK2) and prostate-specific antigen (PSA) messages. Urology 50,
184–188.
17. Corey, E., Arfman, E. W., Liu, A. Y., et al. (1997) Improved protocol for reverse
transcriptase polymerase chain reaction protocol with exogenous internal com-petitive control for prostate specific antigen mRNA in blood and bone marrow.
Clin. Chem. 43, 443–452.
18. Gomella, L. G., Raj, G. V., and Moreno, J. G. (1997) Reverse transcriptase poly-merase chain reaction for prostate specific antigen in the management of prostate
cancer. J. Urol. 158, 326–337.
19. Noguchi, M., Miyajima, J., Itoh, K., et al. (1997) Detection of circulating tumor
cells in patients with prostate cancer using prostate specific membrane-derived
primers in the polymerase chain reaction. Int. J. Urol. 4, 374–379.
20. Wood, D. P. and Banerjee, M. (1997) Presence of circulating prostate cells in the
bone marrow of patients undergoing radical prostatectomy is predictive of dis-ease-free survival. J. Clin. Oncol. 15, 3451.
21. Thiounn, N., Saporta, F., Flam, T. A., et al. (1997) Positive prostate-specific anti-gen circulating cells detected by reverse transcriptase-polymerase chain reaction
does not imply the presence of prostatic micrometastases. Urology 50, 245–250.
22. Ellis, W. J., Vessella, R. L., Corey, E., et al. (1998) The value of a reverse-transcriptase polymerase chain reaction assay in preoperative staging and follow-up of patients with prostate cancer. J. Urol. 159, 1134–1138.
23. Vessela, R. L., Lange, P. H., Blumenstein, B. A., et al. (1998) Multicenter
RT-PCR PSA clinical trial for preoperative staging of prostate cancer.  J. Urol.
159, 292A.
Detection of Prostate Cancer Cells 187
24. Van Nguyen, C., Song, W., Scardino, P. T., et al. (1998) RT-PCR for PSA and
hK2: implications for staging and patient management in men undergoing radical
prostatectomy. J. Urol. 159, 292A.
25. Nejat, R. J., Katz, A. E., Benson, M. C., et al. (1998) Enhanced RT-PCR for PSA
combined with serum PSA predicts pathologic stage and outcome in 300 radical
prostatectomy patients. J. Urol. 159, 291A.
26. Grasso, Y. Z., Gupta, M. K., Levin, H. S., et al. (1998) Combined nested RT-PCR
assay for prostate-specific antigen and prostate-specific membrane antigen and
prostate-specific membrane antigen in prostate cancer patients: correlation with
pathological stage. Cancer Res. 58, 1456–1459.
27. Krafft, A. E., Duncan, B. W., Bijwaard, K. E., Taubenberger, J. K., and Lichy,
J. H. (1997) Optimization of the isolation and amplification of RNA from forma-lin-fixed, paraffin-embedded tissue: the Armed Forces Institute of Pathology ex-perience and literature review. Mol. Diagn. 2, 217.
28. Kaul, K. L., Hanna, W. L., Luke, S., Oda, J., and King, S. T. (1997) Molecular
assays for detecting micrometastases in patient blood. Clin. Chem. 43, S92.
29. Lintula, S. and Stenman, U. (1997) Expression of prostate-specific membrane
antigen in peripheral blood leukocytes. J. Urol. 157, 1969.
30. Gala, J.-L., Heusterspreute, M., Loric, S., Hanon, F., Tombal, B., Van Cangh, P.,
De Nayer, P., and Phillipe, M. (1998) Expression of prostate-specific antigen and
prostate-specific membrane antigen transcripts in blood cells: implications for the
detection of prostate cells and standardization. Clin. Chem. 44, 472–481.
Detection of Clonal Genes in Lymphomas and Leukemias 189
16
Methods to Detect Clonal Gene Rearrangements
in Lymphomas and Leukemias
Naheed Mitha and Ronald C. McGlennen
1. Introduction
The process of lymphocyte differentiation involves structural alterations of
specific genes including those for the immunoglobulin (Ig) and T-cell receptor
(TCR) antigen genes. This process occurs very early in the differentiation of
B- and T-lymphocytes and involves an ordered program for splicing and rear-ranging segments of these genes, depending on cell lineage and level of differ-entiation. Specific DNA cutting and splicing enzymes result in the removal of
a number of constant, joining, and variable segments of the Ig and TCR genes.
Rearrangement of the VDJ and C segments occurs randomly during the pro-cess of B- and T-cell development; hence, the resultant gene rearrangement
varies from cell to cell. This results in a unique rearrangement of these genes
that encode for a specific Ig or TCR protein. A clonal population of lympho-cytes, however, will have a specific molecular structure of rearrangements.
Identification of this clonal population is central to the diagnosis of lympho-mas and lymphocytic leukemias, because virtually all forms of lymphoid
malignancies contain rearrangements of one or more antigen receptor genes.
Furthermore, as a clonal expansion, an individual neoplasm will contain the
identical rearranged gene throughout the population, serving as a unique clonal
marker (1) . However, it is important to be aware that lymphocyte clonality is
not equivalent to malignancy  (2) . Benign and reactive conditions may show
monoclonal rearrangements. Correlation with histology and immunophenotypic
studies is important in order to establish a definitive diagnosis of malignancy.
Similarly, the absence of clonal gene rearrangement may be seen in cases that
189
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
190 Mitha and McGlennen
appear malignant by histologic and immunophenotypic criteria. In these
instances, it is important to be aware of technical limitations of the assays and
sampling errors, which may result in a false-negative result.
Detection of either Ig or TCR gene rearrangements involves examination of
DNA from lymphoid cells that may be obtained from blood, bone marrow, or
tissue specimens. The polymerase chain reaction (PCR) and Southern blot
analysis are methods currently used for detecting Ig and TCR gene rearrange-ments. In the PCR, DNA is amplified with a series of consensus primer pairs
that bind to sequences of variable, diversity, and joining regions of these genes.
In our laboratory, five consensus primers are used for the TCR-`chain gene
rearrangements. These include one V`, two D`, and two J` primers. The prim-ers are combined into four sets as described in Subheading 3. Both V`J`primer
sets will detect only complete VDJ rearrangements and the D`J` primer sets
will primarily detect partial DJ and a few complete rearrangements  (3) .The
PCR assay for the IgH chain gene employs VH primers that anneal to frame-works I, II, or III (4) . In our laboratory, the JH primer binds to the framework
III region and JH2 to the framework II region. The detection rate with these
primers varies according to the type of lymphoma (4) . Framework I primers
are typically not employed because they require several distinct primers to iden-tify the rearrangement (4) . The products are then analyzed by polyacrylamide
gel electrophoresis (PAGE). If a significant population of cells contains a
unique rearrangement of these genes, it appears as a well-defined band within
a molecular size range specific for each set of primers on the electrophoretic
gel. Southern blot analysis utilizes restriction endonucleases that cut genomic
DNA into specific fragments in or around the Ig or TCR genes. These digested
fragments are then separated by gel electrophoresis. The separated DNA is
next transferred to a nylon membrane, denatured, and detected by hybridiza-tion to a radioactive probe. A clonal population of cells containing a unique
rearrangement of  Ig or TCR genes is indicated by the presence of a unique
nongermline band on the autoradiogram.
Both techniques have advantages and disadvantages. Southern blot analysis
of the IgH chain using only one probe and a limited number of restriction en-zymes is highly specific. The sensitivity of Southern blot in detecting a clonal
population ranges from 1 to 5% of specimen cells (5) . Although PCR is more
sensitive in detecting overall clonality (ability to detect 1 neoplastic cell
admixed with 100,000 normal cells), it has a significant risk of contamination
and a higher false-negative rate. In practice, the sensitivity of detecting IgH
chain gene rearrangement by PCR lies between 0.1 and 10%, depending on the
type of background cells, number of PCR cycles, and resolution of the gel
(6 ,7). Southern blot is more expensive and has a longer turnaround time. By
contrast, PCR is less expensive, has a shorter turnaround time, does not require
Detection of Clonal Genes in Lymphomas and Leukemias 191
the use of radioactivity, requires less DNA/RNA, and can be performed on
DNA extracted from fixed specimens. High-quality genomic DNA necessary
for Southern analysis cannot be obtained from paraffin-embedded tissues
owing to the degradation that occurs during the fixation process.
2. Materials
2.1. Polymerase Chain Reaction
2.1.1. Specimens
The following specimen types may be used for PCR-based analysis of  Ig
and TCR gene rearrangements.
1. Blood: 10–15 mL collected in acid citrate dextrose (ACD) or EDTA. Only the
mononuclear cells are required for this test. Therefore, the blood is first separated
by a Ficoll-Hypaque separation procedure and the mononuclear fraction is
extracted and analyzed.
2. Bone marrow: 5 mL (2 mL minimum if the white count is normal) collected in a
syringe containing ACD or EDTA.
3. Tissue: 5-mm3 tissue delivered to the laboratory within 1 h of collection or fro-zen in liquid nitrogen or on dry ice. Paraffin-embedded tissue can also be used
(see Note 1).
4. Other: DNA extracted from any nucleated cells.
2.1.2. Equipment and Supplies
1. Perkin-Elmer thermocycler 480 (Perkin-Elmer Cetus, Norwalk, CT).
2. AmpliTaq DNA Polymerase (Perkin-Elmer). 10X Buffer and 25 mM MgCl2 are
supplied with the AmpliTaq.
3. Bulk dNTPs, 100 mM each of dATP, dCTP, dGTP, dTTP (Perkin-Elmer).
4. Three JH primers have been described. JH-A is the most common primer and is
homologous with all the JH regions (8) . As mentioned in  Subheading 1., five
consensus primers combined into four sets have been described for T-cell `chain
gene rearrangements. They produce bands ranging between 55 and 100 bp. Prim-ers are as follows:
a. T`-V (19mer): 5v-TGT-A(CT)C-TCT-GTG-CCA-GCA-G-3v.
b. T`-D1 (23mer): 5v-CAA-AGC-TGT-AAC-ATT-GTG-GGG-AC-3v.
c. T`-D2 (23mer): 5v-TCA-TGG-TGT-AAC-ATT-GTG-GGG-AC-3v.
d. T`-J1 (17mer): 5v-ACA-GTG-AGC-C(GT)G-GT(CT)-CC-3v.
e. T`-J2 (20mer): 5v-AGC-AC(GCT)-GTG-AGC-C(GT)G-GTG-CC-3v.
f. JH1-S (25mer): 5v-CTG-TCG-ACA-CGG-CCG-TGT-ATT-ACT-G-3v.
g. JH-A (22mer): 5v-AAC-TGC-AGA-GGA-GAC-GGT-GAC-C-3v.
h. JH2-S (20mer): 5v-TGG-(AG)TC-CG(CA)-CAG-(GC)C(TC)-(TC)CN-GG-3v.
i. MBR (20mer): 5v-TTA-GAG-AGT-TGC-TTT-ACG-TG-3v.
j. Vg11 (20mer): 5v-TCT-GG(AG)-GTC-TAT-TAC-TGT-GC-3v.
k. Jg11 (19mer): 5v-CAA-GTG-TTG-TTC-CAC-TGC-C-3v.
l. bcl-1(s) (19mer): 5v-GAA-GGA-CTT-GTG-GGT-TGC-T-3v.
192 Mitha and McGlennen
m. CF20i-5v(s) (21mer): 5v-GGT-CAG-GAT-TGA-AAG-TGT-GCA-3v.
n. CF20i-3v(A) (21mer): 5v-CTA-TGA-GAA-AAC-TGC-ACT-GGA-3v.
5. Mini-Electrophoresis System (Schleicher & Schuell, Keene, NH) or Xcell Mini-Cell Electrophoresis System (Novex, San Diego, CA).
6. FB105 Power supply (Fisher Scientific, Pittsburgh, PA).
7. 6% TBE Precast polyacrylamide gels (cat. no. EC6265; Novex) (see Note 2).
8. Ethidium bromide (EtBr) (cat. no 161-0430; Bio-Rad, Hercules, CA).
9. Molecular weight ladder: D-15 DNA Marker (cat. no. LC5825; Novex).
10. Polaroid type 667 film.
11. MilliQ Water double-distilled H2O (ddH2O).
12. Stock dNTPs (1.25 mmol/L): To prepare a 1.25 mmol/L solution, 400 µL (1.6 mL
total) of each dNTP is mixed with 30.4 mL of sterile water (32.0 mL total vol-ume). The dNTP solution should be aliquoted (1 mL) and stored frozen at
–20°C until needed.
13. Stock primers (100 pmol/µL).
14. Working primers (3.0 pmol/µL): 30 microliters of each stock (100 pmol/µL) gene
rearrangement primer and 12.5 each of CF20 5v and CF20 3v are added and
diluted to 1000 µL with sterile water. These are then stored frozen at –20°C. The
CF20 5v and CF20 3v are primers that amplify a region approx 460–500 bp in
exon 20 of the cystic fibrosis gene; this amplification product serves as an inter-nal control. Primer mixtures are prepared to make the following nine sets. Each
mixture also contains the two CF20 primers: (T`-V) + (T`-JI); (T`-V) + (T`-J2);
(T`-D1) + (T`-J2); (T`-D2) + (T`-J2); Vg11 + Jg11; (JH-S) + (JH-A); MBR + JH-A;
JH2 + JHA; bcl-1 + JH-A.
15. Working PCR mix: A working mix is prepared for the total number of reactions.
It is advisable to prepare a slight excess of master mix to allow for pipeting error.
Typically allowing for a 10% error is adequate (see Table 1).
Table 1
Reaction Cocktail for Gene Rearrangements by PCR
Volume/sample Volume for 10 samples
(does not include pipeting error) (includes 10% pipeting error)
Component (µL)  (µL)
dNTPs 8.0 88
MgCl2 6.0 66
10X Buffer 5.0 55
Sterile H2O 17.15 188.65
Taq I polymerase 0.1 1.1
Total volume 36.25 398.75
Detection of Clonal Genes in Lymphomas and Leukemias 193
16. 18.5X TBE: To a 6-L Erlenmeyer flask, add 216 g of Tris base, 110 g of boric
acid, and 80 mL of 0.5 M EDTA, pH 8.0. Add about 2 to 3 L of MilliQ water and
mix until dissolved. Add MilliQ water to 4 L. Aliquot 500 mL/bottle and store at
room temperature.
17. 1X TBE: Dilute 500 mL of 5X TBE to 2.5 L with MilliQ water. Store at room
temperature.
18. PCR control: A “no template” control reaction in which water is substituted for a
DNA specimen is included with each PCR setup; this is a check for the presence
of amplified DNA contamination. It is recommended to rotate primer pairs
according to the test required.
19. Gene rearrangement analysis controls: Controls may be obtained from investiga-tors or from suppliers of cell lines, e.g., American Type Culture Collection:
a. Ig Genes: JH1, FJO (EBV-transformed lymphoblastoid cell line); mbr,
RLBCL2 cell line; JH2, 95-80 (positive patient sample); bcl-1, 1094 cell line.
b. TCRB: (VJ1), adult T-cell leukemia lymphoma.
c. PCR amplification control: Exon 20 of the CFTR gene is used as an internal
control for PCR amplification.
2.2. Southern Blot Analysis
2.2.1. Specimen
Refer to Subheading 2.1.1.
2.2.2. Equipment and Supplies
1. UV crosslinker (Stratalinker; Stratagene, La Jolla, CA).
2. Hybridization oven (Robbins Scientific, Sunnyvale, CA).
3. Horizontal Electrophoresis System (Bio-Rad).
4. Seal-a-meal bags.
5. Molecular weight ladder: Lambda phage DNA digested with BstEII.
6. X-Ray film (cat. no. 04-441-95; Fisher, Itasca, IL).
7. Whatman 3MM chromatography paper.
8. Nick Translation System (cat. no. 18160-010); Gibco-BRL Life Technologies,
Gaithersburg, MD).
9. Oligolabeling Kit (cat. no. 27-9250-01; Pharmacia Biotech). The kit contains the
following components:
a. Reagent mix: Buffered aqueous solution containing dATP, dGTP, dTTP, and
random hexadeoxyribonucleotides.
b. FPLCpure™ Klenow fragment: Buffered glycerol solution (5–10 U/µL).
c. Control DNA: Aqueous solution or hDNA-HindIII restriction fragments.
d. Carrier DNA: Aqueous solution of calf thymus or salmon sperm DNA at
1 mg/mL.
10. Restriction endonucleases: Purchased from Gibco-BRL Life Technologies. (See
Table 2.)
194 Mitha and McGlennen
11. Recombinant plasmid probes: The plasmids given in Table 3 were obtained in
our laboratory from investigators. However, Dako (Carpinteria, CA) offers sev-eral probes including IGHJ6 (Ig heavy chain) IGKJ5; IGKC and IHKDE (kappa
light chain); TCRBC and TCRBJ2 (TCR` locus); and TCRDJI, TCRDC4,
TCRDRE (TCRblocus). It is beneficial to run several enzyme probe combina-tions to maximize detection of clonal rearrangements (Fig. 1).
12. Blotting Solution II (0.5  M NaOH, 1.5  M NaCl): Dilute 120 g of NaOH and
528 g of NaCl to 6 L with MilliQ water (or 360 g of NaOH and 1584 g of NaCl to
18 L).
13. Blotting Solution III (3 M NaCl, 1 M Tris, pH 7.5): Dilute 1046.4 g of NaCl and
726.6 g of Tris to about 5 L with MilliQ water. Adjust the pH to 7.5 with concen-trated HCl. Start with about 200 mL of acid and then measure the pH as more
acid is added. This requires about 350 mL of acid.
14. EtBr (1 mg/mL): Dilute 1 tablet (11 mg) of EtBr with 11 mL of sterile MilliQ
water. Make in a sterile screw-capped tube. Let it dissolve for about 4 min. Wrap
in foil and store at room temperature.
15. Prehybridization solution (total volume of 150 mL): MilliQ water (77 mL); 20X
SSPE (45 mL); 50X Denhardt’s solution (15 mL); 25% sodium dodecyl sulfate
(SDS) (3 mL); salmon sperm DNA, denatured by boiling for 5 min (10 mL).
16. Hybridization solution (total volume of 10.0 mL): 20X SSPE (3.0 mL),
formamide (5.0 mL), 50% dextran sulfate (1.66 mL), 25% SDS (0.2 mL).
17. Salmon sperm DNA: In a 1-L Erlenmeyer flask add 5 g of DNA sodium salt type
III from salmon testes and dissolve in 500 mL of ddH2O. Shake until evenly
dispersed. Dispense into a wide-mouthed bottle and autoclave to sterilize. Store
refrigerated at 4°C.
18. 6X Loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol, 15% Ficoll.
Dilute 0.25 g of bromophenol blue, 0.25 g of xylene cyanol, and 15.0 g of Ficoll
to 100 mL with sterile water. Store in a 100-mL bottle at room temperature.
19. Equilibration buffer, TE, pH 7.5 (for Pharmacia nick translation columns):
Dilute 5.0 mL of 1 M Tris-HCl, pH 7.5, and 1.0 mL of 0.5 M EDTA, pH 8.0, to
500 mL with MilliQ water and autoclave.
20. 10% SDS: In a 2-L Erlenmeyer flask add 100 g of SDS. Slowly add 1.0 L of
ddH2O. Heat to 68°C to assist in dissolution.
Table 2
Restriction Enzymes Used in Southern Blot Analysis
Restriction enzyme Stock concentration (U/µL) Cat. no.
BamHI 50 15201-049
EcoRI 50 15202-039
BglII 50 15213-036
HindIII 50 15207-038
Detection of Clonal Genes in Lymphomas and Leukemias 195
21. 25% SDS: In a 2-L Erlenmeyer flask add 250 g of SDS. Slowly add 1.0 L of
ddH2O. Heat to 68°C to assist in dissolution.
22. 25X saline sodium citrate (SSC): In a 4-L Erlenmeyer flask add 238.6 g of NaCl
(3.75 M) and 220.8 g of sodium citrate (0.375 M). Dissolve the solid with ddH2O
to 2 L. Autoclave for sterility.
23. 18X SSC + 1 M ammonium acetate: In a 4-L Erlenmeyer flask add 473.4 g of
NaCl (2.7 M) and 238.2 g of sodium citrate (0.27 M). Add 600 mL of 5 M NH4Ac
and ddH2O to a final volume of 3 L.
24. 0.1X SSC + 0.1% SDS: In a 6-L Erlenmeyer flask add 24 mL of 25X SSC, 60 mL
of 10% SDS, and 5.916 L of ddH2O.
25. 0.2X SSC + 0.1% SDS: In a 20-L carboy add 160 mL of 25X SSC, 200 mL of
10% SDS, and 19.64 L of ddH2O.
26. 2X SSC + 0.1% SDS: In a 10-L vessel add 800 mL of 25X SSC, 100 mL of 10%
SDS, and 9.1 L of ddH2O.
27. 20X SSPE: In a 4-L Erlenmeyer flask add 701.2 g of NaCl (3.0 M), 110.4 g of
NaHPO4 ·H2O, and 29.6 g of disodium EDTA. Add approx 3.2 L of ddH2O until
the solute dissolves. Adjust the pH to 7.4 with 10 N NaOH. Add ddH2O to 4 L.
Sterilize by autoclaving.
28. 10X TAE: In a 6-L Erlenmeyer flask add 290.4 mL Tris base and 120 mL of
0.5 M EDTA (pH 8.0). Add approx 5.0 L ddH2O and 68.52 mL of glacial acetic
acid. Adjust to a final volume of 6 L with ddH2O.
29. 1X TE (pH 7.0): In a 4-L Erlenmeyer flask add 40 mL of 1 mol/L Tris (pH 7.5)
and 8 mL of 0.5 mol/L EDTA (pH 8.0). Adjust to pH 7.0 and then add ddH2O to
a final volume of 4 L. Autoclave to sterilize.
30. 1X TE (pH 8.0): In a 4-L Erlenmeyer flask add 40 mL of 1 mol/L Tris (pH 8.0)
and 8 mL of 0.5 mol/L EDTA (pH 8.0). Adjust to pH 8.0 and then add ddH2O to
a final volume of 4 L. Autoclave to sterilize.
31. MilliQ water (ddH2O).
32. Negative (germline) control: Patient samples that have previously been demon-strated to be negative for gene rearrangements are used in our laboratory.
Table 3
Common Probes Used in Gene Rearrangement Analysis: Plasmid Sources
Insert Bacterial Antibiotic Cloning Bacterial
Plasmid  size vector selection site strain
JH 5.6 pUC-19 Amp BamHI-HindIII DH5a
C-g 2.5 pBR322 Amp EcoRI HB101
C-h 0.7 pUC-9 Amp EcoRI-HindIII HB101
TCR-a J1-2 0.7 pUC-18 Amp EcoRI-HindIII HB101
TCR-b V2 0.576 pUC-18 Amp EcoRI-HindIII HB101
TCR-` 0.96 pUC-18 Amp PstI HB101
EBV 2.3 pBR322 Amp EcoRI-BamHI HB101
196 Mitha and McGlennen
3. Methods
3.1. DNA Isolation
The Puregene™ DNA isolation Kit (Gentra Systems, Minneapolis, MN)
allows rapid isolation of high-quality genomic DNA from whole blood, cul-tured cells, and tissue (see Chapter 34). It is based on published salting-out
procedures, thus eliminating the use of toxic organic solvents. Other methods
that yield intact DNA can be used (see Chapters 1 and 2).
3.2. Polymerase Chain Reaction
This protocol is designed for use with 0.5-mL PCR tubes. Adjustments may
be required if 0.2-mL tubes are used.
1. Dilute the genomic DNA to 0.067 mg/mL.
2. Add 36.25 µL of working PCR mix to each tube and add 7.5 µL of patient sample
or control to the respective tubes.
3. Vortex the tubes, spin briefly in a microcentrifuge, and add 1 drop of mineral oil
to each tube before placing the tubes in a thermocycler.
4. Perform “hot start” PCR by setting the temperature of the thermocycler to 80°C.
Place the tubes in the thermocycler in order according to the primer pairs to be
Fig. 1. PCR gel and Southern blot analysis demonstrating TCR`chain gene rear-rangement. In this case Southern blot is used to verify the rearrangement detected by
PCR analysis. Abbreviations: MW = molecular weight marker; Neg = negative con-trol sample; Pat = patient sample; E = EcoRi enzyme digests; H = HindIII enzyme
digests.
Detection of Clonal Genes in Lymphomas and Leukemias 197
added. Note: Nonspecific primer annealing can be minimized by putting in tubes
for two primer sets at a time, adding primers, and then putting in more tubes.
5. Add 6.25 µL of primer pairs to their respective tubes.
6. Start the program for gene rearrangement PCR using the following thermocycler
parameters: 4 min at 94°C (initial denaturation step); 40 cycles of 1 min at 94°C
(denature), 55 s at 60°C (anneal), 1 min at 73°C (extension). The 40 cycles are
followed by a final 5-min extension at 72°C. Samples may be stored indefinitely
at 4°C.
3.2.1. Gel Electrophoresis of PCR Products
1. Load 0.5  µL of the molecular weight marker in lane 1 along with 3 µL of 6X
loading dye and 10 µL of H2O. Then use 3 µL of 6X loading dye and 15 µL of
PCR product (patient samples and controls) for the other lanes.
2. The recommended gel setup is as follows: lane 1: molecular weight ladder; lanes
2–10: VJ1, VJ2, D1J2, D2J2, Vg11Jg11, JH1, mbr, JH2, no template control. Run
for 55 min at 100 V. The bottom dye will be about halfway between the bottom
two lines on the PAGE cassette.
3. Stain with EtBr and photograph the gel using Polaroid type 667 film using f-stop
11 at 1 s.
3.2.2. Interpretation of PCR Results
Results are reported as positive or negative (Fig. 2). A smear pattern indi-cates polyclonality. A positive result is based on the presence of a sharp band
with well-defined edges and a width not more than 1 mm, in the ranges for the
Fig. 2. PCR gels demonstrating TCR and Ig heavy-chain positive and negative
controls. Note the nonspecific banding patterns seen with VJ1, VJ2, JH2 and BCL-1.
MW = molecular weight marker.
198 Mitha and McGlennen
Table 4
Size Range for Interpretation of a Monoclonal Band
and the Expected Nonspecific Bands
Primer Expected size range (bp)   Nonspecific bands
JH1 100–150 50, 65, 100
D1J2   55–100 65, 85, 120, 175, 185
Vg11Jg11   70–110 50, 75
MBR     222         —
JH2 220–250         —
VJ1   55–100         —
VJ2   55–100         —
D2J2   55–100         —
Fig. 3. PCR gels demonstrating clonal TCR`chain gene rearrangements within a
polyclonal smear. The smear most likely represents background-reactive, polyclonal
T-lymphocytes in this case of a T-cell lymphoma.
Detection of Clonal Genes in Lymphomas and Leukemias 199
primer pairs given in Table 4 (see Notes 3–5). A clonal band may be masked
in a polyclonal smear, as is sometimes seen in T-cell lymphomas (see Fig. 3).
A monoclonal population may also have two sharp bands, as is seen in some
cases of chronic lymphocytic leukemia. These are usually a consequence of
rearrangement of both alleles of that particular gene. Cases with more than two
bands or a broad-based smear of bands are interpreted as oligoclonal and
polyclonal, respectively. True clonal bands may also vary with a certain size
range depending on the specific primers used. Empty lanes without smears,
bands, primer/dimer artifact, or amplification of CF exon 20 should be regarded
as failed PCR rather than negative, and repeated if possible.
Distinctive nonspecific primer–dependent banding patterns are seen and posi-tive controls should be routinely performed. The primers used for the TCR-`chain
gene rearrangement produce nonspecific bands at 65, 85, 120, 175, and 185 bp. In
addition, an internal control should be used in each reaction tube to verify amplifi-cation of the individual samples. Tissues with a scant lymphoid inflammatory infil-trate may yield pseudoclonal bands owing to amplifications of normal
rearrangements present in just a few lymphocytes. These polyclonal bands are usually
smaller and less intense than the clonally rearranged ones and often occur in pairs.
False-negative results in true clonal populations, by PCR, are primarily
owing to an inability of the selected primers to anneal effectively to selected
exons of the recombined IgH chain gene. Other reasons include low DNA qual-ity, amplification of nonclonal DNA, suboptimal conditions, chromosomal
translocations, presence of deletions/mutations in the IgH chain genes, and
presence of rearranged DNA below the sensitivity level of the assay (9 ,10) .
One specific protocol modification is to decrease the annealing temperature,
which may increase the detection rate. Increasing the number of PCR cycles
may also increase detection.
PCR has the highest false-negative rate in cases of follicular lymphomas. The
PCR-based assay for the detection of t(14;18) detects a rearrangement in 50–60%
of follicular lymphomas, 60% of cases with a cytogenetically proven t(14;18), and
approx 70% of cases detectable by Southern blot (11–15). Both Southern blot and
PCR can fail to detect a rearrangement if the breakpoint occurs outside the restric-tion enzyme or primer recognition site (16). Our laboratory has identified at least
three bcl2 rearrangements of differing molecular weights by PCR (Fig. 4).
Crossreactivity with EBV DNA may lead to false-positive results, and confirma-tion of these cases by Southern blot or slot blot hybridization is required (17).
“Cross-lineage” rearrangements are not uncommon  (18–20)  (Fig. 5). IgH
chain gene rearrangements have been detected in some T-cell neoplasms.
Approximately 25% of precursor B-cell acute lymphoblastic leukemias (ALLs)
contain rearrangements of the IgH chain genes, and as many as 40% may have
rearranged T-gamma genes (21) . More than 50% of tdt positive acute myeloid
200 Mitha and McGlennen
leukemias show rearrangements of both IgH and TCR-`and TCR-achain genes
(8) . Because of this phenomena of “lineage infidelity,” both Ig and TCR assays
should be routinely performed, and the results should be evaluated with mor-phology and immunophenotypic data. However, rearrangements of both IgH
and light chain genes is a valid marker of B-cell lineage, because coexpression
of these has not clearly been identified in T-lineage cells (22 ,23) .
3.3. Southern Blot Analysis
3.3.1. DNA Isolation
Refer to Subheading 3.1.
3.3.2. Evaluation of Genomic DNA
Both quantitative and qualitative evaluation is done to ascertain evidence of
degradation. Quantitation of DNA is performed by measuring the absorbance at
260 nm, and evidence of degradation is sought by electrophoresis of a small aliquot
in an agarose gel with EtBr staining. These protocols are described in Chapter 2.
Fig. 4. Three PCR gels demonstrating four  bcl2 rearrangements of differing
molecular weight. The slot blot was performed in three of these patients and confirms
the presence of an Ig heavy-chain–bcl2 gene fusion. Abbreviations: MW = molecular
weight marker; Pat = patient sample.
Detection of Clonal Genes in Lymphomas and Leukemias 201
Fig. 5. PCR gels demonstrating a “cross-lineage” rearrangement. The patient had a
history of follicular small cleaved lymphoma diagnosed on a lymph node biopsy.
Molecular testing of the staging bone marrow biopsy (shown above) demonstrated a
bcl2 gene rearrangement as well as TCR-` and TCR-a chain gene rearrangements.
MW = molecular weight marker.
Table 5
Recommended Probe/Enzyme Combinations
Probe Enzymes Expected germline band sizes, kb (3 ,4)
CT-` EcoRI 11 and 4.2
HindIII 7.2, 6.2, and 3.7
JT-a EcoRI 3.4 and 1.5
HindIII 5.4 and 2.2
C-g BamHI 12
HindIII 5.4
JH BglII 4
HindIII 11 and 3.5a
EBV Bam None present
aThe 3.5-kb band represents a crosshybridizing band (4) .
202 Mitha and McGlennen
3.3.3. Restriction Digestion
Make BamHI, EcoRI, BglII, and HindIII restriction endonuclease digests using
20 µg of genomic DNA under the appropriate digestion conditions recommended
by the manufacturer. Table 5 lists the recommended probe/enzyme combinations.
3.3.4. Submerged Gel Electrophoresis
1. Prepare 0.8% agarose gels using TAE buffer as recommended by the manufac-turer. Add EtBr (final concentration of 0.1 µg/mL) to the agarose prior to pouring
the gel.
2. Loading of subgels: In the first and last lanes, load 0.5 µg of molecular weight
ladder prepared by adding the ladder DNA, 6 µL of 6X loading buffer, and
water to a total volume of 36 µL. Leave the adjacent lanes to the ladder blank
to prevent spillover into patient or control lanes. In the remaining lanes, load
the control samples and patient digests. Load 10  µg of DNA diluted in 1X
loading buffer.
3.3.5. Southern Transfer
This method is based on ref. 24 .
1. Photograph the gel at 60–62 cm, f-stop 5.6, for 2 s to check the digest quality and
amount of DNA per lane.
2. Depurinate the DNA in 500 mL of 0.1 N HCl for 30 min on a shaker platform.
Follow depurination with rinsing with tap water.
3. Denature the DNA in 500 mL of Blotting Solution II with shaking for 15 min.
4. Neutralize the DNA in 500 mL of Blotting Solution III with shaking for 30 min.
5. Transfer the DNA to Zetabind membrane as described by the manufacturer. The
layers in the Southern transfer setup are as follows:
a. Layer 1: Two sheets of 3MM Whatman paper to serve as wicks, prewet in
transfer solution.
b. Layer 2: Denatured/neutralized gel, bottom side (DNA side) up.
c. Layer 3: Nylon membrane (Zetabind).
d. Layer 4: Two 3MM Whatman paper wicks.
e. Layer 5: One dry 3MM Whatman paper wick.
f. Layer 6: Stack of dry paper towels, 3–4 in.
g. Layer 7: Evenly distributed weight, e.g., glass plate with two 500-mL reagent
bottles on top or a glass pan.
6. Transfer overnight. Transfer can be accomplished in 4 h by removing soaked
paper towels every 30 min and replacing with dry ones. Do not allow the nylon
membrane to dry out.
7. Perform UV crosslinking of the membrane in a crosslinking apparatus as
described by the manufacturer.
8. Prehybridize the membrane in 150 mL of prehybridization solution at 55°C for
30 min to overnight.
Detection of Clonal Genes in Lymphomas and Leukemias 203
3.3.6. Radiolabeling of Probe
1. Perform nick translation with the Gibco-BRL nick translation system and
_32P-dCTP as described by the manufacturer. Oligolabel smaller probes and
inserts as described by the manufacturer.
2. Isolate radiolabeled probe with the Pharmacia nick column as described by the
manufacturer.
3. To quantitate the radiolabeled DNA, determine the volume in microliters of probe
that is required for 2 × 107 cpm; also calculate the specific activity (disintegra-tions per minute/micrograms).
3.3.7. Hybridization
1. Into each hybridization bottle, add 10 mL of hybridization solution and the
membranes.
2. For every 10 mL of hybridization solution, add boiled probe/salmon sperm mix
(the mixture is prepared by boiling the volume of radiolabeled probe required for
2 × 107 cpm/filter with 100 µL salmon sperm DNA). Note: Do not drip the con-centrated probe onto the filter because background hot spots may occur.
3. Hybridize at 42°C for 18 h overnight.
4. Posthybridization washes are as follows:
a. Wash 1: 500 mL of 2X SSC + 0.1% SDS (made fresh) at room temperature
with shaking for 15 min.
b. Washes 2 and 3 (stringency washes): 500 mL of 0.1X SSC + 0.1% SDS, at
60°C except for PFLI (bcl2), which is washed at 55°C. Perform each wash
with shaking for 30 min.
3.3.8. Autoradiography
Autoradiography is performed by placing the membrane in a plastic bag
(e.g., Seal-a-Meal) and then in an X-ray cassette with film at –70°C for 3 d.
Note: Do not let the membranes dry out.
3.3.9. Interpretation of Southern Blot Results
The presence of germline bands is interpreted only as a negative result. Any
rearrangement of the Ig or TCR genes will appear as a band with variable
intensity and a size different from the germline band. The intensity of the rear-ranged band is proportional to the percentage of clonal cells in the sample
being tested. Generally, a positive result will be seen with both enzymes and is
usually also positive by PCR. Two rearranged bands in a single lane hybrid-ized with one gene probe may be a result of two coexisting clones within the
same tissue or of rearrangements of both alleles for that specific gene (25) . A
marked difference in intensity of the two bands implies different dosages of
204 Mitha and McGlennen
each rearranged gene and is likely to represent more than one clone. The fol-lowing guidelines are useful for interpretation of Southern blots:
1. False-negative results with Southern blotting usually occur because the clonal
population is below the sensitivity level of Southern analysis or because of tis-sue-sampling error. False-positive results may also arise from either a partial
digestion or gene polymorphisms. A common example of partial digestion is
EcoRI-digested DNA hybridized with a TCR`probe. One usually sees an 8.5-kb
band in addition to the usual 11- and 4-kb germline bands, which is owing to a
relatively resistant EcoRI site (3 ,21) . Additionally, certain rearrangements may
produce very large restriction fragments, and hence the DNA may transfer poorly,
causing these rearrangements to be missed. The use of multiple enzyme-probe
combinations helps overcome this problem.
2. Several “benign” clonal lymphoproliferative conditions exist and include immu-nodeficiency settings associated with Wiscott-Aldrich syndrome and autoimmune
deficiency syndrome, angioimmunoblastic lymphadenopathy, posttransplant
immunosuppression, congenital immunodeficiency, and benign monoclonal
gammopathy. It is important to be aware of these conditions to avoid erroneous
diagnoses of malignancy.
Fig. 6. PCR gel and slot blot in a patient with mantle cell lymphoma, demonstrating
the classic  bcl1 gene (t [11;14]) rearrangement. Abbreviations: MW = molecular
weight marker; Pat = patient sample.
Detection of Clonal Genes in Lymphomas and Leukemias 205
3. Restriction fragment length polymorphisms (RFLPs) and partially digested DNA
may produce nongermline bands that are not owing to rearranged antigen recep-tor genes. Thus, laboratories must be aware of RFLPs generated by routinely
used restriction enzymes and DNA probes. It is recommended that two enzymes
be used in combination with each probe. If results from the two digests are not
consistent, a third digest should be performed. This will eliminate misinterpreta-tion of uncommon RFLPs or partial digests as a clonal rearrangement.
4. Comigration occurs when both the germline band and the rearranged band
migrate to the same point in the gel. Comigration of rearranged bands with
germline bands can be a major problem when using the restriction enzyme HindIII
in combination with the constant region probe for the TCR`chain. Restriction
enzymes that produce large germline fragments are more likely to comigrate with
nongermline bands. DNA should be digested with a second enzyme to resolve
comigrations.
Fig. 7. Status post autologous bone marrow transplant for Hodgkin disease of a 39-yr-old female. The patient now presents with a lung mass suspicious for lymphoma.
The PCR gel demonstrates the presence of EBV DNA, and the Southern blot confirms
the clonal nature of the EBV DNA. The molecular findings in conjunction with mor-phology are suggestive of a posttransplant lymphoproliferative disorder. Abbrevia-tions: MW = molecular weight marker; Pat = patient sample.
206 Mitha and McGlennen
4. Notes
1. The quality of DNA affects the PCR-based assays. PCR detection of clonality in
formalin-fixed DNA is 15% less sensitive at detecting IgH gene clonality than in
unfixed DNA (26 ,27) . Tissues obtained in mercurial-based fixations may also be
difficult to amplify.
2. Polyacrylamide gels, 6–8% final concentration, provide the best resolution and
clearest banding patterns.
3. The use of multiple primers improves overall clonality detection rates in the non-Hodgkin lymphomas. The use of both  ` and a chain assays detects rearrange-ments in 90% of T-cell lymphomas. The sensitivity varies depending on the
background cell population.
4. Specific PCR and data protocols for TCR gene rearrangements are not as well
documented as those for B-cell gene rearrangements. However, several studies
suggest that virtually all T-cell chronic lymphocytic and prolymphocytic leuke-mias should have clonal TCR` and TCRa rearrangements (19 ,28) . T-cell ALL
and lymphoblastic lymphomas show TCR` and TCRa rearrangement in more
than 90% of cases  (29–35) . A corresponding IgH rearrangement is seen in
15–20% of these cases  (36) . Seventy to 80% of cases of mycosis fungoides/
Sezary syndrome demonstrate clonality with TCRs. In this case, clonal TCR-a
chain rearrangements are seen more frequently than TCR`chain rearrangements
(24 ,25 ,37) . Peripheral T-cell lymphomas almost always have clonal TCR-arear-rangement and often have a TRC-` rearrangement.
5. Other assays for which PCR is frequently used in our laboratory include the PCR-based assay for the detection of the t(11;14) translocation and Epstein-Barr virus
(EBV) DNA, as illustrated in Figs. 6 and 7.
References
1. Arnold, A., Cossman, J., Bakhshi, A., et al. (1983) Immunoglobulin gene rear-rangements as unique clonal markers in human lymphoid neoplasms. N. Engl. J.
Med. 309, 1593–1599.
2. Collins, R. D. (1997) Is clonality equivalent to malignancy: specifically is immu-noglobulin gene rearrangement diagnostic of malignant lymphoma. Hum. Pathol.
28(7), 757–759.
3. Coad, J. E., Olson, D. J., Lander, T. A., et al. (1997) Molecular assessment of
clonality in lymphoproliferative disorders: II. T-cell receptor gene rearrange-ments. Mol. Diagn. 2, 69–81.
4. Coad, J. E., Olson, D. J., Lander, T. A., et al. (1996) Molecular assessment of
clonality in lymphoproliferative disorders: I. Immunoglobulin gene rearrange-ments. Mol. Diagn. 1, 335–355.
5. Lehman, C. M., Sarago, C., Nasim, S., et al. (1995) Comparison of PCR with
Southern hybridization for routine detection of immunoglobulin heavy chain gene
rearrangements. Am. J. Clin. Pathol. 103, 171–176.
Detection of Clonal Genes in Lymphomas and Leukemias 207
6. Aubin, J., Davi, F., Nguyen-Salomon, F., et al. (1995) Description of a novel FR1
IgH PCR strategy and its comparison with three other strategies for the detection
of clonality in B cell malignancy. Leukemia 9, 471–479.
7. Potter, M. N., Steward, C. G., Maitland, N. J., et al. (1992) Detection of clonality
in childhood B lineage acute lymphoblastic leukemia by polymerase chain reac-tion. Leukemia 9, 289–294.
8. Segal, G. H., Wittwer, C. T., Fishleder, A. J., et al. (1992) Identification of mono-clonal B cell populations by rapid cycle polymerase chain reaction: a practical
screening method for the detection of immunoglobulin gene rearrangements. Am.
J. Pathol. 141, 1291–1297.
9. Cleary, M. L., Meeker, T. C., Levy, S., et al. (1986) Clustering of extensive
somatic mutations in the variable regions of an immunoglobulin heavy chain gene
from a B-cell lymphoma. Cell 44, 97–106.
10. Deane, M., McCarthy, K. P., Wiedemann, L. M., et al. (1991) An improved
method for detection of B-lymphoid clonality by polymerase chain reaction. Leu-kemia 5, 726–730.
11. Kneba, M., Eick, S., Herbst, H., et al. (1991) Frequency and structure of
t(14;18) major breakpoint lesions in Non-Hodgkin’s lymphoma types according
to the Keil classification: analysis by direct DNA sequencing.  Cancer Res. 51,
3243–3250.
12. Ladanyi, M. and Wang, S. (1992) Detection of rearrangements of the bcl-2 major
breakpoint region in follicular lymphomas: correlation of polymerase chain reac-tion results with Southern blot analysis. Diagn. Mol. Pathol. 1, 31–35.
13. Horsman, D. E., Gascoyne, R. D., Coupland, R. W., et al. (1995) Comparison
of cytogenetic analysis, Southern blot analysis, and polymerase chain reaction
for the detection of t(14;18) follicular lymphoma.  Am. J. Clin. Pathol. 103,
472–478.
14. Ngan, B. Y., Nourse, J., and Cleary, M. L. (1989) Detection of chromosomal
translocation t(14;18) within the minor cluster region of bcl-2 by polymerase chain
reaction and direct genomic sequencing of the enzymatically amplified DNA in
follicular lymphomas. Blood 73, 1759–1762.
15. Segal, G. H., Scott, M., Jorgensen, T., et al. (1994) Primers frequently used for
detecting the t(14;18) major breakpoint also amplify Epstein-Barr virus. Diagn.
Mol. Pathol. 3, 15–21.
16. Klein, A., Zemer, R., Manor, Y., et al. (1997) Lymphoma with multi gene rear-rangement on the level of immunoglobulin heavy chain, light chains and T-cell
receptor ` chain. Am. J. Hematol. 56, 219–223.
17. Hanawa, H., Abo, J., Inokuchi, K., et al. (1995) Dual rearrangement of immuno-globulin and T-cell receptor genes in a case of splenic lymphoma with villous
lymphocytes. Leukemia Lymphoma 18, 357–360.
18. Hu, E., Weiss, L., Warnke, R., et al. (1989) Non-Hodgkins lymphoma containing
both B and T cell clones. Blood 70, 287–292.
19. Cossman, J., Uppenkamp, M., Sundeen, J., et al. (1988) Molecular genetics and
the diagnosis of lymphoma. Arch. Pathol. Lab. Med. 112, 117–127.
208 Mitha and McGlennen
20. Cheng, G., Minden, M. D., Toyonaga, B., et al. (1986) T cell receptor and
immunoglobulin gene rearrangements in acute myeloblastic leukemia. J. Exp.
Med. 163, 414.
21. Dick, F. R., Goeken, J. A., Kemp, J. D., et al. (1990) Kappa immunoglobulin light
chain gene rearrangement in a T lineage chronic lymphocytic leukemia. Am. J.
Clin. Pathol. 95, 702.
22. Ebrahim, S. A., Ladanyi, M., Desai, S. B., Offit, K., Jhanwar, S. C., Filippa,
D. A., Lieberman, P. H., and Chaganti, R. S. (1990) Immunohistochemical, molecu-lar, and cytogenetic analysis of a consecutive series of 20 peripheral T-cell lym-phomas and lymphomas of uncertain lineage, including 12 Ki-1 positive
lymphomas. Genes Chromosomes Cancer 2, 27–35.
23. Lust, J. A. (1996) Molecular genetics and lymphoproliferative disorders. J. Clin.
Lab. Anal. 10, 359–367.
24. Ralfkiaer, E., O’Conner, N. T. J., Crick, J., et al. (1987) Genotypic analysis of
cutaneous T cell lymphomas. J. Invest. Dermatol. 88, 762–765.
25. Bachelez, H., Bioul, L., Flageul, B., et al. (1995) Detection of clonal T cell recep-tor gamma gene rearrangements with the use of the polymerase chain reaction in
cutaneous lesions of mycosis fungoides and Sezary syndrome. Arch. Dermatol.
131, 1027–1031.
26. Achille, A., Scarpa, A., Montresor, M., et al. (1995) Routine application of poly-merase chain reaction in the diagnosis of monoclonality of B-cell lymphoid pro-liferations. Diagn. Mol. Pathol. 4, 14–24.
27. Chen, Y.-T., Whitney, K. D., and Chen, Y. (1994) Clonality analysis of B cell
lymphoma in fresh frozen and paraffin embedded tissues: the effects of variable
polymerase chain reaction parameters. Mod. Pathol. 7, 429–434.
28. Knowles, D. M. (1989) Immunophenotypic and antigen receptor gene rearrange-ment analysis in T cell neoplasia. Am. J. Pathol. 134, 761–785.
29. Gill, J. I. and Gulley, M. L. (1994) Immunoglobulin and T cell receptor gene
rearrangement. Hematol. Oncol. Clin. 8, 751–770.
30. Perl, A., DiVincenzo, J., Ryan, D., et al. (1990) Rearrangement of the T cell
receptor alpha, beta, and gamma chain genes in chronic lymphocytic leukemia.
Leukemia Res. 14, 131–137.
31. Davey, M. P., Bongiovanni, K. F., Kaulfersch, W., et al. (1986) Immunoglobulin
and T cell receptor gene rearrangement and expression in human lymphoid leuke-mia cells at different stages of maturation.  Proc. Natl. Acad. Sci. USA 83,
8759–8763.
32. Van Dongen, J. J. M. and Wolvers-Tettero, I. L. M. (1991) Analysis of immuno-globulin and T cell receptor genes. Part 11: possibilities and limitations in the
diagnosis and management of lymphoproliferative diseases and related disorders.
Clin. Chem. Acta. 198, 93–174.
33. De Villartay, J. P., Pullman, A. B., Andrade, R., et al. (1989) Gamma/delta lin-eage relationship within a consecutive series of human precursor T cell neoplasms.
Blood 74, 2508–2518.
Detection of Clonal Genes in Lymphomas and Leukemias 209
34. Korsmeyer, S. J., Arnold, A., Bakhshi, A., et al. (1983) Immunoglobulin gene
rearrangement and cell surface antigen expression in acute lymphocytic leuke-mias of T cell and B cell precursor origins. J. Clin. Invest. 71, 301–313.
35. Falini, B., Flenghi, L., Fagioli, M., et al. (1989) T lymphoblastic lymphomas
expressing the non-disulfide-linked form of the T cell receptor.  Blood 74,
2501–2507.
36. Kitchingman, G. R., Rovigatti, U., Mauer, A. M., et al. (1985) Rearrangement of
immunoglobulin heavy chain genes in T-cell acute lymphoblastic leukemia. Blood
65, 725–729.
37. Mielke, V., Staib, G., Boehncke, W. H., et al. (1994) Clonal disease in early cuta-neous T cell lymphoma. Dermatol. Clin. 12, 351–360.
Bone Marrow Transplant Engraftment 211
17
Monitoring of Bone Marrow Transplant Engraftment
Kristine P. Woronzoff-Dashkoff and Ronald C. McGlennen
1. Introduction
1.1. Background of Engraftment Testing
Bone marrow transplantation is used as a primary treatment for many dis-eases, including leukemia, lymphoma, and inborn errors of metabolism. The
procedure involves ablation of the recipient’s bone marrow by chemotherapy
and/or radiation therapy, followed by transplantation of harvested bone mar-row. In autologous bone marrow transplantation (BMT), the patient’s own
marrow is harvested and treated to remove malignant cells before it is replaced
into the patient. In allogeneic BMT, bone marrow is obtained from a donor
who is a close antigenic match to the patient. In either case, the goal of BMT is
full, permanent replacement of the recipient’s original bone marrow by donor
hematopoietic elements.
Patients who have had bone marrow transplants are carefully monitored for
evidence of disease remission or relapse. In addition, particularly in the early
posttransplant period, it is critical to establish the extent of engraftment of the
transplanted marrow. In patients who have received autologous bone marrow
transplants, a morphologic estimation of bone marrow cellularity is usually
adequate for determining engraftment status. In patients who have received
allogeneic bone marrow transplants, however, bone marrow cellularity may
not reflect true engraftment, because morphologic examination alone cannot
determine whether cells are of donor or recipient origin.
Accurate identification of cell origin is achieved by examining cellular DNA
at either the chromosomal or at the molecular level. Cytogenetic evaluation
affords examination of the full complement of chromosomes; therefore, in
addition to monitoring engraftment status, this method may detect additional
211
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
212 Woronzoff-Dashkoff and McGlennen
abnormalities of clinical importance, such as new or persistent chromosomal
aberrations that portend a certain prognostic significance. In this case, mor-phologic features unique to the donor’s or recipient’s chromosomes, including
differences in sex chromosomes, are followed serially in the posttransplant
period. However, the sensitivity of this method is relatively low. Furthermore,
regenerating marrow specimens, which are often very hypocellular, may not
contain enough dividing cells for accurate cytogenetic analysis.
Molecular evaluation of engraftment involves the evaluation of anonymous
DNA markers, a technique long used in forensic DNA testing. This testing is
based on the fact that the human genome contains small sequences of repetitive
DNA, or markers, located mainly at the ends and centers of chromosomes.
These markers are highly polymorphic; that is, they may vary considerably in
size among individuals. If a bone marrow transplant recipient carries a marker
that is sized differently than the corresponding donor marker, this marker may
be used in the posttransplant period to determine whether the cells repopulat-ing the recipient’s bone marrow are of donor or recipient origin. Although this
type of testing does not allow examination of the entire chromosomal comple-ment, its high sensitivity makes it uniquely suitable for routine laboratory
evaluation of engraftment status.
In the past, the first-line method of monitoring bone marrow transplant
engraftment involved examination of restriction fragment length polymorphisms
(RFLPs) by Southern transfer analysis. In this technique, donor and recipient mark-ers with different enzymatic restriction sites, and thus different molecular weights,
are analyzed by Southern transfer blotting (Fig. 1). The results obtained are usually
relatively unambiguous, with few artifacts, allowing simple interpretation. How-ever, the amount of DNA required for this method of testing is relatively large, the
procedure is time-consuming, and the method is inherently less sensitive than poly-merase chain reaction (PCR)-based methods. Southern transfer analysis for RFLPs
is still used occasionally, particul-arly when PCR analysis fails to detect informa-tive alleles (see Subheading 3.3.).
Currently, the engraftment analysis method used by many laboratories is
PCR amplification of genomic DNA. This method affords several advantages
over RFLP testing, including increased sensitivity, smaller sample require-ments, simpler test setup, shorter turnaround time, elimination of restriction
enzymes and radioisotopes, and substantial cost savings  (1 ,2). As in RFLP
analysis, the targeted DNA regions are highly polymorphic, repetitive DNA
sequences. These sequences may be classified according to the number of
nucleotides in each repeated segment, and the length of the entire array of
repeats. Intermediately sized units (often 0.1–20 kb long) of tandem repeat
sequences are termed minisatellites. Under this category fall the variable num-ber tandem repeat regions (VNTRs), in which each repeat sequence is between
Bone Marrow Transplant Engraftment 213
5 and 64 nucleotides in length. VNTRs are used in both RFLP and PCR analy-sis (Fig. 2). Small units (<0.01 kb long) of tandem repeat sequences are termed
microsatellites. Certain types of microsatellites, called short tandem repeat
regions (STRs), contain very small repeat sequences of 1–4 nucleotides in
length. STRs are evaluated using PCR, for which assay components are now
available in kit format (Fig. 3).
1.2. Laboratory Approach to Engraftment Testing
For analysis of bone marrow transplant engraftment, a simple scheme of
laboratory testing is suggested (Fig. 4). Prior to BMT, donor and recipient DNA
is tested, with the goal of finding an informative marker suitable for following
in the posttransplant period. An informative marker is defined as one for which
at least one unique allele is present in the donor/recipient pair. The ideal marker
would show as many independently segregating alleles as possible, i.e., two
Fig. 1. (A) Diagram of an RFLP. Digestion of allele 1 with a restriction endonu-clease produces a single 7-kb fragment, whereas digestion of allele 2 with the same
restriction endonuclease produces two fragments, measuring 5 and 2 kb. (B) Diagram
of the corresponding possibilities on Southern blotting. Lane 1: Donor (homozygous
for allele 1); lane 2: recipient, pretransplant (homozygous for allele 2); lane 3: recipi-ent, posttransplant, with 50% engraftment of donor marrow; equal amounts of donor-specific and recipient-specific allele are present; lane 4: recipient, posttransplant, with
75% engraftment of donor marrow; relatively more donor-specific allele than recipi-ent-specific allele is present; lane 5: recipient, posttransplant, with equal amounts of
recipient-specific allele and previously unidentified allele. Such results may be spuri-ous (e.g., due to specimen mixup) or attributable to third-party engraftment (e.g., the
patient has received a recent blood transfusion or another bone marrow transplant
from a different donor).
214 Woronzoff-Dashkoff and McGlennen
unique alleles in the donor and two unique alleles in the recipient (abbreviated
D2R2). However, because only a limited number of differently sized alleles have
been described for each marker, it is likely that the donor and recipient will have at
least one allele size in common. At the minimum, at least one unique—preferably
donor-specific—allele must be found in the donor/recipient pair.
In our laboratory, pretransplant specimens from donor and recipient are first
analyzed by PCR for VNTR markers. In the past, analysis of PCR products
was performed manually. The products were electrophoresed on a polyacryla-mide gel, which was stained and photographed for visual analysis (Fig. 2).
Currently, our laboratory uses the ABI373 GeneScan DNA sequencing system
(see Subheading 2.2.) for automated quantification of allele size. Briefly,
fluorescently tagged primers are used to amplify genomic DNA from the speci-men. The PCR products are electrophoresed on a polyacrylamide gel, which is
then analyzed by the ABI instrument using laser-based technology. The sizes
of the donor and recipient alleles are determined by comparing the PCR prod-ucts with internal standards, and a histogram is produced (Fig. 3). Most speci-mens show one or more unique alleles using this method, and therefore no
further pretransplant testing is necessary.
VNTR analysis is used routinely in our laboratory because of our extensive
experience with these markers; in addition, many of our patients’ initial
pretransplant screening assays were performed using VNTR markers. Labora-tories new to bone marrow transplant engraftment analysis may elect to use
Fig. 2. Polyacrylamide gel: PCR analysis of bone marrow engraftment using
ColIIA1 marker. Lane 1, Donor blood; lane 2, recipient blood, pretransplant; lane 3,
control mixture (75% D, 25% R); lane 4, control mixture (90% D, 10% R); lane 5,
recipient marrow, posttransplant (100% engraftment).
Bone Marrow Transplant Engraftment 215
STR markers. If VNTR testing is not informative, the next line of testing in our
laboratory is PCR for STRs. In the unlikely event that no STR marker is infor-mative, Southern transfer analysis is used to identify RFLPs.
The second major segment of engraftment testing occurs following BMT
(Fig. 4). DNA from the recipient’s blood and bone marrow is amplified
by PCR, and the allele sizes in each previously selected marker are deter-mined.
These allele sizes are compared to those in the recipient’s pretransplant mar-row and the donor’s marrow. The percentage of donor engraftment is
determined semiquantitatively by comparing the quantity of donor-specific
allele to the total quantity of donor-plus recipient-specific alleles detected (see
Subheading 3.3.2.). The quantity of allele may be measured by visual estima-tion of band density, fluorescent quantification, or other methods.
Whichever type of assay is chosen for monitoring of engraftment status, it
must be carefully optimized for each marker. The reaction must be robust, par-ticularly since the quantity of DNA available in the posttransplant period may
be very small. Hence, sensitivity controls that employ patient-derived mixes of
donor and recipient cells extracted for DNA mixed in prescribed ratios (5%,
10%) ensure that interassay comparison are meaningful and precise. In addi-tion, accurate, reliable quantification of the percentage of donor and recipient
allele is necessary. In our experience, clinicians place more importance on the
trend of engraftment in a patient, rather than precise numerical quantification
of engraftment at any one particular time. Thus, we report results as a specific
range of percentage of engraftment, rather than as fixed numerical values.
Fig. 3. Histogram of donor and recipient pretransplant alleles showing polymor-phisms in 4 STRs: vWA (D1R1), TH01 (D1R0), TPOX (D1R1), and CSF1PO.
216 Woronzoff-Dashkoff and McGlennen
2. Materials
Markers examined in our assay include the following VNTRs and STRs:
1. Apolipoprotein B (ApoB): This autosomal codominant VNTR lies on the short
arm of chromosome 2 and is located in the 3vflanking region of the 42-kb ApoB
gene. Twelve alleles have been identified, ranging in size from 460–600 bp. The
ApoB VNTR has a heterozygosity of 75% (3) .
2. Collagen Type II alpha 1 (ColIIA1): This autosomal codominant VNTR is
located at 12q14.3, in the 3vflanking region of the 32-kb Type II Collagen gene.
Several alleles have been identified, ranging in size from 600–700 bp. The
COLIIA1 VNTR has a heterozygosity of 81% (4 ,5) .
Fig. 4. Flow chart showing clinical engraftment assessment and accompanying labo-ratory testing.
Bone Marrow Transplant Engraftment 217
3. D1S80: This autosomal codominant VNTR is located on chromosome 1. Sixteen
alleles have been identified, ranging in size from 300–700 bp. The D1S80 VNTR
has a heterozygosity of 80.8% (6) .
4. pYNZ22: This autosomal codominant VNTR is located at 17p23. More than 10
alleles have been identified, ranging in size from 170 –870 bp. The pYNZ22 VNTR
has a heterozygosity of 86% ([7] ; Bruce Balzar, personal communication).
5. CF-17b: This is a dinucleotide intronic STR found in the CFTR gene at chromo-some 7q31. Thirty-three alleles have been identified, ranging in size from
167– 414 bp (8) .
6. CSF1PO: This STR is found within the human c-fms proto-oncogene for CSF-1
receptor gene (HUMCSF1PO) located at 5q33.3-34. Ten alleles have been iden-tified, ranging in size from 295–327 bp.
7. TPOX: This STR is found within the human thyroid peroxidase gene
(HUMTPOX) located at 2p25-1-pter. Eight alleles have been identified, ranging
in size from 224–252 bp.
8. TH01: This STR is found within the human tyrosine hydroxylase gene
(HUMTH01) located at 11p15.5. Eight alleles have been identified, ranging in
size from 179–203 bp.
9. vWA (formerly vWF): This STR is found within the human von Willebrand fac-tor gene (HUMVWFA31) located at 12p12-pter. Ten alleles have been identified,
ranging in size from 139–167 bp.
2.1. Specimen Collection and Processing
Prior to BMT, patient and donor blood specimens are collected. The DNA
from these specimens is amplified by PCR for identification of polymorphism.
These pre-BMT results are later correlated with results from post-BMT speci-men testing.
2.1.1. Blood Specimens
1. Volume: For pre-BMT specimens, collect 30 mL of blood. For post-BMT speci-mens, the volume of blood necessary is dependent on the white blood cell (WBC)
count. For WBC (109/L) of >1.0, 0.5, 0.25, and 0.1 the blood volume is 10, 20,
30, and 50 mL, respectively.
2. Preservative: Acid citrate dextrose (ACD) is preferred, but EDTA is acceptable.
2.1.2. Bone Marrow Specimens
1. Volume: For pre-BMT, bone marrow is not required (blood is recommended for
this procedure). For post-BMT, 5 mL is required (or as little as 2 mL, if marrow
is not markedly hypocellular).
2. Preservative: Collect in a syringe to which ACD has been added (EDTA is
acceptable).
218 Woronzoff-Dashkoff and McGlennen
2.1.3. Tissue Specimens
A minimum volume of 5 mm3 must be delivered to the laboratory within 1 h
of collection or quick frozen in liquid nitrogen or dry ice. Specimens from
outside the hospital should be transported frozen on dry ice.
2.2. Equipment and Supplies
1. Perkin-Elmer Thermocycler 480 (Perkin-Elmer, Norwalk, CT).
2. ABI373 DNA sequencing system (Applied Biosystems, Foster City, CA).
3. The following supplies from Applied Biosystems:
a. GENESCAN™ 672 Software (cat. no. 672-10).
b. Genotyper™ Software (cat. no. 401614).
c. 6-cm S Notched Glass Plate (cat. no. 401623).
d. 6-cm S Plain Plate (cat. no. 401624).
e. 6-cm S Gel Spacer, Pair (cat. no. 401625).
f. GENESCAN 24-tooth comb (cat. no. 401444).
g. GENESCAN 36-tooth comb (cat. no. 401497).
h. PRISM™ GENESCAN-500 Tamra Size Standard (cat. no. 401733).
i. PRISM GENESCAN-2500 Tamra Size Standard (cat. no. 401545).
4. Acrylamide/bis powder (19 1) (cat. no. 161-0120; Bio-Rad, Hercules, CA).
5. Formamide, molecular biology grade (cat. no. BP227-100; Fisher, Fair
Lawn, NJ).
6. TEMED (cat. no. 161-0800; Bio-Rad).
7. Ammonium persulfate (cat. no. 161-0700; Bio-Rad).
8. Urea, ultrapure (cat. no. 15505-035, Life Technologies, Rockville, MD).
9. AG 501X8 resin (cat. no. 143-6424; Bio-Rad).
10. Sterilization filter unit (0.45-µm, 115 mL) (cat. no. 245-0045; Nalge, Roch-ester, NY).
11. Scotch™ electrical tape) (cat. no. 1739-7; 3M, St. Paul, MN).
12. Tris crystallized free base (cat. no. BP152-5; Fisher).
13. Boric acid (cat. no. A73-500; Fisher).
14. EDTA disodium salt (cat. no. BP 120-1; Fisher).
15. Blue dextran powder (cat. no. D-5751; Sigma, St. Louis, MO).
16. Spectrophotometer (Beckman, Palo Alto, CA, or other supplier).
17. Taq DNA polymerase in storage Buffer A, 5  µ/µL (cat. no. M1865; Promega,
Madison, WI).
18. Bulk dNTPs (40  µM) 100 mM/L each of dATP, dCTP, dGTP, dTTP (cat. no.
U1240; Promega).
19. Primers for VNTR and STR sequences: Primers for the ApoB, ColIIA1, D1S80,
pYNZ22, and CF-17b markers are custom synthesized and fluorescently labeled
by Microchemical Facility Laboratory, UMHC Institute of Human Genetics,
Minneapolis, MN. The sequences are as follows:
a. ApoB3v(A) (20mer): 5v-CCT-TCT-CAC-TTG-GCA-AAT-AC-3v.
b. ApoB5v(S) (20mer): 5v-ATG-GAA-ACG-GAG-AAA-TTA-TG-3v.
Bone Marrow Transplant Engraftment 219
c. COL2A3v(A) (20mer): 5v-GTC-ATG-AAC-TAG-CTC-TGG-TG-3v.
d. COL2A5v(S) (20 mer): 5v-CCA-GGT-TAA-GGT-TGA-CAG-CT-3v.
e. D1S80w (28mer): 5v-GAA-ACT-GGC-CTC-CAA-ACA-CTG-CCC-GCC-G-3v.
f. D1S80c (29 mer): 5v-GTC-TTG-TTG-GAG-ATG-CAC-GTG-CCC-CTT-GC-3v.
g. pYNZ223v(A) (22mer): 5v-GCC-CCA-TGT-ATC-TTG-TGC-AGT-G-3v.
h. pYNZ225v(S) (22mer): 5v-GGT-CGA-AGA-GTG-AAG-TGC-ACA-G-3v.
i. CF-17b3v(A) (20mer): 5v-GCT-GCA-TTC-TAT-AGG-TTA-TC-3v.
j. CF-17b5v(S) (20mer): 5v-AAA-CTT-ACC-GAC-AAG-AGG-AA-3v.
Primers for the CSF1PO, TPOX, TH01, and vWA STRs are obtained from
Promega.
2.3. Reagents
1. MilliQ Water (double-distilled H2O): MilliQ is the trade name of the water sys-tem purchased from Millipore. This deionized water is treated with activated car-bon and deionization cartridges, and filtered to remove microorganisms larger
than 0.22 µm. It meets CAP Class II Water Requirements when stored in poly-ethylene carboys. If microbial-free water is needed, a bottle of this water is auto-claved and is referred to as sterile water.
2. PCR master mix: A preparation for either 500 or 2000 reactions is made by com-bining the reagents given in Table 1.
3. Stock primers (100 pmol/µL): Use the following formula to calculate the amount
of filtered, sterilized water to add to the primers (store at –20°C):
Yield (µg)/mol wt (µg/µmol) × (106) pmol/µmol ×µL/100 pmol
= volume needed for resuspension
4. Working primers (2.5 pmol/µL): Add 25 µL of each stock primer and dilute to
1000 µL with sterile water. Store at –20°C.
Table 1
PCR Cocktail for BMT Engraftment Testing
Component 1 rxna (mL) 500 rxn (mL) 2000 rxn (mL)
Peg 0.0425 21.25 85.0
dATP 0.004 2.0 8.0
dCTP 0.004 2.0 8.0
dGTP 0.004 2.0 8.0
dTTP 0.004 2.0 8.0
10X Buffer 0.010 5.0 20.0
Total volume 0.0685 34.25 137.0
arxn = reaction.
220 Woronzoff-Dashkoff and McGlennen
5. Working PCR mix: Prepare a mix for the total number of reactions plus one based
on the amount of DNA template to be used. For 4 µL of DNA template, prepare
the mix as follows: 40.25  µL of PCR master mix, 3.0  µL of MgCl2 (25 mM),
2.5 µL of SIA primer, and 0.25  µL of  Taq polymerase (for a total volume of
46.0 µL).
6. PCR control: A sample containing water in place of specimen is included with
each thermocycler run. This controls for the presence of contaminating DNA in
the reaction reagents.
7. Sensitivity controls: Two cell lines are mixed at a ratio of 95 5 and 10 90. These
product mixes, which determine the sensitivity of the assay, are run as controls
on every gel that includes an engraftment analysis. The sensitivity for a par-ticular specimen is that of the lowest percentage of detectable control, usually
5 or 10%.
3. Methods
3.1. Isolation of DNA
Isolate DNA from the specimen following standard laboratory protocol (see
Note 1).
3.2. Dilution of Genomic DNA
3.2.1. Calculating Amount of DNA to Dilute
3.2.1.1. PRETRANSPLANT RECIPIENT AND DONOR SAMPLES
Because pretreatment recipient and donor samples will be used in every
patient analysis, a dilution of a large quantity of DNA is made (typically
30 µg). However, if the amount of sample is limited, avoid diluting all of it. If
the donor and recipient are not polymorphic by PCR testing, DNA will be
needed for Southern analysis. To determine the volume of DNA needed for
30 µg, take the inverse of the concentration in micrograms/milliliter and multi-ply by 30,000:
1/(µg/mL) × 30 µg × 1000 µL/mL = µL for 30 µg
To determine the amount of H2O to add, dilute 30 µg of DNA into a total
volume of 450 µL to achieve the proper concentration. Then subtract the vol-ume needed for 30 µg from 450.
3.2.1.2. POSTTRANSPLANT RECIPIENT SAMPLES
Because posttransplant recipient samples will be analyzed only once or
twice, a small dilution is made (typically 7.5  µg). If pretransplant analysis
revealed at least one donor-specific and one recipient-specific allele, Southern
transfer analysis will not be required, and the entire posttransplant sample may
Bone Marrow Transplant Engraftment 221
be diluted. If this is not the case, avoid diluting the entire sample. To determine
the amount of DNA needed for 7.5 µg, take the inverse of the concentration in
micrograms/microliter and multiply by 7500:
1/(µg/mL) × 7.5 µg × 1000 µL/mL = µL for 7.5 µg
To determine the amount of H2O to add, dilute 7.5 µg of DNA into a total
volume of 112.5  µL to achieve the proper concentration. Then subtract the
volume needed for 7.5 µg from 112.5.
3.2.2. Making Dilutions for PCR
1. Use 1.5-mL microcentrifuge tubes labeled with the sample number.
2. Make dilutions using designated pipettors and sterile water reserved for
PCR use.
3. Dilute the samples.
4. Vortex the dilutions after making them. Revortex prior to beginning PCR.
3.3. Setting Up Samples According to Type of Analysis
3.3.1. Pretransplant Polymorphism Determination
Pretransplant donor and recipient samples are screened for polymorphisms
in the VNTRs and STRs listed previously in order to find an informative
marker. A marker is considered informative when at least one allele with a
unique number of repeats is found in the donor or recipient. These unique
alleles have different molecular weights and are thus separable on gel
electrophoresis.
Three VNTRs, ApoB, ColIIA1, and D1S80, are evaluated first; most recipi-ent–donor pairs will be polymorphic at one or more of these VNTRs. If no
polymorphism is identified, one additional VNTR (pYNZ22) and an STR
(CF-17b) are examined. If these markers prove uninformative, additional STRs
(CSF1PO, POX, TH01, and vWA) are examined. If these markers are also
uninformative, Southern transfer analysis for RFLPs may be necessary for
polymorphism determination and engraftment analysis.
3.3.2. Posttransplant Engraftment Analysis
Patient samples are analyzed using the markers determined to be informa-tive in the pretransplant polymorphism screen (see Subheading 2.).
1. Add reagents and specimens to tubes; place the tubes in a thermocycler.
2. Into a 0.6-µL microcentrifuge tube, add 85  µL of the working PCR mix (see
Subheading 3.2.), and then add 15µL of DNA template. Vortex the tubes, spin
briefly in a microcentrifuge, and place the tubes in a thermal cycler. Run the
thermal cycler according to predetermined PCR parameters (see Note 2).
222 Woronzoff-Dashkoff and McGlennen
3. Perform electrophoresis according to the ABI373 GeneScan Electrophoresis Pro-cedure using 1–2 µL of PCR product.
4. Analyze the GeneScan collection file according to the ABI373 GeneScan Analy-sis Procedure.
3.4. Formats for Reporting Results
3.4.1. Pretransplant Results
The allele sizes of the donor and the recipient for the informative marker are
recorded in the laboratory. Test results are reported to clinicians as “polymor-phism found” or “polymorphism not found.”
3.4.2. Posttransplant Results
The allele sizes of the recipient and the percentage of engraftment are
recorded in the laboratory. The results are reported to clinicians as ranges, from
the following choices: 0%, 1–10%, 11–25%, 26–50%, 51–75%, 76–99%,
90–94%, 95–99%, and 100%.
3.5. Interpretation of Results
3.5.1. Pretransplant Assays
A useful pretransplant assay will identify at least one unique allele in a
donor/recipient pair. Although some donors will possess two alleles that are
completely different from the recipient’s two alleles, this is not often the case.
Because engraftment testing by PCR is so sensitive, the presence of even one
unique allele in either the donor or recipient is sufficient for distinguishing
between the donor and the recipient.
The unique allele must, of course, be different enough in size to be distin-guished from its partner allele. The necessary size difference depends on the
system used to analyze the PCR products. In our computerized analysis sys-tem, alleles must be at least 4 bp different in size to be resolved by the com-puter. If one is analyzing the PCR products manually, one should choose a
marker that shows at least one unique band in either the donor or recipient that
is easily and distinctly distinguishable from its partner band.
3.5.2. Posttransplant Assays
In the monitoring of bone marrow transplant engraftment, it is essential to
obtain current clinical information about the patient. The date of the bone mar-row transplant should be noted, along with any current clinical information,
such as whether the specimen is sent for routine follow-up examination, or
whether there is a suspicion of disease relapse or graft loss. Morphologic inter-pretation of the specimen, if available, is also quite helpful, particularly when
molecular results are not in concordance with clinical information.
Bone Marrow Transplant Engraftment 223
One morphologic setting in which molecular testing may be particularly
useful is that of a markedly hypocellular marrow. In this situation, cells are too
few in number to accurately assess morphologically or cytogenetically.
Because PCR requires so little DNA to provide an informative result, engraft-ment testing in the molecular laboratory often provides valid, critical informa-tion to the clinician at this vulnerable stage of the posttransplant period.
Following transplant, testing is limited to those unique alleles identified on
pretransplant testing. In most cases, examination of a single marker provides
informative results. If at least one unique allele in each donor and recipient
(abbreviated D1R1) is present, the assay will provide informative results. How-ever, finding one unique allele in the donor and no unique recipient alleles
(D1R0) may also be informative, particularly in the setting of minimal (<25%)
engraftment. Likewise, finding no unique donor alleles but one unique recipi-ent allele (D0R1) is informative in the setting of near-total (>75%) engraft-ment. In the latter two scenarios, correlation of molecular engraftment studies
with bone marrow morphology is particularly crucial.
3.5.3. Semiquantitative Analysis of Data
Based on the amount of donor-specific allele (D) and recipient-specific
allele (R) present, the computer calculates the percentage of bone marrow trans-plant engraftment. This number is reported as a range to allow for gel-to-gel
variation (see Subheading 3.5.2.). The formulas for determining the percent-age of engraftment are different depending on whether the results show one
donor-specific allele and one recipient-specific allele (D1R1), one recipient-specific allele and no donor-specific alleles (D0R1), or one donor-specific
allele and no recipient-specific alleles (D1R0). For D1R1, the formula is D/R
+ D; for D0R1, the formula is 1 – (2R1/R1 + R2); and for D2R0, the formula is
2D2/(D1 + D2).
4. Notes
1. On occasion, posttransplant specimens may be sent for engraftment studies on
patients who have not had pretransplant polymorphism screening. In this situa-tion, tissues such as buccal scrapings or skin biopsies may be used for obtaining
DNA. However, special care must be taken when obtaining skin biopsies in order
to avoid excessive blood contamination, which could lead to amplification of
post-BMT donor leukocyte DNA. Another possible source of pretransplant
recipient DNA is cell lysate samples from the HLA-typing laboratory. These
specimens are used as is, without DNA extraction. Usually, only a very small
volume of this type of specimen is available; the sample is diluted only enough to
allow performance of the three most commonly polymorphic VNTRs: ApoB,
ColIIA1, and D1S80.
224 Woronzoff-Dashkoff and McGlennen
2. Because individual thermal cyclers may have very different characteristics, each
laboratory must optimize its own PCR parameters for each marker. In a
semiquantitative assay, it is particularly important to optimize not only the tem-peratures and segment lengths but also the cycle number. As shown in Fig. 5,
performing only a few cycles under or over the optimum number may result in
misrepresentation of the quantity of template. The following is a sample PCR
protocol for the ApoB marker: 94°C for 4 min; 19 cycles of 94°C for 1 min and
58°C for 6 min; 58°C for 4 min; followed by a 4°C soak.
References
1. Sreenan, J. J., Pettay, J. D., Tbakhi, A., et al. (1997) The use of amplified variable
number of tandem repeats (VNTR) in the detection of chimerism following bone
marrow transplantation: a comparison with restriction fragment length polymor-phism (RFLP) by Southern blotting. Am. J. Clin. Pathol. 107, 292–298.
2. Martinelli, G., Trabetti, E., Farabegoli, P., et al. (1997) Early detection of bone
marrow engraftment by amplification of hypervariable DNA regions.
Hematologica 82, 156–160.
3. Boerwinkle, E., Xiang, W., Fourest, E., et al. (1989) Rapid typing of tandem
repeated hypervariable loci by the polymerase chain: application to the
apolipoprotein B 3v hypervariable region.  Proc. Natl. Acad. Sci. USA 86,
212–216.
4. Seino, S. W. and Bell, G. I. (1990) Human Collagen Type II Alpha I (Col2AI)
Gene: variable number of tandem repeat polymorphism detected by gene amplifi-cation. Nucleic Acid Res. 18,  3102.
5. Stoker, N. G., Cheah, K. E., Griffin, J. R., et al. (1985) A highly polymorphic
region 3v to the human type II collagen gene. Nucleic Acid Res. 13, 4613–4622.
Fig. 5. Amplification kinetics of a mixture of 75% donor and 25% recipient tem-plate DNA, plotted as a function of cycle number. (Reprinted from ref. 9 with permis-sion from W. B. Saunders Company.)
Bone Marrow Transplant Engraftment 225
6. Budowle, B., Chakraborty, R., Giursti, A. M., et al. (1991) Analysis of the vari-able number of tandem repeat locus D1S80 by the polymerase chain reaction fol-lowed by high resolution polyacrylamide gel electrophoresis.  Am. J. Hum.
Genetics 48, 137–144.
7. Horn, T., Richards, B. R., and Klinger, K. W. (1989) Amplification of a highly
polymorphic variable number of tandem repeat segment by the polymerase chain
reaction. Nucleic Acid Res. 17, 2140.
8. Zielinski, J., Markiewicz, D., Rininsland, F., et al. (1991) A cluster of highly
polymorphic dinucleotide repeats in intron 17b of the cystic fibrosis transmem-brane conductance regulator (CFTR) gene. Am. J. Hum. Genet. 49, 1256–1262.
9. Crotty, P. L., Staggs, R. A., Porter, P. T., et al. (1994) Quantitative analysis in
molecular diagnostics. Hum. Pathol. 25, 572–579.
Direct Molecular Diagnosis of MEN1 227
227
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
18
Direct Molecular Diagnosis
of Multiple Endocrine Neoplasia Type 1
Elizabeth M. Petty, Michael Glynn, and Allen E. Bale
1. Introduction
1.1. Epidemiology and Clinical Phenotype
Multiple endocrine neoplasia type 1 (MEN1) is an autosomal dominant syn-drome characterized by the predisposition to develop both peptic ulcer disease
and a wide variety of endocrine tumors usually in adolescence and adulthood.
Specifically, hyperplasia and/or tumors (most often adenomas) of the parathy-roid, pancreatic islet cells, anterior pituitary, and adrenal cortical glands are
classically described in affected individuals who have MEN1 (1 ,2) . MEN1 is a
highly penetrant disorder whose onset is generally during adult life with the
occurrence of at least one, but most often more than one, of the aforementioned
tumors. The age-related penetrance of this disorder based on analysis in 63
unrelated kindreds is 7, 52, 87, 98, 99, and 100% by 10, 20, 30, 40, 50, and
60 yr, respectively (3) . The disorder is estimated to occur in approx 1 in 30,000
to 1 in 50,000 individuals. Most cases are associated with a positive family
history of the disorder, but new germline mutations have been identified in a
small percentage of individuals having a negative family history of the disor-der but classic features of MEN1 (3–7) .
Although the tumors most commonly described in MEN1 are not malignant,
they are often associated with medical conditions that require therapeutic
intervention. Symptoms in this disorder are therefore related to the biology of
the tumors occurring in any one affected individual. Symptoms are most often
related to hypersecretion of various hormones secondary to these tumors. Glu-cose intolerance, hypoglycemia, hypergastrinemia with severe peptic ulcer dis-
228 Petty, Glynn, and Bale
ease, galactorrhea, and full-blown Cushing syndrome are some of the symp-toms that can result from the many different tumors, including pancreatic islet
cell adenomas, gastrinomas, parathyroid adenomas, prolactinomas, glu-cagonomas, insulinomas, and vasointestinal peptide–producing tumors that
occur in this condition (1 ,2) . Epigastric pain with intractable peptic ulcers and/
or ulcerative gastritis (known as Zollinger Ellison syndrome) along with the
steatorrhea and diarrhea are especially problematic for individuals who have
gastrointestinal (GI) and pancreatic manifestations of the disease (8) .
Occasionally tumors will undergo malignant transformation, especially
those that are located in the pancreas, thymus, or bronchi (1 ,2). Thus, malig-nant tumors in the GI tract and the thymic-bronchic tract have been reported,
including bronchial carcinoids or carcinomas, thymic carcinoids, and duode-nal carcinoids. Cutaneous lesions, ovarian tumors, adrenocortical carcinomas,
and primary nervous tissue tumors including malignant schwannomas have
also been observed. In fact, angiofibromas, similar to those seen in tuberous
sclerosis, are present in many affected individuals. Thus, the effect of germline
mutations in the MEN1 gene is pleiotropic and there is both inter- and intrafamilial
variability in the phenotypic expression of the highly penetrant disorder.
Because MEN1 is a medical condition in which regular clinical screening
studies of at-risk individuals have proven useful in helping to reduce morbidity
and mortality related to the disease, owing to early symptom management and
medical or surgical intervention to minimize development of symptoms, DNA
or molecular diagnosis of MEN1 has demonstrated clear clinical utility in fami-lies in which the MEN1 disease phenotype segregates. At-risk individuals can
be evaluated for development of parathyroid, pituitary, and pancreatic islet cell
abnormalities by monitoring calcium, parathyroid hormone, prolactin, and gas-trin levels. Radiographic studies are useful to screen for pulmonary carcinoids.
1.2. MEN1 Tumor Suppressor Gene and Menin Protein
Studies of both linkage analysis and loss of heterozygosity pinpointed the
location of the MEN1 gene to chromosome 11q13 (9–15) . Studies of early loss
of heterozygosity suggested that the gene would be a tumor suppressor based
on the two-hit model according to Knudson (16) . In 1997, the MEN1 gene was
cloned (17) . The gene contains 10 exons spanning approx 9 kb of genomic
DNA. Exon 1 and the last 832 bp at the 3vend of exon 10 remained untranslated.
Therefore, there are nine coding exons that contribute to the 2.9-kb MEN1
transcript that is ubiquitously expressed in most human tissues. An additional
4-kb transcript was identified in the pancreas and the thymus.
The MEN1 gene is ubiquitously transcribed and encodes a 610 amino acid
(67 kDa) protein called menin (17) . Recently, clues to the possible function of
menin were reported  (18 ,19) . Menin is a nuclear protein that interacts with
Direct Molecular Diagnosis of MEN1 229
JunD. JunD may have transactivating and transrepressor activity through the
AP-1 transcription factor depending on the specific Fos partner or promoter,
but the actual role of menin and the tissue specificity of tumor involvement in
individuals with germline MEN1 mutations is not yet understood.
No one predisposing mutation has been identified in the coding region of
the MEN1 to account for the majority of the disease in patients studied to date.
However, founder mutations have been described in some populations stem-ming from geographic isolates (20–23) . Germline mutations have been reported
scattered throughout the coding region of the gene and include nonsense, mis-sense, and frameshift mutations. Mutations often predict premature truncation
of the protein owing to the nonsense or frameshift mutations. Missense muta-tions account for approx 30% of the cases (3–7 ,24 ,25) . To date, there are no
well-defined genotype/phenotype correlations or prognostic predictions that
can be made based solely on mutational analysis that are useful in the clinical
management of individuals. In general, it appears that germline mutations can
be identified in approx 90% of classic MEN1 families and slightly more than
80% of patients who themselves have isolated MEN1 with no known family
history (3–7 ,25) . MEN1 mutations have also been identified in a wide range
of related endocrine disorders and sporadic endocrine tumors  (5 ,7 ,26–35)
(see Note 1).
1.3. Considerations for DNA Diagnostic Testing of MEN1
Because the vast majority of MEN1 mutations reported to date are scattered
throughout the coding region of the gene, screening the nucleotide sequence of
each coding exon and adjacent intron/exon boundaries is a relatively robust
method for identifying mutations in the MEN1 gene (see Note 2). This type of
exon-by-exon mutation screening has the potential drawback, however, of
missing large deletions or rearrangements involving one allele. The MEN1
phenotype in its classic form does not display significant genetic heterogene-ity, making linkage analysis a useful diagnostic tool in large families in whom
multiple blood samples are available for analysis. In fact, in families in whom
a MEN1 mutation cannot be found by direct analysis, further studies to analyze
MEN1 linkage in these kindreds may be quite useful in helping to make a
diagnosis. Presumably in families in whom MEN1 mutations cannot be identified
in the coding regions, mutations may lie in promoter or other regions important to
the expression of the gene that have not been fully characterized in these patients
or there may be a large deletion or rearrangement involving several exons.
A variety of well-described methods to screen for point mutations have been
widely published in the primary scientific literature as well as in molecular
genetics or molecular biology protocol manuals (36–52) . These different strat-egies, often involving modifications of single-strand conformation polymor-
230 Petty, Glynn, and Bale
phism (SSCP) analysis, have varying degrees of sensitivity, depending on the
specific techniques employed. Given that there are only nine relatively small
coding exons in the MEN1 gene, direct sequence analysis proves to be the most
robust, cost-effective, and sensitive strategy to screen for unknown mutations.
Thus, polymerase chain reaction (PCR)-based manual or automated sequence
analysis provides the most useful means for the diagnosis of MEN1 in our
laboratories. Although there is no particular hot spot of mutations in MEN1,
early studies suggested that a slightly higher percentage of mutations have been
found in exons 2 and 10, potentially making these exons ideal starting places
for sequence-based mutation analysis. Exons 2 and 10 are the largest of the
MEN1 coding exons, which may help account for the relatively high number
of mutations in these regions. Given that there is no one exon where the major-ity of mutations are located, full sequencing through all coding exons in a
molecular diagnostic laboratory is recommended.
2. Materials
2.1. Reagents for DNA Sample Preparation
1. Reagents from peripheral blood using either standard laboratory protocols or
commercial DNA extraction/purification kits that reliably yield PCR quality
DNA, e.g., Puregene (Gentra Systems, Minneapolis, MN).
2.2. PCR Reagents
1. Taq polymerase (5 U/µL) (Perkin-Elmer).
2. 10X PCR Buffer II (Perkin-Elmer).
3. KlenTaq1 (Ab peptides) (25 U/µL) and buffer.
4. Cloned Pfu Polymerase (2.5 U/µL) (Stratagene).
5. LA 16 (15 parts KlenTaq and 1 part Pfu polymerase).
6. Double-distilled H2O (ddH2O).
7. dNTPs (2 mM).
8. Spermidine (50 mM).
9. MgCl2 (25 mM).
10. Dimethyl sulfoxide (DMSO).
11. 5 M Betaine.
12. PCR primers (10 µM) (see Table 1 for sequences) (see Note 3).
13. For direct mutation analysis: men1x2 (F and R), men1x3 (F and R), men1x4
(F and R), men1x5 + 6 (F and R), men1x7 (F and R), men1x8 (F and R), men1x9
(F and R), men1x10 (F and R).
2.3. Reagents for Agarose Gel Electrophoresis
1. Molecular biology grade agarose.
2. Ethidium bromide (10 mg/mL) stock.
3. 5X TBE buffer stock: 54g of Tris base, 27.59g of boric acid, 0.5 M Na2 EDTA,
20 mL in 1 L of ddH2O (pH 8.0).
Direct Molecular Diagnosis of MEN1 231
Table 1
MEN1 Primer Pairs for Mutation Analysis
Primer Primer sequence Product size Annealing Number    PCR
Exon name 5v to 3v (bp) temperature (°C) of cycles     conditionsa
2 men1 2F GGT GGA ACC TTA GCG GAC 592 55 30 Betaine
men1 2R GGT TTT GAA GAA GTG GGT C
3 men1 3F CCC ATG TTA AAG CAC AGA G 299 60 30 Standard
men1 3R ACA GTA TGA AGG GGA CAA G
4 men1 4F TGT CAT TCC CTG AAG CAG 252 60 30 Standard
men1 4R CCC ACA GCA AGT CAA GTC
5 and 6 men1 5 and 6F AAG GAC CCG TTC TCC TCC C 320 60 30 Standard
men1 5 and 6R CCT GCC TCA GCC ACT GTT AG
7 men1 7F GCA TTT GTG CCA GCA GGG 268 60 30 Standard
men1 7R GGG TGG TTG GAA ACT GAT GGA G
8 men1 8F GCT ACC CCC GAT GGT GAG AC 261 55 30 Betaine
men1 8R ATG GCC CTG TGG AAG GGA G
9 men1 9F CTG CTA AGG GGT GAG TAA GAG AC 260 50 30 Betaine
men1 9R ACC ACC TGT AGT GCC CAG AC
10 men1 10FB TGC CGA TGG GAC TGA GAC 714 55 or 60 30 Betaine
men1 10RB CTA GGG TTT GGG TAG AGG TG
aStandard and Betaine conditions are described in Subheading 3.1. The Betaine conditions enhance amplification of GC-rich sequences.
231
232 Petty, Glynn, and Bale
2.4. Sequencing Reagents
Sequencing reagents will vary because they will be specific to the
laboratory’s available sequencing equipment. Several different commercial kits
are available for manual sequencing of PCR products. In our hands, incorpo-rating PCR cycle sequencing with  33P-radiolabeled ddNTPs provides quick,
strong, optimally readable manual sequencing results as visualized after over-night autoradiography compared to methods using end-labeled primers with
T4 kinase, but lacks the advantage of being able to use one radiolabeled primer
mix for a number of sample reactions over a few weeks. We routinely use the
Thermosequenase kit (US 79750; Amersham Life Science Reagents), which
we have found provides reproducible high-quality sequence. A 6% polyacryla-mide sequencing gel (a commercial preparation mix for long-range sequencing
polyacrylamide gel mix) has proven optimal for manual sequencing. For auto-mated sequencing protocols, obtain reagents and methods for optimal analysis
recommended by the manufacturer of the equipment (see Note 4).
2.5. Equipment
1. Thermocycler.
2. Agarose gel electrophoresis equipment (power supply and gel boxes).
3. Manual sequencing gel-running apparatus.
4. Sequencing gel-running apparatus.
5. Glass plates, spacers, combs.
6. Power supply to support electrophoresis.
7. Gel dryer.
8. Pipets with accurate pipetting between 1 and 200 µL.
9. Centrifuge.
10. Or automated sequencer with associated gel and analysis components.
3. Methods
Because the equipment and technical expertise available within different
molecular diagnostic and research laboratories vary considerably, detailed
methods for DNA extraction and sequencing are not provided here. Several
different reliable protocols have been previously published in manuals on
molecular genetic techniques as well as in other chapters in the  Methods in
Molecular Medicine series (53 ,54) . Several different highly reliable commer-cial kits are available for both DNA extraction and sequencing.
After PCR-quality DNA has been prepared from the biological samples of
interest, which are generally leukocytes (although any nucleated specimen con-taining nondegraded DNA such as cultured amniocytes, chorionic villus
samples, buccal cells, fibroblasts, or fresh or archival tumor tissue may be used
as appropriate), PCR amplification of coding exons and flanking sequences
using the primers and conditions discussed next can be conducted.
Direct Molecular Diagnosis of MEN1 233
The methods outlined will specifically focus on providing detailed informa-tion about useful primer pairs and optimal PCR conditions to provide appropri-ate template for MEN1 analysis and general discussion of the analysis of point
mutations in the MEN1 gene by direct sequencing.
3.1. PCR of MEN1 Coding Regions and Splice Sites
A 50-µL (this volume may be decreased by adjusting all components
appropriately, but a 50-µL sample provides an optimal amount of specimen for
complete analysis and reanalysis if necessary) PCR is set up for each pair of
primers using one of the two following conditions, as noted in Table 1. LA 16,
a 15 1 mix of KlenTaq1 Pfu polymerase, may be mixed ahead of time and
stored at –20°C.
1. Standard conditions:
a. Template DNA (0.1–1 µg): 1 µL.
b. H2O: 34.5 µL.
c. 10X buffer: 5 µL.
d. dNTP (2 mM ): 5 µL.
e. Spermidine (50 mM ): 0.25 µL.
f. Primer 1(10 µM ): 1 µL.
g. Primer 2 (10 µM ): 1 µL.
h. Taq polymerase (5 U/µL): 0.25 µL.
i. MgCl2 (25 mM ): 3 µL.
2. Betaine conditions (to enhance amplification of GC-rich templates:
a. Template DNA (0.1–1 µg): 1 µL.
b. H2O: 14.3 µL.
c. 10X KlenTaq buffer: 5 µL.
d. dNTP (2 mM ): 5 µL.
e. DMSO: 2.5 µL.
f. Primer 1 (10 µM ): 1 µL.
g. Primer 2 (10 µM ): 1 µL.
h. LA 16: 0.2 µL.
i. 5 M Betaine: 20 µL.
3.2. General PCR Cycling Conditions
Annealing temperatures and cycling conditions may vary depending on the
specific thermocycler used. The following conditions were optimized using an
Ericomp dual-block, water-cooled thermocycler.
1. Initial denaturation step: 97°C for 2 min.
2. Cycling steps: 1: 96°C at 30 s; 2: For annealing temperatures of exon flanking
primers, see Table 1. 30 s; 3: 72°C at 40 s (total cycles: 30).
3. Final extension: 72°C for 5 min.
234 Petty, Glynn, and Bale
3.3. Initial Analysis of PCR Products
Prior to direct mutation analysis by sequencing, analyze 5 µL of each PCR
product in a 4% agarose gel to ensure that all PCR reactions generated specific
products (a single band) of the appropriate size and concentration. If a nonspe-cific PCR product is obtained, adjustment of the PCR conditions to increase
specificity will be necessary. Often, the simplest way to increase the specific-ity is to increase the annealing temperature by a few degrees or by employing a
“touchdown” method of annealing. Adjustments in the PCR reaction compo-nents, such as the buffer salt or magnesium concentration, can be employed to
enhance specificity if desired. Approximation of the size and concentration of
the PCR products can be easily done by comparing the migration distance of
the band and the band intensity of the PCR product to a well-quantified, appro-priately loaded, commercially available size standard marker designed to pro-duce bands in the 100- to 1000-bp size range. Ideally, the PCR reaction should
yield approx 10–30 ng/µL of a specific amplified product.
3.4. Sequence Analysis
The PCR sample generally must be purified prior to sequencing. This can be
done through commercially available spin columns designed for purifying PCR
products (Qiagen or Promega) or simply by treating 5 µL of the PCR product
with 1 µL of exonuclease I (10 U/µL) and 2 µL of shrimp alkaline phosphatase
(1 U/µL) for 15 min at 37°C. Stop the enzyme activity by incubating the sample
at 80°C for 15 min. Use approx 10–20 ng per each 100 bp of product for
sequencing. Follow the specific protocol for manual or automated sequencing
in your laboratory. We routinely use the forward primer fused in the initial
PCR reaction for sequencing.
3.5. Analysis of Results
Sequence chromatographs and or sequencing gel autoradiographs should be
read very carefully to look for any base changes. Always read the sequence
against a normal control sequence for comparison of peak height in chromato-graphs. For optimal analysis of manual sequencing autoradiographs, it is often
helpful to run multiple samples against a normal control in which samples are
grouped by the ddNTP. For instance, all “A” samples should be run next to one
another, all “G” samples next to one another, and so on so that mutations are
easy to identify visually. Any missense mutations should be checked against a
database or published literature describing previously identified polymor-phisms and mutations (3–7 ,24 ,25) .
If an individual who has features of MEN1 is found to have a previously
undescribed missense mutation that is not a known polymorphism, other fam-
Direct Molecular Diagnosis of MEN1 235
ily members, including parents, should be evaluated to determine whether it is
seen in affected or unaffected individuals. If the missense mutation is not seen
in unaffected parents of an affected individual, it is quite likely that it may be a
new disease-causing mutation. Any mutations that have been reported previ-ously as being associated with the disease are presumed to be disease-causing
mutations. Mutations that alter splice sites or cause frameshifts and nonsense
mutations are presumed to be disease causing mutations. Missense mutations
that affect conserved amino acid residues or make a significant change in the
polarity of the residue may be associated with disease in an affected individual
especially if they are not found in unaffected parents. Missense mutations that
cannot be identified as clear polymorphisms based on segregation with unaf-fected individuals in the pedigree or polymorphism database comparisons, or
that cannot be presumed to be disease causing based on the aforementioned
criteria, are not diagnostic. Once a mutation is found, confirm the mutation by
restriction endonuclease digestion (if the mutation changes a restriction site) or
reverse sequencing using the reverse primer. If a restriction site confirms the
mutation, a simple restriction digest can be used for direct mutation DNA
analysis in other family members (see Note 5).
3.6. Linkage-Based Diagnosis
When multiple family members are available and the clinical affection sta-tus can be clearly documented, linkage analysis may still be useful especially
in cases in which no mutation is found by sequencing analysis. Flanking and
intragenic primer sequences are readily available through the Genome Data-base. We routinely use primers from the following loci:  PYGM, DS11S913,
D11S970, D11S971, and D11S987. Although methods for diagnostic linkage
studies are beyond the scope of this chapter, incorporating 0.2  µL of  32P
_-dCTP in the PCR reaction and running samples on a 5% polyacrylamide gel
allow for clear visualization of genotypes after autoradiography. The methods
for interpretation of linkage analysis are also beyond the scope of this chapter,
but note that this type of testing is dependent on many variables that must be
considered in the computer analysis of linkage, including the penetrance of the
disorder for the ages of the individuals in the analysis and the prior probability
that this family’s disease is caused by a mutation in the MEN1 gene. Inherent
potential problems in linkage analysis include problems or errors in typing
owing to nonpaternity, formation of new alleles, misclassification or diagnosis
of affected individuals who may have a sporadic phenocopy of the disease, and
misclassification of currently nonexpressing gene carriers as unaffected.
Despite these problems, linkage remains quite useful for individuals in large
kindreds with MEN1 in whom no specific mutation can be identified.
236 Petty, Glynn, and Bale
3.7. Benefits and Pitfalls of Diagnosis
The medical benefit of testing for MEN1 molecular genetic susceptibility
for individuals at 50% risk in early diagnosis and treatment is sometimes
accompanied by the fear that this information may be misused by providing a
basis for discrimination against individuals who carry a predisposing muta-tion. However, given the availability of screening programs, medical manage-ment based on genetic predisposition, specific genetic counseling, provision of
accurate recurrence risks, patient education, and referral to appropriate support
resources for patients and their at-risk relatives, there are clear benefits for
identifying individuals at highest risk for MEN1. There are no clear guidelines
regarding at what age MEN1 mutation testing is optimal. Because most young
children do not develop symptoms of MEN1, it could be argued that testing
could wait until an individual is old enough to make an informed, educated
choice about having the test. Most medical screening programs for signs and
symptoms of MEN1 do not begin until late adolescence or early adulthood.
However, because some tumors can occur earlier, there may be benefits in
testing children, but there are not yet well-defined guidelines for MEN1 screen-ing in young pediatric populations.
3.8. Importance of Genetic Counseling
It is critically important to remember that genetic testing encompasses more
than a simple laboratory analysis. It needs to include pretesting counseling and
education; provision of informed consent; accurate interpretation of the test
results and their implications; follow-up conveyance of test results; and
posttesting education, management, and support. This is especially true when
DNA-based testing is used to determine more accurately a healthy individual’s
genetic risk as in predictive testing of an asymptomatic individual who, by
virtue of his or her family history, is at risk of having inherited MEN1 and
seeks to learn whether or not he or she has indeed inherited a MEN1 mutation.
4. Notes
1. Of the 166 mutations in MEN1 entered in the Human Gene Mutation Database at
the Institute of Medical Genetics in Cardiff, Wales (http://www.uwcm.ac.uk/
uwcm/mg/hgmd0.html), 43% are single-base substitutions, either nonsense or
missense. Small deletions, generally of 1–5 bp, occur in 34%, and small inser-tions (up to 15 bp) occur in 13%. Nucleotide substitutions that affect splicing
occur in 6%. Large deletions, insertions, or duplications have been reported in
<1% of cases reported to date, but this may be because the type of mutational
analysis used is biased against finding large deletions and rearrangements. Thus,
at present it seems that the vast majority of mutations are substitutions of a single
nucleotide or small deletions or insertions of one or more base pairs. These muta-
Direct Molecular Diagnosis of MEN1 237
tions have been found throughout the coding exons of the gene, and there is no
one hot spot that accounts for the majority of mutations in nonfounder popula-tions. To date, exon 2, the first coding exon and the second largest of the coding
exons (exon 10 is the largest), has the highest number of mutations reported, but
only 13% of the reported mutations fall in this exon, further highlighting the fact
that mutations are scattered throughout the sequence.
2. A variety of strategies to find base pair substitutions and minute insertions
and deletions to identify  MEN1 mutations has been described in the literature.
Standard mutational analyses including SSCP analyses and heteroduplex analy-sis, bi-dideoxy fingerprinting, and direct manual and automated sequencing have
been employed in our research laboratories with good results. These methods are
well described in protocol manuals as well as in the primary scientific literature
describing MEN1 mutations (4 ,5 ,17 ,36–52) . A combination of SSCP/heterodu-plex analysis identifies approx 80–85% of MEN1 mutations. Given the advances
in technology and the increased sensitivity of direct sequencing, the use of direct
sequencing, whether manual or automated, has been favored in our laboratories
for both analysis of research and diagnostic samples. Thus, we propose here that
direct sequencing of coding regions of the gene and intron/exon boundaries is the
analysis of choice for relatively small genes such as MEN1 in which mutations
are scattered throughout the coding region. However, if rapid high-throughput
sequencing is not available in the laboratory, a modified SSCP method can be
used with the primers cited in  Table 1 and analyzed using autoradiography if
0.2  µL 32P _-dCTP is added to the PCR reaction. Advances in microarray and micro-chip hybridization technology for analyzing sequences may become an optimal way
for screening sequences at a much more rapid pace than current automated methods.
3. Several other primer pairs have been reported for use in MEN1 mutation analysis
(6 ,17 ,29) . The primers reported here were specifically designed at the Yale Uni-versity DNA Diagnostic Laboratory to allow running PCR reactions in two gen-eral PCR buffers with the majority of samples having the same annealing
temperature. However, we have also used the other reported primers with suc-cess, including those listed through the National Institutes of Health Internet
database (http://www.nhgri.nih.gov/DIR/LGT/MEN1/table1.html).
4. Because sequencing one strand of a PCR product is usually sufficient to detect
mutations, we do not routinely use bidirectional sequencing because it has not
been proven to be cost-effective in most cases. Our diagnostic sequencing is done
using an ABI-373XL automated sequencer with which we routinely are able to
read through the 700 bp of exon 10 and flanking splice sites. When using manual
sequencing, bidirectional sequencing using both the forward and reverse primers
of the two largest exons, exons 2 and 10, is necessary. By running samples on 5%
long-range sequencing polyacrylamide gels at 55 W in which the sample runs for
both 1.5 and 4.0 h by doing a second loading on the same gel, we are routinely
able to read approx 400 bp of DNA, spanning each exon completely.
5. A useful Web site is Online Mendelian Inheritance in Man (OMIM) (http://
www3.ncbi.nlm.nih.gov/omim/searchomim.html), a regularly updated database
238 Petty, Glynn, and Bale
of inherited disorders with information on clinical presentation, molecular basis,
cytogenetics, mapping, and population genetics. OMIM also has direct links to
the National Library of Medicine’s MEDLINE database, DNA sequence data-bases, and the Genome Database. As stated in Note 1, the Human Gene Mutation
Database at the Institute of Medical Genetics in Cardiff is a useful catalog of
known mutations in MEN1. Published reports of known mutations and polymor-phisms are also available (3–7 ,24 ,25) . The GeneTests™, a searchable database
(http://www.genetests.org/), provides contact and test information regarding
clinical and research laboratories worldwide that are doing molecular testing for
MEN1. It is a password-protected (passwords are issued free of charge to health
care professionals who register) database in which laboratories offering either
clinical or research-based testing for any genetic disease, including MEN1, can
list their services and contact information, making their services readily avail-able to health care professionals. Contact and discussion with other diagnostic
laboratories doing MEN1 molecular testing may help provide additional useful
new information for those individuals setting up  MEN1 testing in their own
molecular genetic laboratories. Currently, five laboratories worldwide (includ-ing the Yale University DNA Diagnostic Laboratory) are listed as offering clini-cal testing, and three additional laboratories are listed as providing research-based
testing.
References
1. Miller, J. A. and Norton, J. A. (1997) Multiple endocrine neoplasia. Cancer Treat.
Res. 90, 213–225.
2. Thakker, R. V. (1998) Multiple endocrine neoplasia—syndromes of the twentieth
century. J. Clin. Endocrinol. Metab. 83, 2617–2620.
3. Bassett, J. H., Forbes, S. A., Pannett, A. A., et al. (1998) Characterization of
mutations in patients with multiple endocrine neoplasia type 1. Am. J. Hum. Genet.
62, 232–244.
4. Agarwal, S. K., Kester, M. B., Debelenko, L. V., et al. (1997) Germline mutations
of the MEN1 gene in familial multiple endocrine neoplasia type 1 and related
states. Hum. Mol. Genet. 6, 1169–1175.
5. Giraud, S., Zhang, C. X., Serova-Sinilnikova, O., et al. (1998) Germ-line muta-tion analysis in patients with multiple endocrine neoplasia type 1 and related dis-orders. Am. J. Hum. Genet. 63, 455–467.
6. Sato, M., Matsubara, S., Miyauchi, A., Ohye, H., Imachi, H., Murao, K., and
Takahara, J. (1998) Identification of five novel germline mutations of the MEN1
gene in Japanese multiple endocrine neoplasia type 1 (MEN1) families. J. Med.
Genet. 35, 915–919.
7. Teh, B. T., Kytola, S., Farnebo, F., et al. (1998) Mutation analysis of the MEN1
gene in multiple endocrine neoplasia type 1, familial acromegaly and familial
isolated hyperparathyroidism. J. Clin. Endocrinol. Metab. 83, 2621–2626.
Direct Molecular Diagnosis of MEN1 239
8. Yu, F., Venzon, D. J., Serrano, J., Goebel, S. U., Doppman, J. L., Gibril, F., and
Jensen, R.T. (1999) Prospective study of the clinical course, prognostic factors,
causes of death, and survival in patients with long-standing Zollinger-Ellison syn-drome. J. Clin. Oncol. 17, 615–630.
9. Bystrom, C., Larsson, C., Blomberg, C., Sandelin, K., Falkmer, U., Skogseid, B.,
Oberg, K., Werner, S., and Nordenskjold, M. (1990) Localization of the MEN1
gene to a small region within chromosome 11q13 by deletion mapping in tumors.
Proc. Natl. Acad. Sci. USA 87, 1968–1972.
10. Bale, S. J., Bale, A. E., Stewart, K., Dachowski, L., McBride, O. W., Glaser, T.,
Green, J. E. III, Mulvihill, J. J., Brandi, M. L., and Sakaguchi, K. (1989) Linkage
analysis of multiple endocrine neoplasia type 1 with INT2 and other markers on
chromosome 11. Genomics 4, 320–322 (published erratum appears in Genomics
1989;5[1], 166).
11. Courseaux, A., Grosgeorge, J., Gaudray, P., et al. (1996) Definition of the mini-mal MEN1 candidate area based on a 5-Mb integrated map of proximal 11q13.
The European Consortium on Men1 (GENEM 1; Groupe d’Etude des Neoplasies
Endocriniennes Multiples de type 1). Genomics 37, 354–365.
12. Guru, S. C., Agarwal, S. K., Manickam, P., et al. (1997) A transcript map for the
2.8-Mb region containing the multiple endocrine neoplasia type 1 locus. Genome
Res. 7, 725–735.
13. Larsson, C., Skogseid, B., Oberg, K., Nakamura, Y., and Nordenskjold, M. (1988)
Multiple endocrine neoplasia type 1 gene maps to chromosome 11 and is lost in
insulinoma. Nature 332, 85–87.
14. Lemmens, I., Merregaert, J., Van de Ven, W. J., et al. (1997) Construction of a
1.2-Mb sequence-ready contig of chromosome 11q13 encompassing the multiple
endocrine neoplasia type 1 (MEN1) gene. The European Consortium on MEN1.
Genomics 44, 94–100.
15. Nakamura, Y., Larsson, C., Julier, C., Bystrom, C., Skogseid, B., Wells, S., Oberg,
K., Carlson, M., Taggart, T., and O’Connell, P. (1989) Localization of the genetic
defect in multiple endocrine neoplasia type 1 within a small region of chromo-some 11. Am. J. Hum. Genet. 44, 751–755.
16. Knudson, A. G. (1971) Mutation and cancer: statistical study of retinoblastoma.
Proc. Natl. Acad. Sci. USA 68, 820–823.
17. Chandrasekharappa, S. C., Guru, S. C., Manickam, P., et al. (1997) Positional
cloning of the gene for multiple endocrine neoplasia-type 1.  Science 276,
404–407.
18. Guru, S. C., Goldsmith, P. K., Burns, A. L., Marx, S. J., Spiegel, A. M., Collins,
F. S., and Chandrasekharappa, S. C. (1998) Menin, the product of the MEN1 gene,
is a nuclear protein. Proc. Natl. Acad. Sci. USA 95, 1630–1634.
19. Agarwal, S. K., Guru, S. C., Heppner, C., et al. (1999) Menin interacts with the
AP1 transcription factor JunD and represses JunD-activated transcription.  Cell
96, 143–152.
240 Petty, Glynn, and Bale
20. Bear, J. C., Briones-Urbina, R., Fahey, J. F., and Farid, N. R. (1985) Variant mul-tiple endocrine neoplasia I (MEN IBurin): further studies and non-linkage to HLA.
Hum. Hered. 35, 15–20.
21. Farid, N. R., Buehler, S., Russell, N. A., Maroun, F. B., Allerdice, P., and Smyth,
H. S. (1980) Prolactinomas in familial multiple endocrine neoplasia syndrome
type I: relationship to HLA and carcinoid tumors. Amer. J. Med. 69, 874–880.
22. Petty, E. M., Green, J. S., Marx, S. J., Taggart, R. T., Farid, N., and Bale, A. E.
(1994) Mapping the gene for hereditary hyperparathyroidism and prolactinoma
(MEN1Burin) to chromosome 11q: evidence for a founder effect in patients from
Newfoundland. Am. J. Hum. Genet. 54, 1060–1066.
23. Olufemi, S. E., Green, J. S., Manickam, P., et al. (1998) Common ancestral muta-tion in the MEN1 gene is likely responsible for the prolactinoma variant of MEN1
(MEN1Burin) in four kindreds from Newfoundland. Hum. Mutat. 11, 264–269.
24. Krawczak, M. and Cooper, D. N. (1997) The Human Gene Mutation Database.
Trends Genet. 13, 121–122.
25. Lemmens, I., Van de Ven, W. J., Kas, K., et al. (1997) Identification of the mul-tiple endocrine neoplasia type 1 (MEN1) gene. The European Consortium on
MEN1. Hum. Mol. Genet. 6, 1177–1183.
26. Carling, T., Correa, P., Hessman, O., Hedberg, J., Skogseid, B., Lindberg, D.,
Rastad, J., Westin, G., and Akerstrom, G. (1998) Parathyroid MEN1 gene muta-tions in relation to clinical characteristics of nonfamilial primary hyperparathy-roidism. J. Clin. Endocrinol. Metab. 83, 2960–2963.
27. Debelenko, L. V., Brambilla, E., Agarwal, S. K., et al. (1997) Identification of
MEN1 gene mutations in sporadic carcinoid tumors of the lung. Hum. Mol. Genet.
6, 2285–2290.
28. Farnebo, F., Teh, B. T., Kytola, S., Svensson, A., Phelan, C., Sandelin, K.,
Thompson, N. W., Hoog, A., Weber, G., Farnebo, L.O., and Larsson, C. (1998)
Alterations of the MEN1 gene in sporadic parathyroid tumors. J. Clin. Endocrinol.
Metab. 83, 2627–2630.
29. Görtz, B., Roth, J., Speel, E. J. M., Krähenmann, A., DeKrijger, R. R., Matias-Guiu, X., Muletta-Feurer, S., Rütmann, K., Saremaslani, P., Heitz, P. U., and
Komminoth, P. (1999) MEN1 gene mutation analysis of sporadic adrenocortical
lesions. Int. J. Cancer 80, 373–379.
30. Shou, W., Seol, J. H., Shevchenko, A., Baskerville, C., Moazed, D., Chen, Z. W. S.,
Jang, J., Shevchenko, A., Charbonneau, H., and Deshaies, R. J. (1999) Exit from
mitosis is triggered by Tem1-dependent release of the protein phosphatase Cdc14
from nucleolar RENT complex. Cell 97, 233–244.
31. Prezant, T. R., Levine, J., and Melmed, S. (1998) Molecular characterization of
the men1 tumor suppressor gene in sporadic pituitary tumors. J. Clin. Endocrinol.
Metab. 83, 1388–1391.
32. Sawicki, M. P., Wan, Y. J., Johnson, C. L., Berenson, J., Gatti, R., and Passaro,
E. Jr. (1992) Loss of heterozygosity on chromosome 11 in sporadic gastrinomas.
Hum. Genet. 89, 445–449.
Direct Molecular Diagnosis of MEN1 241
33. Vortmeyer, A. O., Boni, R., Pak, E., Pack, S., and Zhuang, Z. (1998) Multiple
endocrine neoplasia 1 gene alterations in MEN1-associated and sporadic lipomas.
J. Natl. Cancer Inst. 90, 398–399.
34. Teh, B. T., Esapa, C. T., Houlston, R., Grandell, U., Farnebo, F., Nordenskjold,
M., Pearce, C. J., Carmichael, D., Larsson, C., and Harris, P. E. (1998) A family
with isolated hyperparathyroidism segregating a missense MEN1 mutation and
showing loss of the wild-type alleles in the parathyroid tumors. Am. J. Hum. Genet.
63, 1544–1549.
35. Zhuang, Z., Ezzat, S. Z., Vortmeyer, A. O., et al. (1997) Mutations of the MEN1
tumor suppressor gene in pituitary tumors. Cancer Res. 57, 5446–5451.
36. Korf, B. R. and Pagon, R. A. (1998) Overview of molecular genetic diagnosis
(Unit 9.1), in Current Protocols in Human Genetics (Dracopoli, N. C., Haines,
J. L., Korf, B. R., et al., eds.), John Wiley & Sons, New York, pp. 9.1.1–9.1.9.
37. Smooker, P. M. and Cotton, R. G. (1993) The use of chemical reagents in the
detection of DNA mutations. Mutat. Res. 288, 65–77.
38. Bottema, C. D. and Sommer, S. S. (1993) PCR amplification of specific alleles:
rapid detection of known mutations and polymorphisms. Mutat. Res. 288, 93–102.
39. Hayashi, K. (1991) PCR-SSCP: a simple and sensitive method for detection of
mutations in the genomic DNA. PCR Methods Appl. 1, 34–38.
40. Loda, M. (1994) Polymerase chain reaction-based methods for the detection of
mutations in oncogenes and tumor suppressor genes. Human Pathol. 25, 564–571.
41. Sidransky, D. (1997) Nucleic acid-based methods for the detection of cancer.
Science 278, 1054–1058.
42. Liu, Q., Feng, J., and Sommer, S. S. (1996) Bi-directional dideoxy fingerprinting
(Bi-ddF): a rapid method for quantitative detection of mutations in genomic
regions of 300–600 bp. Hum. Mol. Genet. 5, 107–114.
43. Ben-Ezra, J. (1995) Amplification methods in the molecular diagnosis of genetic
diseases. Clin. Lab. Med. 15, 795–815.
44. McPherson, R. (1995) Molecular basis of genetic disease and molecular methods.
Clin. Lab. Med. 15, 779–794.
45. Wagener, C., Epplen, J. T., Erlich, H., Peretz, H., and Vihko, P. (1994) Molecular
biology techniques in the diagnosis of monogenic diseases. Clin. Chim. Acta 225,
S35–S50.
46. Grompe, M. (1993) The rapid detection of unknown mutations in nucleic acids.
Nat. Genet. 5, 111–117.
47. Blaszyk, H., Hartmann, A., Schroeder, J. J., McGovern, R. M., Sommer, S. S.,
and Kovach, J. S. (1995) Rapid and efficient screening for p53 gene mutations by
dideoxy fingerprinting (ddF). BioTech 18, 256–260.
48. Warren, W., Hovig, E., Smith-Sorensen, B., Borresen, A. L., Fujimura, F. K., Liu,
Q., Feng, J., and Sommer, S. S. (1997) Detection of mutations by single-strand
conformation polymorphism (SSCP) analysis and SSCP-hybrid methods (Unit
7.4), in Current Protocols in Human Genetics (Dracopoli, N. C., Haines, J. L.,
Korf, B. R., et al., eds.), John Wiley & Sons, New York, pp. 7.4.1–7.4.23.
242 Petty, Glynn, and Bale
49. Mashal, R. D., Koontz, J., and Sklar, J. (1995) Detection of mutations by cleavage
of DNA heteroduplexes with bacteriophage resolvases. Nat. Genet. 9, 177–183.
50. Sekiya, T. (1993) Detection of mutant sequences by single-strand conformation
polymorphism analysis. Mutat. Res. 288, 79–83.
51. Dracopoli, N., ed. (1994) Current Protocols in Human Genetics, John Wiley &
Sons, New York.
52. Cotton, R.G. (1993) Current methods of mutation detection.  Mutat. Res. 285,
125–144.
53. Thierfelder, L. (1998) Mutation detection by cycle sequencing (Unit 7.7), in Cur-rent Protocols in Human Genetics (Dracopoli, N. C., Haines, J. L., Korf, B. R., et
al., eds.), John Wiley & Sons, New York, pp. 7.7.1–7.7.6.
54. Shuldiner, A. R., LeRoith, D., and Roberts, C. T. (1995) DNA sequence analysis,
in Molecular Endocrinology: Basic Concepts and Clinical Correlations
(Weintraub, B. D., ed.), Raven Press, New York, pp. 13–21.
Molecular Detection of MEN2 243
19
243
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
Molecular Detection of Multiple
Endocrine Neoplasia Type 2
Edward H. Rowsell and Myra J. Wick
1. Introduction
1.1. Overview
Multiple endocrine neoplasia type 2 (MEN2) comprises three autosomal
dominant disorders: MEN2A, familial medullary thyroid carcinoma (FMTC),
and MEN2B. Clinical features common to both MEN2A and MEN2B include
C-cell hyperplasia, medullary thyroid cancer (MTC), and pheochromocytoma.
Other features of MEN2B are ocular and mucosal neuromas, gastrointestinal
ganglioneuromatosis, and skeletal abnormalities. FMTC is characterized by
MTC in the absence of parathyroid or adrenal disease.
Germline mutations within the RET proto-oncogene are definitively associ-ated with MEN2A, FMTC, and MEN2B. RET encodes a member of the
tyrosine kinase receptor family. The RET protein contains an extracellular
cadherin-like domain, a cysteine-rich domain, a transmembrane domain, and
two intracellular tyrosine kinase domains (Fig. 1). The RET-encoded protein
is activated by interaction with the glial cell line-derived neurotropic factor
GDFN/GDFN receptor-_ complex (1) . This interaction results in RET dimer-ization, autophosphorylation, and phosphorylation of intracellular substrates.
Approximately 95% of MEN2A and 85% of FMTC patients have a germline
point mutation within one of five cysteine residues in exon 10 (codons 609,
611, 618, and 620) or exon 11 (codon 634) of the RET gene. Rare noncysteine
mutations have been identified in exons 11 (codon 631), 13 (codon 768), and
14 (codons 804 and 844) of patients with FMTC/MEN2A; each of these muta-tions accounts for <1% of MEN2A/FMTC families  (2) . Recently, two addi-
244 Rowsell and Wick
tional germline mutations within codons 790 and 791 (exon 13) were identi-fied in 1.6 and 1.1%, respectively, of German MEN2A/FMTC families (2). In
contrast to the variety of mutations associated with MEN2A/FMTC, a single
point mutation within exon 16 (codon 918, ATGAACG) is detected in 95% of
MEN2B patients. Current data indicate that these MEN2/FMTC-associated
RET mutations are dominant gain-of-function mutations, resulting in increased
kinase activity of the mutant RET-encoded protein (3) .
The MEN literature includes numerous articles with guidelines for evalua-tion and testing of MEN2/FMTC family members as well as for patients pre-senting with apparently sporadic cases of MTC (4–6) . The laboratory director
should be aware of these guidelines to assist physicians who might be unfamil-Fig. 1. The RET proto-oncogene–encoded tyrosine kinase. The protein contains
extracellular cadherin-like and cysteine-rich domains, a transmembrane domain, and
intracellular tyrosine kinase domains. ATP, adenosine triphosphate binding site.
Molecular Detection of MEN2 245
iar with the evaluation process. Therefore, we briefly summarize these guide-lines here. For MEN2A/FMTC families, mutational analysis should initially
be performed on an affected family member. Once the mutation has been iden-tified, testing should be offered to at-risk family members. Molecular-based
testing should be done at an early age, because thyroidectomy by age 6 is indi-cated in individuals carrying mutations  (7) . It is also recommended that all
patients with apparent sporadic MTC be screened for MEN2A/FMTC muta-tions, because approx 24% of such cases will demonstrate a germline RET
mutation (8) . Phenotypic features of MEN2B usually enable the clinician to
make an accurate diagnosis in the absence of molecular testing. However,
molecular-based testing is useful for individuals who lack some of the pheno-typic features of MEN2B, or in very young children who are presymptomatic
(3) . Also note that although de novo MEN2A/FMTC-associated mutations are
rare, as many as 50% of MEN2B cases are the result of  de novo germline
mutations in codon 918 (9) .
1.2. Choice of Methods
A variety of methods have been developed for the molecular diagnosis of
MEN2A/FMTC in the clinical molecular genetics laboratory, including
sequencing, denaturing gradient gel electrophoresis (10) , single-strand confor-mation polymorphism (SSCP) analysis  (11 ,12) , heteroduplex (HET) forma-tion (13 ,14) , and restriction enzyme (RE) analysis  (14 ,15) . Included in this
chapter are protocols for manual sequencing of exons 10 and 11; SSCP analy-sis of exons 10 and 11; HET analysis for mutations in exons 10, 11, and 13; and
RE analysis for the codon 918 mutation associated with MEN2B.
Each of the methods for MEN2A/FMTC mutation detection has inherent
advantages and disadvantages. Sequencing allows for definitive identification
of the mutation, without the need for additional confirmatory testing. The
sequencing protocol described herein, however, is relatively time- and labor-intensive (compared to screening methods such as HET) and involves the use
of radioisotopes. Adaptation of this sequencing protocol to a semiautomated
system such as the Li-Cor Model 4200 or PE Applied Biosystems Model 377
system would increase efficiency and eliminate the use of radioisotopes.
Sequencing of exons 10 and 11 results in a detection rate of approx 95% of
MEN2A patients and 85% of FMTC families.
The SSCP protocol described herein has the advantage of utilizing a kit and
SSCP equipment, both of which can be purchased from Pharmacia Biotech,
Piscataway, NJ. The kit includes premade SSCP gels and the silver staining
reagents. This SSCP protocol was reported to detect 20 of 21 different muta-tions that represent approx 90% of known exon 10 and 11 mutations (12) .
246 Rowsell and Wick
The HET protocol  (14) utilizes a modified Mutation Detection Enhance-ment matrix enabling efficient screening for MEN mutations without the use
of radioisotopes. Although this method simultaneously detects at least 16 dif-ferent RET mutations, the specific mutation must be confirmed by RE analysis
or sequencing (in cases in which a mutation has previously been identified in a
family member, HET analysis alone is adequate). HET analysis of exons 10,
11, and 13 results in detection rates of 95% and slightly greater than 85% for
MEN2A and FMTC patients, respectively.
2. Materials
2.1. Samples
Peripheral blood is the sample type most frequently submitted for molecular
genetic analysis of RET mutations, although other specimen types containing
nucleated cells may also be used (e.g., buccal swabs, amniocytes, cord blood).
It is preferable that peripheral blood be collected in tubes containing EDTA or
acid citrate dextrose as the anticoagulant.
The Puregene® DNA Isolation Kit (nos. D-5500, D-5500A, and D-7000A;
Gentra Systems, Minneapolis, MN) allows rapid isolation of high-quality
genomic DNA from whole blood, cultured cells, and tissue, respectively. It is
based on published salting-out procedures, thus eliminating the use of toxic
organic solvents.
2.2. Reagents and Equipment for Sequencing (MEN2A)
1. Purified specimen DNA in water or 1X TE buffer (pH 8.0) at 250 µg/mL.
2. Gene Amplification Kit (no. N801-0043; PE Applied Biosystems, Foster City,
CA) containing 5 U/µL of AmpliTaq® Taq DNA polymerase; 10 mM dATP,
dCTP, dGTP, and dTTP; and GeneAmp® 10X PCR Buffer (500 mM KCl,
100 mM Tris-HCl, pH 8.3, 15 mM MgCl2, and 0.01% [w/v] gelatin, in a volume
of 1.5 mL).
3. 5X reaction buffer with dNTPs. This buffer is made in-house as follows: 100 µL
of dATP (10 mM), 100 µL of dCTP (10 mM), 100 µL of dGTP (10 mM), 100 µL
of dTTP (10 mM), 100 µL of sterile water, and 500 µL of GeneAmp 10X PCR
Buffer total volume of 1000 µL). The 5X buffer can be stored at –70°C for up to
6 mo. Thus, it is convenient to make batches of the 5X buffer and to store them as
1-mL aliquots.
4. Previously prepared primers stored (–70°C) at a concentration of 20 µM. Refer to
Table 1 for primer sequences.
5. High-Strength Analytical Grade Agarose (no. 162-0125; Bio-Rad, Hercules, CA).
6. NuSieve® GTG® Agarose (no. 50081; FMC BioProducts, Rockland, ME).
7. TAE buffer, prepared as 50X TAE (2 M Tris-acetate, 50 mM EDTA): 968.0 g of
Tris base, 228.4 g of glacial acetic acid, 400.0 mL of 0.5 M EDTA, pH 8.0. Dilute
to 4 L with sterile water. The 50X buffer is stable at room temperature for 3 mo.
Dilute the appropriate quantity of 50X buffer with sterile water to 1X before use.
Molecular Detection of MEN2 247
8. 6X Sample buffer for agarose gel electrophoresis: 0.125 g of bromophenol blue,
7.5 g of Ficoll® 400 (no. 17-0400-01; Pharmacia Biotech), in sufficient quantity to
50 mL in deionized, distilled water. This buffer is stable for 1 yr at room temperature.
9. Ethidium bromide (EtBr).
10. pGEM® DNA markers (no. G1741; Promega, Madison, WI).
11. Exonuclease I (10 U/mL) (no. E 70073Z; Amersham, Arlington Heights, IL).
12. Shrimp Alkaline Phosphatase (no. E 70092Y; Amersham).
13. ThermoSequenase™ cycle sequencing kit (no. US 78500; Amersham). The kit
includes 33P-radiolabeled terminators.
14. 40% Acrylamide/bis-acrylamide solution (19 1) (no. 161-0144; Bio-Rad).
Table 1
RET Oligonucleotide PCR Primers
Sequencing analysis
Locus-specific amplification: ~1000-bp amplicon (amplicon contains exons
10 and 11)
Forward (RET A2, 21mer): 5v-CAA CAT TTG CCC TCA GGA CTG-3v
Reverse (CRT 19A, 20mer): 5v-CTT GAA GGC ATC CAC GGA GA-3v
Sequencing
Exon 10 (RET D, 22mer): 5v-TTG GGA CCT CAG ATG TGC TGT T-3v
Exon 11 (CRT 19B, 19mer): 5v-GCA TAC GCA GCC TGT ACC C-3v
HET analysis
Exon 10: 232-bp amplicon
Forward (RET I, 19mer): 5v-CGC CCC AGG AGG CTG AGT G-3v
Reverse (RET I, 22mer): 5v-TTG GGA CCT CAG ATG TGC TGT T-3v
Exon 11: 150-bp amplicon
Forward (CRT 19B, 19mer): 5v-GCA TAC GCA GCC TGT ACC C-3v
Reverse (CRT 2C, 20mer): 5v-GAC AGC AGC ACC GAG ACG AT-3v
Exon 13: 296-bp amplicon
Forward (CRT 4F, 22mer): 5v-GCA GGC CTC TCT GTC TGA ACT T-3v
Reverse (CRT 4E, 20mer): 5v-GGA GAA CAG GGC TGT ATG GA-3v
SSCP analysis
Exon 10
Forward (21mer): 5v-GGG GCA GCA TTG TTG GGG GAC-3v
Reverse (19mer): 5v-CGT GGT GGT CCC GGC CGC C-3v
Exon 11
Forward (20mer): 5v-CCT CTG CGG TGC CAA GCC TC-3v
Reverse (21mer): 5v-GAA GAG GAC AGC GGC TGC GAT-3v
RFLP analysis
Exon 16: 195-bp amplicon
Forward (Ret 16F, 20mer): 5v-AGG GAT AGG GCC TGG GCT TC-3v
Reverse (Ret 16R, 20mer): 5v-TAA CCT CCA CCC CAA GAG AG-3v
248 Rowsell and Wick
15. TBE buffer (5X stock): 216.0 g of Tris base, 110.0 g of boric acid, 80.0 mL of
0.5 M EDTA, pH 8.0. Add deionized water to a final volume of 4 L. The 5X stock
is stable at room temperature for 1 yr. Dilute the appropriate quantity of 5X buffer
to 1 L with deionized water before use.
16. 6% Polyacrylamide/7  M urea gel mix (for sequencing gels): 150 mL of 40%
acrylamide/bis stock, 200 mL of 5X TBE stock, 420 g of urea. Add water to a
total volume of 1 L, and degas under vacuum when freshly made. Store refriger-ated in the dark for up to 6 mo.
17. Urea.
18. TEMED.
19. Ammonium persulfate (APS), prepared as a 10% (w/v) solution: 0.1 g of APS diluted
to a final volume of 1.0 mL. This may be stored in the refrigerator for 1 wk.
20. Sequencing gel-loading buffer: 45% formamide, 20 mM EDTA, 0.05% bro-mophenol blue, 0.05% xylene cyanol. Bring to a final volume of 10 mL with
deionized water.
21. Camera and Polaroid® Polapan® 667 film (Polaroid, Cambridge, MA).
22. Kodak X-Omat AR5 X-ray film (35  × 43 cm) (no. 165 1512; Eastman Kodak
Co., Rochester, NY).
23. Thermocycler and polymerase chain reaction (PCR) tubes (see Note 1).
24. Submarine gel electrophoresis system such as the Bio-Rad Wide Mini-Sub Cell
GT System (no. 170-4405).
25. Sequencing gel apparatus with required accessories (plates, spacers, sharkstooth
combs, clamps). The sequencing described in this protocol was performed with a
Gibco-BRL Model S2 Sequencing Gel Electrophoresis Apparatus (no. 21105-010; Life Technologies, Gaithersburg, MD).
26. Electrophoresis power supplies for submarine gel electrophoresis and sequenc-ing gel electrophoresis.
27. UV transilluminator.
28. Gel dryer, such as Bio-Rad Model 583 Gel Dryer (no. 165 1745).
2.3. Reagents and Equipment for HET Analysis
(MEN2A and FMTC)
1. Purified specimen DNA in water or 1X TE buffer (pH 8.0) at 250 µg/mL.
2. Taq DNA Polymerase in Storage Buffer A Kit (nos. M2861–M2868; Promega)
including Taq DNA polymerase (5 U/µL) and 10X Reaction Buffer A with
15 mM MgCl2 (500 mM KCl; 100 mM Tris-HCl, pH 9.0, at 25°C; 1.0% Triton
X-100; and 15 mM MgCl2).
3. Bulk dNTPs (no. U1330; Promega) containing 100 mM dATP, dCTP, dGTP, and
dTTP each in water at pH 7.5.
4. 5X Reaction Buffer A with dNTPs. This buffer is made in the lab as follows: 12.5 µL
of 100 mM bulk dNTPs, 487.5 µL of sterile water, and 500.0 µL of 10X Reaction
Buffer A (total volume of 1000 µL). The 5X buffer may be stored at  –70°C for
up to 6 mo. Thus, it is convenient to make batches of the 5X buffer and store
them as 1-mL aliquots.
Molecular Detection of MEN2 249
5. Working primers (10 µM ) prepared in water from 100 µM stock primer solutions
and stored at –70°C. Refer to Table 1 for primer sequences.
6. 0.5 M EDTA (pH 8.0).
7. MDE™ gel-loading buffer (50% [w/v] sucrose, 0.6% [w/v] bromophenol blue,
0.6% [w/v] xylene cyanol in 1X TE buffer, pH 8.0): Dissolve 25.0 g of sucrose,
0.3 g of bromophenol blue, 0.3 g of xylene cyanol, and 5 mL of 10X TE buffer
(pH 8.0) in about 30 mL of deionized water. Dilute with deionized water to a
50-mL final volume. This buffer is stable for 1 yr at room temperature.
8. pGEM DNA markers (no. G1741; Promega).
9. MDE™ gel solution (2X concentrate) (no. 50620; FMC BioProducts).
10. 10X TBE stock solution: 216.0 g of Tris base, 110.0 g of boric acid, 80.0 mL of
0.5 M EDTA (pH 8.0). Add deionized water to a final volume of 2.0 L. The 10X
stock is stable at room temperature for 1 yr.
11. Urea.
12. Formamide.
13. TEMED.
14. APS, prepared as a 10% (w/v) solution: 0.1 g of APS diluted to a final volume of
1.0 mL. Prepare fresh each time.
15. Ethidium bromide.
16. Camera and Polaroid Polapan 667 film.
17. Thermocycler and PCR reaction tubes.
18. Heating block.
19. Gel electrophoresis apparatus with accessories, including spacer sets, combs, and
glass plates. A power supply is also required.
20. UV transilluminator box.
2.4. Reagents and Equipment for SSCP Analysis
1. Purified specimen genomic DNA in water or 1X TE buffer at 10 µg/mL.
2. 5X Reaction Buffer N from PCR Optimization Kit (no. K1220-01; Invitrogen,
San Diego, CA).
3. GeneAmp® dNTP mix (no. N808-0007; PE Applied Biosystems) including
10 mM each of dATP, dCTP, dGTP, and dTTP.
4. AmpliTaq Taq DNA polymerase (5 U/µL) (no. N801-0060; PE Applied
Biosystems).
5. Working primers (10 µM) prepared in water from 100 µM stock primer solutions
and stored at –70°C. Refer to Table 1 for primer sequences.
6. Thermocycler and PCR reaction tubes.
7. Heating block.
8. PhastSystem™ gel electrophoresis system (no. 18-1018-23; Pharmacia Biotech),
consisting of the Separation and Control Unit (electrophoresis chambers, power
supply, microprocessor) and the Development Unit.
9. PhastGel™ sample applicators: either 6/4 (6 samples, 4  µL each) or 8/1
(8 samples, 1  µL each) (nos. 18-0012-29 and 18-1618-01, respectively;
Pharmacia Biotech).
250 Rowsell and Wick
10. PhastGel™ Homogeneous 20: native 20% polyacrylamide gel media (no.
17-0624-01; Pharmacia Biotech).
11. PhastGel™ native buffer strips (no. 17-0517-01; Pharmacia Biotech).
12. PhastGel™ Silver Kit gel-staining kit (no. 17-0617-01; Pharmacia Biotech).
2.5. Reagents and Equipment for MEN2B Analysis
(PCR and RE Digestion)
1. Purified specimen genomic DNA in water or 1X TE buffer at 10 µg/mL.
2. Gene Amplification Kit (no. N801-0043; PE Applied Biosystems) containing
5 U/µL Taq DNA polymerase; 10 mM dATP, dCTP, dGTP, and dTTP; and
GeneAmp 10X PCR Buffer (500 mM KCl, 100 mM Tris-HCl, pH 8.3, 15 mM
MgCl2 and 0.01% [w/v] gelatin, in a volume of 1.5 mL).
3. 5X Reaction buffer with dNTPs. This buffer is made in-house as follows: 100 µL
of dATP, 100 µL of dCTP, 100 µL of dGTP, 100 µL of dTTP, 100 µL of sterile
water, and 500 µL of GeneAmp 10X PCR Buffer (total volume of 1000 µL). The
5X buffer may be stored at –70°C for up to 6 mo. Thus, it is convenient to make
batches of the 5X buffer and store them as 1-mL aliquots.
4. Previously prepared primers stored (–70°C) at a concentration of 20 µM. Refer to
Table 1 for primer sequences.
5. FokI and NEB Buffer 4 (no. 109; New England Biolabs, Beverly, MA).
6. High-Strength Analytical Grade Agarose (no. 162-0125; Bio-Rad).
7. NuSieve GTG Agarose (no. 50081; FMC BioProducts).
8. TAE buffer, prepared as 50X TAE (2 M Tris acetate, 50 mM EDTA): 968.0 g of
Tris base, 228.4 g of glacial acetic acid, 400.0 mL of 0.5 M EDTA, pH 8.0. Dilute
to 4 L with sterile water. The 50X buffer is stable at room temperature for 3 mo.
Dilute the appropriate quantity of 50X buffer with sterile water to 1X before use.
9. 6X Sample buffer for agarose gel electrophoresis: 0.125 g of bromophenol blue,
7.5 g of Ficoll 400 (no. 17-0400-01; Pharmacia Biotech), in sufficient quantity to
50 mL in deionized, distilled water. This buffer is stable for 1 yr at room temperature.
10. EtBr.
11. Camera and Polaroid Polapan 667 film.
12. Thermocycler and PCR reaction tubes (see Note 1).
13. Submarine gel electrophoresis system such as Bio-Rad Wide Mini-Sub Cell GT
System (no. 170-4405).
14. Electrophoresis power supplies for submarine gel electrophoresis.
15. UV transilluminator.
3. Methods
3.1. Sequencing of RET Exons 10 and 11
3.1.1. PCR Amplification of Exons 10 and 11
1. Label PCR reaction tubes for patient specimens, positive controls, and the
no-DNA control.
2. Prepare the PCR reaction master mix. It is advisable to prepare a slight excess of
master mix to allow for pipetting error. Typically, allowing for a 10% error is
Molecular Detection of MEN2 251
adequate (e.g., the total master mix volume prepared for 10 samples [24 µL of
master mix/sample] would be 264  µL rather than 240  µL) (see Table 2). All
reagents should be kept on ice. Mix gently and aliquot 24 µL of the master mix
into each of the PCR reaction tubes.
3. Using a clean pipet tip for each specimen, aliquot 1 µL of DNA (at a concentration
of 250 µg/mL) into the appropriate PCR reaction tube. Thermocycling reaction con-ditions are as follows: an initial denaturation step for 2 min at 95°C; and an amplifi-cation step consisting of 30 cycles of denaturation for 30 s at 94°C, annealing for
30 s at 65°C, extension for 30 s at 72°C and a final extension step for 10 min at 72˚C.
After PCR amplification, samples may be stored indefinitely at 5°C.
4. PCR products are detected by agarose gel electrophoresis. Run samples on either
a 20 × 14 cm or 20 × 20 cm 2% agarose gel (1% agarose/1% NuSieve). Prepare
the gel by mixing 1.2 g each of agarose and NuSieve with 300 mL of 1X TAE.
Heat the resulting slurry in a microwave until the agarose is dissolved. Cool to
approx 50°C (just cool enough to touch), and add 30 µL of EtBr.
5. Pour the agarose into the gel mold. Use a comb to form the appropriate number of
sample wells.
6. To prepare the samples for electrophoresis, combine 2 µL of 6X sample buffer
and 10 µL of PCR product in a microcentrifuge tube. Load approx 10 µL of each
sample (including the positive control, the no-DNA control, and the pGEM lad-der) into the gel. Run the gel for 2 h at 90 V.
7. Following electrophoresis, take a Polaroid picture of the gel by placing the gel on
the UV transilluminator. Use the camera setting of f8 at 1 s. The template size is
approx 997 bp.
3.1.2. Sequencing of Exons 10 and 11
Refer to Fig. 2 for a diagram of the protocol.
1. Prepare the DNA template in a microcentrifuge tube by mixing 5 µL of each PCR
product with 1 µL of Exonuclease I (10 U/mL) and 1 µL of shrimp alkaline phos-Table 2
PCR Master Mix for PCR Amplification of RET Exons 10 and 11
Volume/sample Volume/10 samples
(does not include (includes 10%
pipetting error) (µL) pipetting error) (µL)
Sterile water 17.75 195.25
5X PCR buffer 5.00 55
Forward primer RET A2 0.5 5.5
Reverse primer CRT 19A 0.5 5.5
Taq DNA polymerase (5 U/µL) 0.25 2.75
Total volume 24.0 264.0
252 Rowsell and Wick
phatase. Incubate the mixture at 37°C for 15 min, and then heat the sample to
80°C for 15 min. Store the samples at 4°C or on ice until the remainder of the
sequencing reagents are prepared.
2. Prepare batches of each of the four termination mixes (_-33P-ddATP mix,
_-33P-ddCTP mix, _-33P-ddGTP mix, and _-33P-ddTTP mix) by mixing 2 µL of
dGTP nucleotide master mix (from the Amersham kit) with 0.5 µL of each of the
four _-33P-ddNTPs. Each individual sequencing reaction requires 2.5  µL of
the termination mix. Thus, it is more convenient to prepare enough of each of the
four termination mixes for the entire sequencing run. Termination mixes should
be prepared on ice.
3. Prepare a sequencing master mix for each sequencing reaction with the following
reagents (all volumes in microliters/reaction). As with the PCR master mix, it is
advisable to prepare a 10% excess of sequencing master mix; the volumes given
do not include the 10% pipetting error volume. Prepare the reactions on ice. Add
the ThermoSequenase last: 12.8 µL of sterile water, 2.0 µL of reaction buffer, 0.2 µL
of sequencing primer—Ret D or CRT 19B (20 µM), 2.0 µL of ThermoSequenase
DNA polymerase (total volume of 17.0 µL/reaction).
Fig. 2. Diagram of the MEN2A sequencing protocol. (Left) Preparation of a sample
mix (see Subheading 3.1.2., step 4c) for each patient by mixing the prepared DNA
template see Subheading 3.1.2., step 1) with the sequencing master mix (see Sub-heading 3.1.2, step 3); (Right) Aliquoting of the patient sample mix into each of the
individual sequencing reactions (A, C, G, T) for exons 10 and 11 of the RET gene (see
Subheading 3.1.2., step 4d).
Molecular Detection of MEN2 253
4. Prepare the sequencing reactions on ice as follows:
a. For each specimen (including the positive control) and for each exon to be
sequenced, label four PCR tubes with the specimen number (or identifier) and
either A, C, G, or T.
b. Add 2.5 µL of the appropriate d/ddNTP termination mix (see step 2) to the
appropriate tube.
c. Prepare a sample mix for each patient specimen (and the positive control) by
placing 3  µL of treated template (see step 1) into a microcentrifuge tube
labeled with the patient identifier. Then add 17 µL of the sequencing master
mix (see step 3) to each tube to create a sample mix. Vortex gently and spin
briefly in the microcentrifuge.
d. Add 4.5 µL of each sample mix (see step 3) to the appropriate A-, C-, G-, and
T-labeled tubes. Mix by repeatedly pipetting up and down.
5. Thermocycling conditions for the cycle sequencing reactions (both exons 10 and
11) are as follows (do not place samples into the thermocycler until the machine
has reached 95°C): an initial denaturation step for 2 min at 95°C; an amplifica-tion step consisting of 30 cycles of denaturation for 30 s at 94°C and annealing
and extension for 30 s at 70°C; there is no final extension. After thermocycling,
the reactions should be stored at –20°C.
6. Prepare the sequencing gel plates and apparatus as recommended by the
manufacturer.
7. Prepare the acrylamide gel mixture by mixing 60 mL of 6% polyacrylamide (at
room temperature) with 0.6 mL of 10% APS and 10.2 µL of TEMED. The APS
and TEMED should be added immediately prior to pouring the gel; a plastic
bottle with a nozzle works well for mixing and pouring the gel. The manufacturer
of the gel apparatus should provide instructions for pouring the gel. The gel
should polymerize within 1 h.
8. Conduct gel electrophoresis of sequencing reaction products as follows:
a. Prepare the samples for gel analysis by diluting 7  µL of each sample with
4 µL of loading buffer; mix by pipetting up and down.
b. Heat the samples in the thermocycler for 2 min at 72°C. Quickly cool the
samples in an ice/water slurry and keep them on ice until ready for loading.
Load the gel within 1 h of heating the samples.
c. It is important to flush the urea from the wells before loading the specimens.
Load 4 µL of each specimen into each well. To detect mutations more easily,
load all of the A reactions in adjacent wells, followed by all C reactions, all
G reactions, and all T reactions (Fig. 3).
d. Run the gel at 70 W for approx 100 min (about 15 min after the bromophenol
blue dye runs off the end of the gel).
e. Transfer the gel to blotting paper. Dry the gel on the gel dryer for 1 h.
f. Place the gel in an X-ray cassette, expose an X-ray film overnight at room
temperature, and develop the film.
254 Rowsell and Wick
3.1.3. Interpretation of Results
Mutations within exon 10 of the RET gene will appear as new bands on the
sequencing gel at codon 609, 611, 618, or 620; mutations within exon 11 will
appear as new bands at codon 634. Because patients are heterozygous for the
mutation, the intensity of the corresponding normal band will be decreased
(Fig. 3).
3.2. Heteroduplex Analysis of RET Exons 10, 11, and 13
3.2.1. PCR Amplification of Exons 10, 11, and 13
1. Set up the appropriate number of PCR reaction tubes, including a positive control
for each RET exon being analyzed, a negative control, and a blank (no DNA)
control.
2. Prepare a PCR master mix containing the following (in microliters per tube):
78.5 µL of sterile water, 20.0 µL of 5X PCR Reaction Buffer with dNTPs, 0.5 µL
of exon 10/11/13 forward primer, 0.5 µL of exon 10/11/13 reverse primer, 0.5 µL
of Taq DNA polymerase (total master mix volume per tube of 99.5  µL). It is
Fig. 3. Sequencing analysis of MEN2A. PCR amplification and sequencing of exon
10 demonstrates mutations in codon 618. (Note that exon 10 was sequenced in the
reverse direction from 3vA5v. The base designations have been modified to reflect the
sequence of the sense strand.) Lanes N and 2, a normal control and a normal patient,
respectively; lanes 1 and 3, the codon 618 TGCAAGC mutation; lane 4, the codon
618 TGCACGC mutation. The “extra” band in lane C4 (denoted by the bracket on the
right of the gel) is a sequencing artifact; it is thought to arise from mutation-induced
secondary structural changes in the DNA. (Figure 3 was provided by S. Thibodeau.)
Molecular Detection of MEN2 255
advisable to prepare a slight excess of master mix to allow for pipetting error.
Typically, allowing for a 10% error is adequate (e.g., if there are 10 samples,
prepare adequate master mix for 11). All reagents should be kept on ice. Mix
gently and aliquot 99.5 µL of the master mix into each PCR reaction tube.
3. Using a clean pipet tip for each specimen, aliquot 0.5  µL of patient, positive
control, and negative control template DNA (at a concentration of 250 µg/mL)
into the appropriate separate PCR reaction tubes. Aliquot 0.5 µL of sterile water
instead of template DNA into the blank tube.
4. Thermocycling reaction conditions are as follows: an initial denaturation step for
5 min at 94°C; an amplification step consisting of 35 cycles of denaturation for
1 min at 94°C, annealing for 1 min at 56°C, extension for 2 min at 72°C, and a
final extension step for 10 min at 72°C.
5. After PCR amplification, add 1  µL of 0.5  M EDTA (pH 8.0) to each tube
to inactivate the Taq DNA polymerase. PCR products may be stored indefinitely
at 5°C.
3.2.2. MDET Gel Electrophoresis of PCR Products
1. Prepare and pour the 1.2X MDET gel as follows:
a. MDE gel solution is supplied as a 2X concentrate. A 40 cm × 20 cm × 1 mm
gel requires about 80 mL of 1.2X gel solution. Prepare the correct amount for
your plates by adjusting each of the following components proportionally:
60 mL of MDE gel solution (2X), 6 mL of 10X TBE, 15 g of urea, 15 mL of
formamide, 100 mL of deionized water (fill to), 40 µL of TEMED, 400 µL of
10% APS (prepared fresh).
b. Mix the first five components in a clean beaker by gently swirling.
c. After the urea has dissolved, filter the solution through Whatman No. 1 filter
paper.
d. Add the specified amounts of TEMED and fresh 10% APS; mix the solution
by gently swirling.
e. Pour the gel solution into the plates using the standard procedure for
acrylamide. Insert a comb. Allow the gel to polymerize for at least 60 min at
room temperature.
2. Enhance heteroduplex formation by denaturing the PCR products for 5 min at
94°C and then annealing for 30 min at room temperature.
3. Mix 5  µL of PCR product with 1  µL of MDE gel loading buffer in a micro-centrifuge tube. Load the PCR product/buffer mixture onto the 1.2X MDE gel.
4. Run the gel at 20 V/cm for 35,000 V-h (the power supply should be set at 800 V
and run 43.75 h for a 40-cm plate).
5. After the run is completed, remove one of the glass plates. Leave the gel attached
to the other plate to ease handling during the staining and destaining steps.
6. Stain the gel for 10–15 min at room temperature in 1  µg/mL of EtBr solution
(made in 0.6X TBE buffer) (see Note 3). Adjust the staining, destaining, and
photography steps as necessary to achieve the desired results.
256 Rowsell and Wick
7. Destain the gel for 15–20 min in 0.6X TBE buffer. Destaining for up to an hour
may be necessary to eliminate background fluorescence, which may obscure faint
bands.
8. Invert the plate (gel side down) on a UV transilluminator to visualize the DNA
bands. (For easier handling, cut out the area of the gel containing the DNA bands
of interest, and then place the gel onto the UV transilluminator.)
9. Photograph the gel using Polaroid Polapan 667 film.
3.2.3. Interpretation of Results
The HET control DNA lane should contain two bands: a slower moving
HET DNA band and a faster moving homoduplex band (HET bands often run
as a single band) (Fig. 4). Homozygous normal or mutant samples are expected
to migrate as a single band (homoduplex). Depending on the mutation, the
PCR products of heterozygous DNA will form up to four bands: mutant/
mutant, normal/normal homoduplexes, plus two mutant/normal HETs. When
using EtBr staining, bands may appear broader because this staining technique
requires more DNA to be loaded onto the gel.
Fig. 4. HET analysis of MEN2A by multiplex PCR amplification of RET proto-oncogene exons 10, 11, and 13. Normal individuals (+/+) have single homoduplex
bands for all three exons. Additional bands represent chimeric HETs generated by the
presence of MEN2A-, MTC-, and FMTC-associated mutations in exons 10, 11, and
13. A normal polymorphism in exon 13 is present in a number of individuals. Muta-tions are indicated by codon number and altered nucleotide sequence. (Reprinted from
ref. 14  with permission by John Wiley & Sons, Inc. © 1996 Wiley-Liss, Inc.)
Molecular Detection of MEN2 257
3.3. SSCP Analysis of RET Exons 10, 11, and 13
3.3.1. PCR Amplification of Exons 10 and 11
1. Set up the appropriate number of PCR reaction tubes, including a positive control
for each RET exon being analyzed, a negative control, and a blank (no DNA)
control.
2. Prepare a PCR master mix containing the following (in microliters per tube):
11.25 µL of sterile water, 5.00  µL of 5X Reaction Buffer N, 2.00  µL of
dNTPs, 0.625 µL of exon 10/11 forward primer, 0.625 µL of exon 10/11 reverse
primer, 0.50 µL of AmpliTaq DNA polymerase (total master mix volume per tube
of 20.00 µL). Do not add the template DNA at this time. It is advisable to prepare
a slight excess of master mix to allow for pipetting error. Typically, allowing for
a 10% error is adequate (e.g., if there are 10 samples, prepare adequate master
mix for 11). All reagents should be kept on ice. Mix gently and aliquot 20 µL of
the master mix into each of the PCR reaction tubes.
3. Using a clean pipet tip for each specimen, aliquot 5 µL of patient, positive con-trol, and negative control template DNA (at a concentration of 250 µg/mL) into
the appropriate separate PCR reaction tubes. Aliquot 5 µL of sterile water instead
of template DNA into the blank tube.
4. Thermocycling reaction conditions are as follows: an initial denaturation step for
2 min at 94°C; an amplification step consisting of 35 cycles of denaturation for
1 min at 94°C, annealing for 1 min at 65°C, extension for 1 min at 72°C, and a
final extension for 10 min at 72°C. After PCR amplification, PCR products may
be stored indefinitely at 5°C.
5. To check amplification quality, analyze the PCR products by nondenaturing poly-acrylamide electrophoresis. Mix 10 µL of PCR product with 3 µL of gel-loading
buffer and run in 1X TBE on an 8% polyacrylamide gel.
6. Photograph the gel using Polaroid Type 667 film.
3.3.2. SSCP Gel Electrophoresis of PCR Products
1. Make a 1 10 dilution of each PCR product by mixing 1 µL of product with 10 µL
of sterile deionized water. Denature the diluted PCR products for 5 min at 95°C
in either a boiling water bath or a thermocycler and then quickly chill them on ice
for at least 15 min.
2. While the samples are on ice, prepare the PhastSystem.
3. Turn the instrument on and program it to cool down to ~10°C.
4. Clean the platform and place a dime-sized drop of water (approx 200 µL) on each
side of the separation beds.
5. Using forceps, place a 20% homogeneous polyacrylamide native PhastGel over
the drops of water. Make sure that no air bubbles are trapped under the gel.
6. Lower the applicator arm over the gel and place two PhastGel buffer strips in the
slots at each end of the gel.
7. Program the unit to cool down to 3 ± 1°C and start the programmed pre-run.
258 Rowsell and Wick
8. After the unit has prerun ~50 V-h, prepare the samples for automatic loading as
follows.
a. Either of two sample applicator sizes can be used: the 6/4 (6 samples, 4 mL
each per gel) or the 8/1 (8 samples, 1 mL each per gel).
b. Lay the applicator down on a piece of Parafilm® laboratory film (American
National Can™, Neenah, WI) and place ~1 mL of loading dye in alignment
with each applicator sample well.
c. Pipet the diluted, denatured, quick-chilled PCR product (11 mL) onto each
microliter of dye and gently mix. Take care not to introduce air bubbles into
the drop or mix the drop with adjacent drops.
d. Lift the applicator and place it vertically over each sample until the sample is
picked up.
e. Place the sample applicator with the loaded sample into the appropriate slot
on the separation unit.
f. After 100 V-h of prerunning, the sample will be automatically applied to the
gel.
g. Actual electrophoresis conditions must be determined empirically and pro-grammed into the unit. Example conditions are as follows:
a. Prerun: 400 V/5.0 mA/1.0 W/3°C for 100 V-h.
b. Application: 25 V/5.0 mA/1.0 W/3°C for 2 V-h.
c. Separation: 400 V/5.0 mA/1.0 W/3°C for 350–390 V-h.
9. After electrophoresis, wearing gloves and using only forceps, place the gel into
the PhastSystem Developing Unit (see Note 3).
10. Silver stain the gel using the PhastGel Silver Kit staining reagents and predeter-mined conditions.
11. Air-dry the gel for several hours at room temperature before mounting it in a
Kodachrome™ slide holder.
3.3.3. Interpretation of SSCP Gel Electrophoresis Results
The positive control DNA lane should contain bands with altered mobility,
as should the homozygous mutant and heterozygous patient sample lanes. By
contrast, homozygous normal patient lanes should have no bands with altered
mobility (Fig. 5).
3.4. RFLP-Based Detection of MEN2B
3.4.1. PCR Amplification of Exon 16
1. Set up the appropriate number of PCR tubes, including a positive control, a nor-mal control, and a blank (no DNA) control.
2. Prepare a PCR master mix containing the components in Table 3. It is advisable
to prepare a slight excess of master mix to allow for pipetting error. Typically,
allowing for a 10% error is adequate (e.g., the total master mix volume prepared
for 10 samples [24  µL of master mix/sample] would be 264  µL rather than
240 µL). All reagents should be kept on ice. Mix gently and aliquot 24 µL of the
master mix into each of the PCR reaction tubes.
Molecular Detection of MEN2 259
3. Using a clean pipet tip for each specimen, aliquot 1 µL of DNA (250 µg/mL) into
the appropriate tube.
4. Thermocycling reaction conditions are as follows: an initial denaturation step for
2 min at 95°C; an amplification step consisting of 35 cycles of denaturation for
30 s at 94°C, annealing for 30 s at 65°C, extension for 30 s at 72°C, and a final
extension for 10 min at 72°C. After PCR amplification, PCR products may be
stored indefinitely at 5°C.
Fig. 5. SSCP analysis of MEN2A by PCR amplification, denaturation, and gel elec-trophoresis (Phast system) of exon 11. The gel illustrates the wild-type SSCP pattern
in a normal control (NL), an abnormal SSCP pattern in a positive control (Pos),
an affected father (■ ), and his affected daughter (?). Arrows indicate the abnormal
conformer.
Table 3
Master Mix for PCR Amplification of RET Exon 16
Volume/sample Volume/10 samples
(does not include  (includes 10%
pipetting error) (µL) pipetting error) (µL)
Sterile water 14.25 156.75
50% glycerol 2.5 27.5
5X PCR buffer 5.00 55
Primer Ret 16F 1.0 11
Primer Ret 16R 1.0 11
Taq polymerase (5 U/µL) 0.25 2.75
Total volume 24.0 264.0
260 Rowsell and Wick
3.4.2. RE Digestion
1. Prepare a RE master mix containing the following (in microliters/per/sample)
3.0 µL of sterile water, 1.5 µL of 10X NEB Buffer 4, 0.5 µL of FokI. Mix gently
and aliquot 5 µL of the master mix into each of the reaction tubes. (As previously
noted, it is advisable to prepare a volume of master mix that allows for a 10%
pipetting error.)
2. Using a clean pipet tip for each specimen, aliquot 10 µL of amplified DNA into
the appropriate tube.
3. Carry out the digestion reaction at 37°C for a maximum of 2 h; longer digestion
may result in a decreased signal.
3.4.3. Agarose Gel Electrophoresis
1. To detect the PCR products by agarose gel electrophoresis, run the samples on
either a 20 × 14 cm or 20 × 20 cm 3% agarose gel (1% agarose/2% NuSieve).
Fig. 6. RE analysis of MEN2B. PCR amplification of exon 16 results in a 195-bp
fragment (uncut control), which is restricted by FokI into 109- and 86-bp fragments
(cut control). The  FokI site is destroyed by the MEN2B mutation in codon 918.
Affected individuals (patients) demonstrate a 195-bp fragment, which represents the
abnormal allele, and 109- and 86-bp fragments, which represent the normal allele.
Molecular Detection of MEN2 261
Prepare the gel by mixing 1.2 g of agarose and 2.4 g of NuSieve with 300 mL of
1X TAE. Heat the resulting slurry in the microwave until the agarose is dissolved;
cool to ~50°C (just cool enough to touch) and add 30 µL of EtBr.
2. Pour the agarose into the gel mold; use a comb to form the appropriate number of
sample wells.
3. To prepare the samples for gel loading, mix 15  µL of digest with 3  µL of 6X
sample buffer. Load the entire volume (18 µL) onto the gel.
4. Run the gel at 80 V for 2 to 3 h.
5. After electrophoresis, place the gel on a UV transilluminator. Take a Polaroid
picture of the gel using a camera setting of f11 and 1 s.
3.4.4. Interpretation of Results
PCR amplification (see Subheading 3.4.1.) results in a 195-bp amplicon.
FokI digestion of the amplicon in normal individuals results in two restriction
fragments of 109 and 86 bp. Because affected individuals are heterozygous, all
three bands will be present on the gel (Fig. 6).
4. Notes
1. The PCR and sequencing protocols described here were optimized with the PE
Applied Biosystems GeneAmp® PCR System 9600 and MicroAmp® Reaction
Tubes with Caps (nos. N801-0001 and N801-0540, respectively; PE Applied
Biosystems). The use of other systems may require reoptimization.
2. Other DNA stains may be used (and may be more sensitive).
3. Rinse the gloves with water prior to handling the gel. Glove powder can leave
artifacts on gels that appear after silver staining.
References
1. Trupp, M., Arenas, E., Fainzilber, M., et al. (1996) Functional receptor for GDFN
encoded by the c-ret proto-oncogene. Nature 381, 785–789.
2. Berndt, I., Reuter, M., Saller, B., Frank-Raue, K., Groth, P., Grussendorf, M.,
Raue, M., Ritter, M., and Höppner, W. (1998) A new hot spot for mutations in the
ret proto-oncogene causing familial medullary thyroid carcinoma and multiple
endocrine neoplasia type 2A. J. Clin. Endocrinol. Metab. 83, 770–774.
3. Marsh, D. J., Mulligan, L. M., and Eng, C. (1997) RET Proto-oncogene mutations
in multiple endocrine neoplasia type 2 and medullary thyroid carcinoma. Horm.
Res. 47, 168–178.
4. Decker, R. A. and Peacock, M. L. (1997) Update of the Profile of Multiple Endo-crine Neoplasia Type 2a RET Mutations. Cancer 80(Suppl.), 557–568.
5. Eng, C., Mulligan, L. M., Smith, D. P., et al. (1995) Mutations of the RET Proto-oncogene in sporadic medullary thyroid carcinoma.  Genes Chrom. Cancer 12,
209–212.
6. Heshmati, H. M. and Hofbauer, L. (1997) Multiple endocrine neoplasia
type 2: recent progress in diagnosis and management. Eur. J. Endocrinol. 137,
572–578.
262 Rowsell and Wick
7. Wells, S. A. Jr., Chi, D. D., Toshima. K., Dehner, L. P., Coffin, C. M., Dowton,
S. B., Ivanovich, I. L., DeBenedetti, M. K., Dilley, W. G., Moley, J. F., et al.
(1994) Predictive DNA testing and prophylactic thyroidectomy in patients at
risk for multiple endocrine neoplasia type 2A. Ann. Surg. 220, 237–247.
8. Decker, R. A., Peacock, M. L., Borst, M. J., Sweet, J. D., and Thompson,
N. W. (1995) Progress in genetic screening of multiple endocrine neoplasia type
2A: is calcitonin testing obsolete? Surgery 118, 257–264.
9. Carlson, K. M., Bracamontes, J., Jackson, C. E., et al. (1994) Parent of origin
effects in multiple endocrine neoplasia type 2B.  Am. J. Hum. Genet. 55,
1076–1082.
10. Schuffenecker, I., Billuad, M., Calendar, A., Chambe, B., Ginet, N., Calmettes,
C., Modigliani, E., and Lenoir, G. M. (1994) Ret Proto-Oncogene mutations in
French MEN2A and FMTC families. Hum. Mol. Genet. 3, 1939–1943.
11. Ceccherini, I., Hofstra, R. M. W., Luo, Y., et al. (1995) DNA polymorphisms and
conditions for SSCP analysis of the 20 exons of the RET proto-oncogene.
Oncogene 9, 3025–3029.
12. Siegelman, M. S., Mohabcer, A. J., Fahey, T., Tomlinson, G., Mayambala, C.,
Jafari, S., Noil, W. W., Thibodeau, S. N., and Dawson, D. B. (1997) Rapid, non-radioactive SSCP analysis for mutations in exons 10, 11, and 15 of the RET proto-oncogene associated with inherited medullary thyroid carcinoma. Clin. Chem. 43,
453–457.
13. Anonymous. (1998) Heteroduplex Analysis with MDE™ solution. FMC
BioProducts Catalog, FMC BioProducts, ME, pp. 93–98.
14. Kambouris, M., Jackson, C. E., and Feldman, G. L. (1996) Diagnosis of Multiple
Endocrine Neoplasia [MEN] 2A, 2B and Familial Medullary Thyroid Cancer
[FMTC] by multiplex PCR and heteroduplex analyses of RET proto-oncogene
mutations. Human Mutat. 8, 64–70.
15. Feldman, G. L., Kambouris, M., Talpos, G. B., Mulligan, L. M., Ponder, B. A. J.,
and Jackson, C. E. (1994) Clinical value of direct DNA analysis of the RET proto-oncogene in families with multiple endocrine neoplasia type 2A.  Surgery 116,
1042–1047.
Detection of I1307K within the APC Gene 263
20
Assay for Detecting the I1307K Susceptibility
Allele within the Adenomatous Polyposis Coli Gene
Stephen B. Gruber
1. Introduction
Most germline mutations of the adenomatous polyposis coli (APC) tumor
suppressor gene result in a classic inherited cancer syndrome called familial
adenomatous polyposis (FAP). FAP is characterized by thousands of colonic
polyps, well-defined extracolonic manifestations that may include pigmented
lesions of the ocular fundus, supernumerary teeth, osteomas, odontomas,
desmoid tumors and epidermoid cysts, and a 100% lifetime risk of developing
colorectal cancer. Shortly after the  APC gene was cloned in 1991  (1 ,2) the
molecular basis of an attenuated form of FAP was recognized to be related to
germline mutations within APC that were most likely to be found in the 5vand
3vends of the gene (3 ,4) . The truncating mutations leading to classic FAP and
attenuated FAP are quite rare, but recently a polymorphism of the APC gene
was found among 6 to 7% of Ashkenazi Jews that approximately doubles the
risk of colorectal cancer (5) .
The APC I1307K allele confers an increased risk of colorectal cancer by
creating a hypermutable tract of eight contiguous A residues, in contrast to the
wild-type (WT) sequence (A)3T(A)4. This poly-A tract functions as a
premutation that makes the surrounding DNA sequence especially vulnerable
to subsequent somatic mutations. Furthermore, this enhanced somatic muta-tion rate is completely restricted to the mutant (MU) allele, and the somatic
mutations are almost always insertions (6) . It is clear that I1307K increases the
carrier’s risk of colorectal cancer (7) , but recent evidence suggesting that this
allele also increases the risk of breast cancer remains controversial  (8 ,9) .
263
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
264 Gruber
Together these findings suggest that the I1307K allele may have relevant
implications for genetic counseling and risk prediction, even though the posi-tive predictive value for this test is relatively low.
Several techniques have been described for detecting the presence of the
APC I1307K allele (9) . Allele-specific oligonucleotide (ASO) hybridization is
used most commonly, because the technique permits MU and WT alleles to be
accurately distinguished at low cost for many samples (10) . Some laboratories
prefer to screen samples with single-strand conformation polymorphism or
conformation-sensitive gel electrophoresis  (11) followed by sequencing in
order to detect the I1307K allele, but in our experience each of these tech-niques is more time-consuming and expensive than ASO hybridization. Varia-tions of the radiolabeled ASO technique using chemiluminescent enzymatic
detection are also relatively straightforward to set up, but are more expensive
than the radioactive assay.
2. Materials
2.1. General Reagents
1. MilliQ purified, autoclaved double-distilled H2O (ddH2O).
2. 10X PCR Buffer: 500 mM KCl, 0.1% gelatin, 1% Triton X-100, 100 mM
Tris-HCl, pH 9.0, 15 mM MgCl2.
3. dNTP mix: final concentration of 25 mM dATP, 25 mM dCTP, 25 mM dGTP,
25 mM dTTP.
4. Taq polymerase (Perkin-Elmer).
5. 50X TAE buffer: 242 g of Tris base, 57.1 mL glacial acetic acid, 100 mL of 0.5 M
EDTA, pH 8.0, in 1 L of distilled water.
6. Loading buffer for agarose gels: 6X buffer consisting of 15% Ficoll, 60 mM
EDTA, 3% sodium dodecyl sulfate (SDS), 0.25% bromophenol blue, 0.25%
xylene cyanol (1.5 g of Ficoll, 1.2 mL 0.5 M EDTA, 3 mL 10% SDS, 25 mg of
bromophenol blue, 25 mg of xylene cyanol, in 10 mL of distilled water).
7. 2X saline sodium citrate (SSC): Diluted from a 20X SSC stock solution consist-ing of 175.3 g of NaCl and 88.2 g of sodium citrate in 800 mL of water. Adjust
the pH to 7.0 with a few drops of a 10 N solution of NaOH. Adjust the volume to
1 L with H2O and autoclave).
8. 20X SSPE: 175.3 g of NaCl, 27.6 g of NaH2PO4 ·H2O and 7.4 g of EDTA in
800 mL of water. Adjust the pH to 7.4 with NaOH (approx 6.5 mL of 10 N solu-tion), in 1 L of distilled water.
2.2. Sample Preparation
1. Lymphocyte separation media (LSM®; ICN Pharmaceuticals, Aurora, OH).
2. Phosphate-buffered saline (PBS) (Sigma, St. Louis, MO).
3. Puregene DNA Isolation kit (Gentra Systems).
4. Isopropanol stored at –20°C.
5. 70% Ethanol.
6. LoTE: 3 mM Tris-HCl, 0.2 mM EDTA, pH 7.6.
Detection of I1307K within the APC Gene 265
2.3. ASO Hybridization
1. 10X Polynucleotide kinase (PNK) buffer (New England Biolabs); store at –20°C.
2. PNK (New England Biolabs); store at –20°C.
3. a32P-ATP (Amersham Pharmacia).
4. 50X DET: DET is Denhardt’s with 10 mM EDTA and 10 mM Tris pH 8.0. 50X
Denhardt’s is made with 5 g of Ficoll (Type 400; Pharmacia), 5 g of polyvinyl-pyrrolidone, 5 g of bovine serum albumin (Fraction V; Sigma), and water to
500 mL.
5. 10% SDS.
6. Sheared, single-stranded salmon sperm DNA.
7. 0.4 M NaOH, 25 mM EDTA.
8. Hybond N+ nylon membrane (Amersham Pharmacia).
9. Trichloroacetic acid (TCA).
10. Metricel membrane filter (P/N 63068; Gelman Sciences).
11. Scintillation fluid (Beckman-Coulter).
2.4. Equipment
1. Centrifuge.
2. Thermocycler.
3. Agarose gel-running aparatus.
4. Dot-blot apparatus.
5. Adjustable temperature water bath with shaker.
6. Scintillation counter.
7. Heat sealer.
3. Methods
The technique for detecting the I1307K allele follows four steps:
1. Sample preparation.
2. Polymerase chain reaction (PCR) amplification of the relevant region of the
APC gene.
3. Allele-specific hybridization of oligonucleotide probes to PCR products that have
been immobilized on nylon membranes.
4. Washes at a critical temperature and stringency.
The ASO technique takes advantage of the difference in melting tempera-tures of oligonucleotides with differing sequences. Even a single nucleotide
change is sufficient to distinguish two variant alleles at a critical hybridization
temperature, and nonspecific hybridization is typically avoided by using oligo-nucleotide primers between 16 and 20 nucleotides long (10) . The performance
of oligonucleotide probes can sometimes be enhanced by designing probes
corresponding to the noncoding strand; the probes we use for the I1307K assay
are noncoding (L. Brody, personal communication).
266 Gruber
3.1. Sample Preparation
1. Transfer 3 mL of defibrinated or heparinized blood into a 15-mL conical tube
containing 3 mL of PBS and mix gently by inverting or pipetting.
2. Gently mix LSM by inverting. Then transfer 3.5 mL of LSM to a 15-mL conical tube.
3. Carefully layer the blood/PBS solution onto the LSM, keeping a sharp interface
between the dilute blood and the LSM.
4. Centrifuge the tube at 400g at room temperature for 15–30 min.
5. Aspirate the top layer of clear plasma to within 2 to 3 mm above the lymphocyte
layer.
6. Transfer the lymphocyte layer and about half of the LSM layer below it to
a centrifuge tube. Add an equal volume of PBS to the tube and centrifuge for
10 min at 200g, in order not to damage the cells. Remove the supernatant.
7. Wash the cells a second time by resuspending in PBS and centrifuge for 10 min
at 200g. Aspirate the supernatant, and vortex briefly in the small remaining
amount of residual supernatant to resuspend the cells.
8. For an average-sized cell pellet, add 3 mL of Cell Lysis Solution (Gentra Sys-tems) to the lymphocytes and pipet up and down to lyse the cells.
9. Add 1 mL of Protein Precipitation Solution and vortex for 20 s.
10. Centrifuge at 13,000–16,000g for 3 min.
11. Pour off the supernatant into ice-cold isopropanol.
12. Invert the tube about 40 times until the DNA precipitates. The DNA precipitate
should be visible as a white precipitate and can be collected by spooling the DNA
on a glass Pasteur pipet. Wash in 70% ethanol and resuspend the DNA in 200 µL
of LoTE.
13. If no precipitate can be seen, centrifuge for 1 min at 13,000g, pour off the isopro-panol, let the tube dry, and then add 150 µL of LoTE.
14. Measure the concentration of the DNA by removing an aliquot and measuring
the absorbance at 260 and 280 nm. Dilute an aliquot of DNA in ddH2O to a
concentration of 4 ng/µL.
3.2. PCR Amplification of APC
The oligonucleotide primers that are used to amplify the segment of APC
containing the I1307K polymorphism cover a larger region of the APC gene
than was originally reported by Laken et al. (5) . The primers shown in Table 1
give a single 372-bp PCR product visible as a single band on a 1% agarose gel.
Although the originally published primers work quite well by amplifying a
110-bp product, the primers shown in  Table 1 provide a longer product for
those investigators who are interested in using one set of primers to study somatic
mutations in a larger region surrounding the hypermutable tract of I1307K.
1. Aliquot 5 µL of DNA sample (4 ng/µL) into each well of a 96-well plate.
2. Make a PCR reaction mix for the appropriate multiple of PCR reactions (add the
Taq polymerase last, and keep reactions on ice prior to loading on a thermocycler)
using the following protocol (for each 20-µL individual reaction):
Detection of I1307K within the APC Gene 267
a. PCR buffer: 2 µL of 10X PCR buffer for a 1X final concentration.
b. dNTP: 0.16 µL of 25 mM dNTP mix.
c. Forward primer: 0.2 µL of 50 ng/µL primer for a total of 10 ng per reaction.
d. Reverse primer: 0.2 µL of 50 ng/µL primer for a total of 10 ng per reaction.
e. Taq polymerase: 0.2 µL of 5U/µL Taq polymerase for a total of 1 U/reaction.
f. ddH2O: 12.24  µL ddH2O brings the total volume of the reaction to 20  µL
when 5 µL of sample DNA is used.
3. Distribute 15 µL of the PCR mix to each sample and cover with mineral oil.
4. Program the thermocycler to denature at 95°C for 5 min; followed by 35 cycles
of denaturing at 95°C for 1 min; annealing at 53°C for 1 min, and extension at
72°C for 1 min; and ending with a final extension at 72°C for 10 min.
5. Visualize an aliquot of the PCR product on 1% agarose/TAE gel to ensure
adequate amplification. The product is visualized as a single band at ~370 bp.
3.3. ASO Hybridization
Figure 1 summarizes the general technique for ASO hybridization. PCR-amplified DNA products are denatured and immobilized on a nylon membrane,
usually in duplicate in order to make it more convenient to hybridize with WT
and MU probes at the same time. The membrane is prehybridized with solution
that contains a blocking agent such as Denhardt’s or BLOTTO (5% nonfat
dried milk in water with 0.02% sodium azide) to prevent nonspecific binding
of probes to the membrane. Radiolabeled oligonucleotide probes are hybrid-ized to the immobilized DNA fragments at a temperature that permits specific
nucleotide pairing. Excess probe is removed by washing each membrane at a
carefully titrated stringency to eliminate nonspecific binding.
1. Denature 10 µL of PCR product in 93 µL of 0.4 M NaOH and 25 mM EDTA.
2. Transfer the PCR product to a nylon membrane (Hybond N+) using a dot-blot
apparatus, rinsing each well twice with 50 µL of 2X SSC. Disassemble the appa-ratus and neutralize the membrane for 5 min in 2X SSC.
3. Crosslink the DNA to the membrane with 1200 J of UV irradiation.
Table 1
Primer and Oligonucleotide Probe Sequences
for APC I1307K ASO Hybridization Assay
Product length/
Primer/probe Sequence hybridization temperature
APC forward TCC ACA CCT TCA TCT AAT GCC 372 bp
APC reverse TAA ACT AGA ACC CTG CAG TCT GC
Wild type CTT TTC TTT TAT TTC TGC 44°C
Mutant CTT TTC TTT TTT TTC TGC 44°C
268 Gruber
4. Prepare a prehybridization solution with 25 mL of 20X SSPE, 10 mL of 50X
DET, 5 mL of 10% SDS, 1 mL of single-stranded salmon sperm  denatured DNA,
and add ddH2O to bring the total volume to 100 mL.
5. Heat the prehybridization solution to exactly 44°C, reserving 15 mL of the
prehybridization solution for hybridization.
6. Prehybridize at exactly 44°C for 1 h.
7. While the membranes are in the prehybridization solution, end-label the wild-type and mutant probes with a32P-ATP. Label each probe by incubating a 25-µL
labeling reaction containing 16.5 µL of ddH2O, 2.5 µL of 10X PNK buffer, 2 µL
of oligonucleotide probe (WT or MU), 1 µL of PNK, and 3 µL of a32P-ATP at
37°C for 30 min. Quench the reaction at 68°C for 10 min.
8. Determine the activity of each probe to ensure adequate incorporation of 32P and
comparability of WT and MU probes. This is done by adding 1 µL aliquots of
WT and MU probes to separate tubes containing 1 mL of 10% TCA on ice, and
Fig. 1. (A) PCR products immobilized on nylon membranes in duplicate; (B) ASO
hybridization performed separately with WT and MU probes.
Detection of I1307K within the APC Gene 269
then slowly pouring this over a Metricel membrane filter (Gelman Sciences). The
filter is rinsed several times with TCA and transferred to a scintillation vial con-taining Beckman scintillation cocktail. The activity is counted using a scintilla-tion counter. Activities >500,000 cpm are acceptable. Lower counts may give
weaker signals. The labeled oligonucleotide probes may also be separated from
unbound label using spin columns or other techniques.
9. Heat the reserved 15 mL of prehybridization solution to exactly 44°C before add-ing WT and MU probes to equal aliquots of this solution.
10. Hybridize at exactly 44°C for 1 h.
3.4. Washes and Detection of Mutation
1. Wash membranes briefly with 1X SSC/0.05% SDS at room temperature, and
then wash again in 1X SSC/0.05% SDS at 44°C for 20 min.
2. Wrap membranes in plastic wrap and expose to film for 2–4 h at –80°C, depend-ing on the activity of the probe.
3. Read the genotypes by visual inspection. Genotypes are generally not difficult to
interpret in the presence of adequate controls, but it is important to make sure that
the signal intensity of the WT and MU blots are comparable for known samples
before proceeding with interpretation. WT/WT homozygotes are recognized by a
strong signal on the WT blot and the absence of a signal on the MU blot. WT/MU
heterozygotes are detectable by observing signals on both blots. I1307K homozy-gotes are rare, but this genotype is recognized by a signal on only the MU blot.
4. Notes
1. The sample preparation technique is not critical to this assay, and virtually any
method of isolating genomic DNA is sufficient.
2. Resuspending the isolated lymphocytes in the small amount of residual PBS that
remains after aspiration makes the cell lysis much more efficient. Vortexing for
5–10 s is adequate.
3. There are simpler ways to lyse erythrocytes and isolate white cells for DNA
preparations, but we prefer this method because it yields a clean fraction of lym-phocytes that can be frozen for future transformation or harvested immediately.
4. Treating the cell lysate with RNase is not necessary, and eliminating the RNase
treatment saves the expense of a costly reagent.
5. Some investigators prefer to store genomic DNA in ddH2O because EDTA can
chelate magnesium in the buffer for PCR reactions. We prefer to use LoTE in
order to preserve the DNA stock solutions for long-term storage, and dilute
aliquots for analysis in ddH2O. This does not interfere with PCR reactions.
6. The membrane transfer is not difficult and can be performed with many different
apparatuses and conditions. Some laboratories prefer using a slot-blot apparatus
rather than a dot-blot apparatus; we prefer to use a 96-well dot blot to facilitate
simultaneous analysis of many samples. Neutralizing the membranes and DNA fol-lowing the denaturing reaction is recommended but is not critical because the assay
does not appear to be sensitive to whether the neutralization step is performed.
270 Gruber
7. Prehybridization of both membranes can be performed in the same solution, but
it is important for the DNA sides of the membranes to be exposed to the solution.
Keeping one membrane faceup and the other facedown permits excellent
prehybridization.
8. The hybridization temperature is critical to the success of the assay. Lower tem-peratures permit nonspecific hybridization, and higher temperatures may lower
the sensitivity of the test by making the hybridization conditions too stringent.
Washing at the correct stringency is the second most important aspect of the
assay, and for this ASO a temperature of 44°C is appropriate for both hybridiza-tion and washing.
9. Membranes that are sealed in plastic wrap (and have not dried out completely)
can be stripped and reprobed many times.
References
1. Groden, J., Thliveris, A., Samowitz, W., et al. (1991) Identification and character-ization of the familial adenomatous polyposis coli gene. Cell 66, 589–600.
2. Kinzler, K. W., Nilbert, M. C., Su, L. K., et al. (1991) Identification of FAP locus
genes from chromosome 5q21. Science 253, 661–665.
3. Spirio, L., Olschwang, S., Groden, J., Robertson, M., Samowitz, W., Joslyn, G.,
Gelbert, L., Thliveris, A., Carlson, M., and Otterud, B. (1993) Alleles of the APC
gene: an attenuated form of familial polyposis. Cell 75, 951–957.
4. Soravia, C., Berk, T., Madlensky, L., Mitri, A., Cheng, H., Gallinger, S., Cohen,
Z., and Bapat, B. (1998) Genotype-phenotype correlations in attenuated
adenomatous polyposis coli. Am. J. Hum. Genet. 62, 1290–1301.
5. Laken, S. J., Petersen, G. M., Gruber, S. B., et al. (1997) Familial colorectal can-cer in Ashkenazim due to a hypermutable tract in APC. Nat. Genetics 17, 79–83.
6. Gryfe, R., Di Nicola, N., Gallinger, S., and Redston, M. (1998) Somatic instabil-ity of the APC I1307K allele in colorectal neoplasia. Cancer Res. 58, 4040–4043.
7. Gruber, S. B., Petersen, G. M., Kinzler, K. W., and Vogelstein, B. (1999) Cancer,
crash sites, and the new genetics of neoplasia. Gastroenterology 116, 210–212.
8. Woodage, T., King, S. M., Wacholder, S., Hartge, P., Struewing, J. P., McAdams,
M., Laken, S. J., Tucker, M. A., and Brody, L. C. (1998) The APCI1307K allele
and cancer risk in a community-based study of Ashkenazi Jews. Nat. Genet. 20,
62–65.
9. Redston, M., Nathanson, K. L., Yuan, Z. Q., et al. (1998) The APCI1307K allele
and breast cancer risk. Nat. Genetics 20, 13–14.
10. Nollau, P. and Wagener, C. (1997) Methods for detection of point mutations: per-formance and quality assessment. Clin. Chem. 43, 1114–1128.
11. Ganguly, A., Rock, M. J., and Prockop, D. J. (1993) Conformation-sensitive gel
electrophoresis for rapid detection of single-base differences in double-stranded
PCR products and DNA fragments: evidence for solvent-induced bends in DNA
heteroduplexes. Proc. Natl. Acad. Sci. USA 90, 10,325–10,329.
Detection of HPVs by PCR and ISH 271
21
Detection of Human Papillomaviruses
by Polymerase Chain Reaction
and In Situ Hybridization
Elizabeth R. Unger and Suzanne D. Vernon
1. Introduction
The clinical utility of human papillomavirus (HPV) testing continues to be
the focus of much debate. The clear epidemiologic link of HPV infection with
the development of cervical intraepithelial neoplasia and invasive cervical can-cers (1) leads to widespread expectation that testing for HPV could improve
cervical cancer screening or aid in the triage of patients with abnormal cytol-ogy. The results of clinical trials of HPV testing are contradictory  (2) , and
widespread implementation of HPV testing is currently not recommended.
However, HPV testing is important in epidemiologic studies of cervical and
anogenital disease, and a large National Cancer Institute trial is currently evalu-ating HPV as an adjunct to cytology in cervical cancer screening.
Methods for diagnosing HPV are all dependent on detection of viral DNA,
because the agent cannot be cultivated in routine tissue culture and antibody
methods lack sensitivity. Detection of HPV DNA therefore requires analysis
of cellular material from the viral lesion. Methods for HPV detection are com-plicated by the fact that HPV is not a single virus, but a group of more than 75
closely related viruses. Typing is based on the viral genomic sequence and at
least 30 types infect the genital area. The types are considered high or low risk,
based on the frequency of their association with malignant lesions. The assay
format and sampling method both influence how often HPV will be detected.
The many variations of testing and sampling that have been and are being used
make comparisons among studies problematic.
271
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
272 Unger and Vernon
Southern blot hybridization was considered the gold standard for HPV
detection; however, this technique is no longer frequently employed. More
rapid methods that can easily accommodate limited patient samples are
favored. In this chapter, we include protocols for an L1 consensus polymerase
chain reaction (PCR) assay, as the most commonly used amplification assay,
and colorimetric in situ hybridization (ISH) assay, as a nonamplified method
that yields complementary information to PCR. ISH and PCR assays are both
applicable to DNA analysis in formalin-fixed archival tissues. Both assay for-mats require small amounts of tissue and thus conserve material essential for
clinical management. In addition, both formats tolerate some degradation of
target nucleic acids and can utilize nonradioactive detection methods (3) .
The HybridCapture HPV test (Digene Diagnostics) is Food and Drug
Administration approved and was designed for clinical laboratories. The assay
is a solution phase hybridization of sample DNA with an RNA probe. The
specific hybrids are selected (“captured”) using an antibody bound to a tube or
microtiter plate that specifically recognizes DNA-RNA hybrid molecules. The
same antibody linked to an enzyme is then used for detection with a chemilu-minescent substrate. Although there is no control for input DNA, the assay
yields semiquantitative values for the amount of HPV DNA. As currently mar-keted, the assay groups multiple HPV probes into high- and low-risk groups,
so type-specific of information is not obtained. The manufacturer supplies
detailed instructions with the kit and trains laboratories in the proper use of the
kit on request. Therefore, despite its clinical applicability, the HybridCapture
assay is not discussed in this chapter.
1.1. L1 Consensus PCR Assay
Laboratories are increasingly using PCR to detect HPV. Since there are more
than 75 different types of HPV (4) , primers designed to amplify the conserved
L1 region have become the most widely used in clinical and epidemiologic
studies. The MY09 and MY11 primers contain several degenerate nucleotides
and amplify a 450-bp region of L1 of at least 25 different types of anogenital
HPV (5) . The GP5+-GP6+ primers are a mixture of fixed nucleotide oligo-nucleotides that amplify a 150-bp region of L1 of a wide range of HPV types
by lowering the annealing temperature during PCR  (6) . The utility of both
primer sets has been compared and contrasted with each set having
advantages or disadvantages over the other (7–9) . The protocol described
herein is an optimized method for amplifying HPV DNA from fresh cervi-cal swab specimens, cervical lavage specimens, or formalin-fixed paraffin-embedded tissues using the MY09/MY11 primer set. The most reproducible
results are achieved when DNA is extracted from samples, and for this
Detection of HPVs by PCR and ISH 273
reason, sample  preparation is described in this protocol. Type-specific
PCR assays are described in the literature and for specific applications may
have advantages.
1.2. ISH for HPV
Colorimetric ISH is applicable to routinely processed, formalin-fixed paraf-fin-embedded tissue sections. The chief advantage of the method is that HPV
DNA is demonstrated within a morphologic context, allowing the tissue distri-bution of virus to be evaluated. In addition, the integration status of HPV cor-relates with the pattern of signal produced in the assay (3,10) . The methods of
colorimetric detection and interpretation are the same as those used for immu-nohistochemistry, making ISH the molecular technique with the greatest
potential for ready incorporation into diagnostic histopathology laboratories.
ISH is labor-intensive and requires extensive optimization of conditions in order
to achieve maximum sensitivity. The method described herein uses capillary gap
technology to introduce and remove reagents, minimizing the individual han-dling of slides (11–13) . Empirical adjustment of conditions is required if other
methods of reagent application, slide denaturation, and incubation are used.
ISH is conceptually straightforward, requiring tissue pretreatment (to allow
for probe penetration), denaturation of probe and tissue nucleic acids, hybrid-ization, washes, and detection. The method described is for formalin-fixed par-affin-embedded tissues. Methods of preparation of cell block controls and
probe labeling are also included, because these are essential to obtaining opti-mal results on a routine basis.
2. Materials
2.1. L1 Consensus PCR Assay
2.1.1. Preparation of Samples
1. Pro-par clearant (Anatech, Battle Creek, MI).
2. Proteinase K buffer: 50 mM Tris-HCl, pH 8.3, 1 mM EDTA, 0.5% Tween-20,
200 µg/mL of proteinase K.
3. MµlTI-Lid-Locs (cat. nos. C5000 and C6000; Marsh Biomedical).
4. Phenol/chloroform/isoamyl alcohol (25 24 1) (v/v/v).
5. Phase Lock Gel I Light (5 Prime-3 Prime, Boulder, CO).
6. 100% Ethanol (4°C).
7. 10 M Ammonium acetate.
2.1.2. HPV L1 Consensus and `-Globin PCR
1. Sterile double-distilled H2O (ddH2O).
2. 10X MgCl2-free PCR buffer (Roche).
3. 25 mM MgCl2 (see Note 1).
274 Unger and Vernon
4. 10 mM dNTPs.
5. 50 µM Working oligonucleotide primers (see Table 1).
6. Template DNA (test samples, HPV-positive and -negative control DNA).
7. Taq DNA polymerase (5 U/µL).
8. Mineral oil.
9. DNA-Erase™ (ICN Biomedicals, Aurora, OH).
2.1.3. Typing of HPV L1 Consensus Amplicons
2.1.3.1. PROBE LABELING
1. Reagents for PCR (see Subheading 2.2.).
2. Template HPV DNA: Clones of HPV-6, -11, -16, -18, -31, and -35 are available
from American Type Culture Collection. Other clones are available by request
from investigators.
3. PCR DIG Labeling Mix (cat. no. 1 585 550; Boehringer Mannheim).
4. Positively charged nylon membrane (cat. no. 1 417 240; Boehringer Mannheim).
5. 20X saline sodium citrate (SSC): 3 M NaCl, 0.3 M sodium citrate, pH 7.0.
2.1.3.2. PREPARATION AND HYBRIDIZATION OF AMPLICON DOT BLOTS
1. 10X Denaturation solution: 4 M NaOH, 100 mM EDTA.
2. 0.4 M NaOH.
3. 2X SSC.
4. Nylon membrane.
5. DIG-Easy Hyb (cat. no. 1 603 558; Boehringer Mannheim).
6. 2X Wash solution (2X SSC, 0.1% sodium dodecyl sulfate [SDS]).
7. 0.5X Wash solution (0.5X SSC, 0.1% SDS).
2.1.3.3. CHEMILUMINESCENT DETECTION
1. DIG Wash and Block Buffer Set (cat. no. 1 585 762; Boehringer Mannheim).
2. Antidigoxigenin-conjugated with alkaline phosphatase (cat. no. 1 093 274;
Boehringer Mannheim).
3. CDP-Star (cat. no. 1 759 051; Boehringer Mannheim).
4. Lumi-film (cat. no. 1 666 657; Boehringer Mannheim).
Table 1
Oligonucleotide Primer Information
Amplicon
Primer Origin Sequence (5v–3v) size Reference
MY09 HPV CGT CCM ARR GGA WAC TGA TCa approx 450 5
MY11 GCM CAG GGW CAT AAY AAT GGa
PC04 `-Globin GAA GAG CCA AGG ACA GGT AC   286
GH20 CAA CTT CAT CCA CGT TCA CC
aM = A or C, R = A or G, W = A or T, Y = C or T.
Detection of HPVs by PCR and ISH 275
2.2. In Situ Hybridization
Cervical cancer cell lines with well-defined characteristics and a known copy
number of HPV are suggested as a dependable and consistent control for many
HPV assays. When prepared as formalin-fixed paraffin-embedded cell blocks,
the cell lines form quite effective tissue controls for monitoring the sensitivity
of the ISH reaction. Suggested lines include Caski, 400–600 copies
HPV-16/cell; Hela, 10–50 copies of HPV-18/cell; SiHa, 1 to 2 copies of
HPV-16/cell; HTB-31, HPV negative. The HPV-16 in SiHa cells is near the
limits of detection of the assay, and results on this cell block clearly demon-strate when any component of the assay is failing.
2.2.1. Preparation of Cell Block
1. Dulbecco’s phosphate-buffered saline (D-PBS).
2. 10% Neutral buffered formalin.
3. Collodion (no. 4560-1; Mallinckrodt).
2.2.2. Probe Labeling
1. 10X Nick translation buffer: 500 mM Tris, pH 8.0, 50 mM MgCl2, 100 mM
`-mercaptoethanol, 100 µg/mL of nuclease-free bovine serum albumin (BSA).
Filter sterilize (0.2-µ filter) and store in 0.5-mL aliquots at –20°C.
2. DNase activation buffer: 10 mM Tris-HCl, pH 7.6, 5 mM MgCl2, 1 mg/mL of
nuclease-free BSA. Store at –20°C.
3. DNase I (10 µg/µL (Gibco-BRL, Gaithersburg, MD).
4. DNA Polymerase I/DNase I (0.4 U of polymerase and 40 pg of DNase/µL
(Gibco-BRL).
5. Nucleotide buffer solution: 0.5 mL of 10X nick translation buffer; 0.5 mL of
sterile ddH2O, 10 µL each of 10 mM solutions of dATP, dCTP, dGTP (Gibco-BRL). Store in 100-µL aliquots at –20°C.
6. Bio-11-dUTP (1.0 mM ) (Enzo).
2.2.3. Verification of Probe Size and Labeling
1. Biotinylated DNA ladder (Gibco-BRL).
2. Agarose, Sigma Type 2 (Sigma, St. Louis, MO).
3. 10X Running buffer: 0.3 M NaOH, 30 mM EDTA (store at room temperature).
4. 10X Gel buffer: 0.3 M NaCl, 30 mM EDTA (store at room temperature).
5. Ficoll dye solution: 50 mM Tris-HCl, pH 7.5, 20% Ficoll, 5 mM EDTA, 0.9%
bromophenol blue, 0.9% xylene cyanole FF (store at room temperature).
6. 20X SSC: 3 M NaCl, 0.3 M sodium citrate.
7. TS Brij 7.5: 0.1 M Tris-HCl, pH 7.5, 0.1 M NaCl, 5 mM MgCl2, 2.5 mL of 30%
Brij 35/L.
8. TS Brij 9.5: 0.1 M Tris-HCl, pH 9.5, 0.1 M NaCl, 50 mM MgCl2, 2.5 mL of 30%
Brij 35/L.
9. ddH2O.
276 Unger and Vernon
10. Neutralizing buffer: 3 M NaCl, 0.5 M Tris-HCl, pH 7.5.
11. Nitroblue tetrazolium chloride (NBT) stock: 50 mg of NBT (grade III, no. N6876;
Sigma), 0.5 mL of sterile dH2O, 0.5 mL of N,N-dimethylformamide (DMF). Store
at room temperature.
12. 5-Bromo-4-chloro-3-indolyl phosphate p-toluidine salt (BCIP) stock: 50 mg of
BCIP (no. 0885; Amresco) and 1.0 mL DMF. Store at 4°C.
13. Avidin–alkaline phosphatase conjugate (Dako). Store at 4°C.
14. 1% BSA/TS Brij 7.5: 1 g of BSA fraction V (no. A4503; Sigma) and 0.1 g of
sodium azide to a final volume of 100 mL in TS Brij 7.5. Filter sterilize and store
at 4°C.
15. McGadey reagent: 67 µL of NBT stock, 33 µL of BCIP stock, 10 mL of TS 9.5
Brij. Prepare within 4 h of use.
2.2.4. ISH Assay
1. 0.01 N HCl, 2.5 mL of 35% Brij 35/L.
2. TS Brij 7.5: see Subheading 2.2.3, item 7.
3. TS Brij 9.5: see Subheading 2.2.3, item 8.
4. ddH2O Brij: 2.5 mL of 30% Brij 35 to 1 L with ddH2O (store at room
temperature).
5. 2X SSC SDS/Brij: 2X SSC, 0.1% SDS, 2.5 mL of 35% Brij 35/L.
6. 0.2X SSC SDS/Brij: 0.2X SSC, 0.1% SDS, 2.5 mL of 35% Brij 35/L.
7. 0.1X SSC SDS/Brij: 0.1X SSC, 0.1% SDS, 2.5 mL of 35% Brij 35/L.
8. Nuclear Fast Red counterstain: 0.1 g of Nuclear Fast Red (Sigma) dissolved
with heating and stirring in 5% aluminum sulfate. Cool and filter through
Whatman no. 1 filter paper. Add one crystal of Thymol (Sigma). Store at room
temperature.
9. Cytoseal 60 mounting media (cat. no. 48212-154; VWR Scientific).
10. CrystalMount (no. M02; Biomeda).
11. NBT stock: see Subheading 2.2.3, item 11.
12. BCIP stock: see Subheading 2.2.3, item 12.
13. McGadey reagent: see Subheading 2.2.3, item 15.
14. Pepsin (no. P7012; Sigma)
15. Xylene (histologic grade).
16. Absolute alcohol.
17. 95% Alcohol.
18. Avidin–alkaline phosphatase conjugate (Dako). Store at 4°C.
19. HPV Probe set: All probes are nick translated with biotin (see Subheading 2.2.2)
and mixed in 1 mL of 45% formamide hybridization cocktail (Amresco). Store at
–20°C. Other HPV probe mixtures or single HPV probes may be prepared as
required. Final probe concentration is 1–1.5 µg/mL.
a. HPV-6/11 probe: 0.5 µg each of biotinylated HPV-6 and HPV-11.
b. HPV-16/18 probe: 0.5 µg each of biotinylated HPV-16 and HPV-18
c. HPV-31/33/35 probe: 0.5  µg each of biotinylated HPV-31, HPV-33, and
HPV-35.
Detection of HPVs by PCR and ISH 277
20. Control probe set: All probes are nick translated with biotin (see Subheading
2.2.2) and mixed in 1 mL of 45% formamide hybridization cocktail (Amresco).
Store at –20°C. Final probe concentration is 1 µg/mL.
a. HG probe (endogenous positive control probe): 1.0 µg of biotinylated human
placental DNA (Sigma).
b. pBR probe (negative control probe): 1.0  µg of biotinylated pBR322 DNA
(Gibco-BRL).
2.3. Equipment
2.3.1. L1 Consensus PCR Assay
1. Thermal cycler.
2. Horizontal gel electrophoresis apparatus.
3. UV transilluminator and camera for gel image documentation.
4. Heat block.
5. SpeedVac.
6. Microcentrifuge.
7. Bio-Dot Apparatus (cat. no. 170-6545; Bio-Rad, Hercules, CA).
8. UV crosslinker.
9. X-ray film developer.
10. Hybridization bags or roller bottles.
2.3.2. ISH Assay
1. MicroProbe Slide Stainer and accessories (reagent buckets, rubber and glass
isolons, and blotting pads) or Fisher Codon Automated Slide Stainer (see Fig. 1).
2. TissueTek slide carrier, staining rack, and buckets.
3. Convection oven.
4. Water bath, 37°C, with test tube rack.
3. Methods
3.1. L1 Consensus PCR Assay Methods
3.1.1. Preparation of Samples: Formalin-Fixed
Paraffin-Embedded Tissues
To monitor the histology of the lesion assayed in the PCR reaction, it is
good practice to request that serial 5-µ sections be cut, with levels 1 and 4
mounted on a glass slide and stained with hematoxylin and eosin (H&E) and
levels 2 and 3 combined into one sterile microcentrifuge tube for PCR analy-sis. To prevent carryover between cases, a new disposable microtome blade
should be used for each case, and the sections for PCR analysis should be
transferred from the microtome directly to the tube using sterile disposable
sticks. Sections cut for PCR are stored at room temperature until ready for
processing.
278 Unger and Vernon
1. Add 1 mL of Pro-par clearant to each sample and incubate for 5 min at 55°C.
2. Centrifuge at 10,000g in a microfuge for 5 min at room temperature. Remove the
Pro-par clearant and discard into an organic waste container. Repeat steps 1 and
2 twice to remove all paraffin.
3. To remove traces of Pro-par clearant, wash the pellet twice with 1 mL of ethanol,
followed by centrifugation at 10,000g in a microfuge for 5 min at room temperature.
4. Dry the pelleted cellular material briefly in a speed vac and resuspend in 200 µL
of proteinase K buffer.
5. Place a lid-loc on each tube and incubate overnight in a heat block at 55°C. Inac-tivate the proteinase K by heating at 100°C for 10 min.
6. Store the samples at –20°C until ready for amplification. Prior to PCR, centrifuge
each sample briefly to pellet any remaining cellular material. An aliquot (usually
5–10 µL) of the clarified sample is used in the amplification.
3.1.2. Preparation of Samples: Swabs or Cervicovaginal Lavage
1. Place cervical samples collected on dacron swabs or cytobrushes in 1 mL of col-lection fluid and transport to the laboratory. Briskly agitate or vortex the tubes
for several minutes to dislodge material from the collection device. Press and
swirl the swab or brush against the side of the tube to remove excess fluid. Dis-card the collection device into biohazard waste. Use a 100 µL aliquot of the fluid
for DNA extraction and store the remainder at –20°C.
2. For cervical vaginal lavage samples, collect the cellular material by low-speed
centrifugation. Discard the supernatant into biohazard waste. Resuspend the pel-let at a 1 10 dilution (v/v) in PBS or a minimum volume of 450 µL. Use 100 µL
for DNA extraction and store the remainder at –20°C.
Fig. 1. Schematic drawing of (A) the MicroProbe system demonstrating the stain-ing station,  (B) programmable temperature-controlled incubation chamber, and  (C)
the slide holder. (From ref. 17  with permission.)
Detection of HPVs by PCR and ISH 279
3. Add 200 µL of proteinase K buffer to each 100-µL sample, mix, and digest for
1 h at 55°C in a heat block.
4. Transfer the digested samples to a Phase-Lock tube and add an equal volume of
phenol chloroform isoamyl alcohol. Vortex to mix thoroughly and centrifuge at
10,000g for 10 min to separate the phases.
5. Transfer the aqueous (upper) phase containing the DNA to a clean sterile
microfuge tube and add 2 vol of ice-cold 100% ethanol and 1/10 vol of 10 M
ammonium acetate. Allow the DNA to precipitate at –20°C for at least 1 h. Pellet
the DNA at 10,000g for 15 min in a microcentrifuge. Remove the alcohol super-natant and allow the pellets to air-dry briefly.
6. Resuspend the precipitated DNA in 100 µL of sterile distilled deionized water.
Use 5–10 µL in the amplification reaction.
3.1.3. HPV L1 Consensus and `-Globin PCR
Excellent technique is required to avoid contamination of PCR samples and
the possibility of false-positive results. At least three distinct sites within the
laboratory should be used for the different steps of routine PCR. All surfaces
should be thoroughly cleaned with a DNA contaminant removal solution. The
initial PCR preparation including reagent master mixes is completed in the
first location. Test samples are added in a second location (we routinely use a
laminar flow hood without the airflow on and turn on the UV lamp when com-plete). Amplified samples are handled and analyzed in the third location. Each
location should have a dedicated set of micropipettors, and sterile, disposable,
aerosol block tips should be used. Finally, talking should be minimal when
setting up the reactions.
A set of positive, negative, and contamination controls should be included
for every 12 samples. For the HPV consensus PCR, an HPV-containing cell
line such as SiHa or CaSki (both containing HPV-16, described in Subhead-ing 2.2.) are good controls. Avoid the use of cloned HPV DNA unless used at
a very high dilution, because the high copy number easily results in cross con-tamination of samples. Human placental DNA is used as the negative control.
A tube that contains all the reagents, with water in place of the test sample,
serves as the contamination control. Another important control performed on
all samples in parallel with the HPV consensus PCR is a PCR that amplifies a
normal gene such as `-globin. These reactions test for inhibitors or insufficient
quantity of test sample.
1. Prepare a work sheet for the appropriate number of 50-µL reactions. (An
example worksheet is shown in Fig. 2.)
2. At the designated setup location, label the lid of the PCR tube (0.5-mL Eppendorf
tubes) with the sample number from the work sheet and the date.
3. Following the formula shown in Table 2, prepare two separate reagent master
mixes in 1.5-mL Eppendorf tubes, one for the HPV primers and the other for the
280 Unger and Vernon
Fig. 2. An example worksheet for HPV L1 consensus PCR that allows calculation
of reagent volumes, prompts inclusion of required controls, directs loading of agarose
gel, and documents PCR results. A Polaroid photograph of the ethidium bromide
(EtBr)–stained gel (see Fig. 3) should be attached to the worksheet.
Detection of HPVs by PCR and ISH 281
globin primers (the calculations are based on 12 test samples, 3 control samples,
plus one extra).
4. Dispense 45 µL of each master mix into the appropriate set of 0.5-mL microfuge
tubes.
5. Add two drops of mineral oil to each tube. (This step can be omitted if the
thermocycler has a condenser lid.)
6. Move to the designated area for handling patient samples. As directed on the
worksheet, add 5 µL of each test DNA to the appropriate tubes. Also add positive
and negative control samples and water blank to the appropriate tubes. (The con-centration of positive control DNA needs to be determined during optimization
of the protocol so that the resulting product is clearly detectable but does not
overwhelm the system, leading to cross contamination. We use 100 ng of CaSki
DNA as the positive control sample.)
7. Tightly cap the tubes and mix gently. Place the samples in the thermocycler
(should be located in a separate room) and program for the following
thermocycling profile: hold at 95°C for 5 min then cycle with 94°C to denature
for 1 min, ramp 10 s to 55°C to anneal for 1 min, ramp 10 s to 72°C to elongate
for 1 min, ramp 10 s back to 94°C; repeat the cycle profile 30 times. Follow the
30 cycles with a “topping off” elongation step at 72°C for 5 min.
8. After amplification, take the sealed sample tubes to the location designated for
analysis. Samples may be frozen at –20°C until needed.
9. Electrophorese 10 µL of each sample in a 1.5% agarose gel along with an appro-priate marker. Stain briefly in an EtBr solution containing 1 mg/L EtBr.
10. Examine the stained gel on a UV transilluminator and photograph for
documentation.
Table 2
Master Mixes for 16 Reactions Each
of MY09/MY11 and `-Globin Primer Sets
Master mix (µL)
Final Volume
Components  concentration per reaction (µL) HPV `-Globin
10X PCR buffer 1X 5 80 80
25 mM MgCl2 2.5 mM 58080
dNTP (each) 0.2 mM 1          16 (each) 16 (each)
50 µM MY09 0.5 µM 0.5 8 —
50 µM MY11 0.5 µM 0.5 8 —
50 µM PC04 0.5 µM 0.5 — 8
50 µM GH20 0.5 µM 0.5 — 8
Taq Polymerase 1.25 U 0.25 4 4
H2O — 29.75 476 476
282 Unger and Vernon
Fig. 3. Arrangement of dot-blot assay. The dot blot uses the 96-well format. (A)
The dot blot for probe and hybridization optimization can be arranged as shown. (B)
Template for evaluating test samples. Ten samples (t1–t10) can be placed on each
strip. Each strip is hybridized with the probe corresponding to the positive control dot
at the top of the strip.
Detection of HPVs by PCR and ISH 283
3.1.4. Typing of HPV L1 Consensus Amplicons
The advantage of an HPV consensus reaction is that all (or most) types of
HPV will yield a positive result; thus, one reaction may be used to determine
whether a sample has HPV DNA. However, further testing is required to deter-mine the type(s) of HPV in the sample. There are several different approaches
for typing the amplicon, such as restriction fragment length polymorphism (14) ,
single-strand conformation polymorphism (15) , solid-phase or solution hybrid-ization (SHARP Signal System, Digene Diagnostics) with either probe cock-tails or single type-specific probes, and, finally, direct sequencing of the PCR
product (Advanced Biotechnologies Inc.).
This section presents a solid-phase dot-blot hybridization assay with type-specific hybridization and chemiluminescent detection. This format permits
detection of multiple types of HPV present in a single sample, and because of
the chemiluminescent detection, it is highly sensitive. The limitation is that a
separate hybridization must be performed for each type. The following sec-tions include the protocols for synthesis of digoxigenin-11-dUTP-labeled HPV
probes, synthesis of the HPV DNA positive control PCR products, preparation
and hybridization of dot blots, and chemiluminescent detection.
At least one company is developing a simplified approach to typing L1
consensus PCR products. A reverse dot-blot format, called a line probe
assay, has HPV type-specific DNA immobilized as lines on a plastic strip
(16) . The patient sample is amplified and labeled during the consensus PCR
reaction (modified to include biotinylated primers). The labeled patient
material is then directly hybridized to the strip to detect the types of HPV
present in the assay.
3.1.5. Probe Labeling
To make HPV-specific probes, known HPV templates are amplified and
labeled with MY09/MY11 primers using a modified nucleotide mix that
includes digoxigenin-dUTP (DIG-dUTP). Include one PCR tube for each HPV
DNA template, plus a negative control and no-template control reaction for
every 10 tubes. The following procedure is designed for 100-µL reactions.
1. Add the components listed in Table 3 to a sterile 0.5-mL microfuge tube.
2. Move to the designated location for addition of template and add 5 µL (100 ng)
of the appropriate HPV DNA template to its respective tube.
3. Follow the PCR protocol outlined in Subheading 3.1.3.
4. Analyze 10 µL of each reaction by agarose gel electrophoresis and visualize the
products after EtBr staining. The molecular size of the products will be consider-ably larger than 450 bp because of the DIG-dUTP incorporation.
284 Unger and Vernon
5. After gel documentation, retain the gel. This gel is used to determine the effi-ciency of DIG-dUTP incorporation.
6. Blot DIG-labeled PCR products from the gel by downward capillary transfer to a
positively charged nylon membrane using 20X SSC. Allow the transfer to pro-ceed overnight.
7. After the transfer, rinse the membrane in 2X SSC and UV-crosslink the DNA to
the membrane.
8. Detect the DIG-labeled probes (as described in Subheading 3.1.7.) to estimate
the efficiency of the DIG-dUTP incorporation during the PCR.
3.1.6. Preparation and Hybridization of Amplicon Dot Blots
The sensitivity and specificity of the dot-blot assay using the DIG-labeled
probes must be validated before testing patient samples. The assay optimiza-tion dot-blot format should include unlabeled HPV type-specific amplicons
prepared from known HPV templates in the MY09/MY11 protocol. The opti-mized dot-blot assay should not allow cross hybridization between HPV types
to be detected, and type-specific hybridization should be easily detected. Fig-ure 3A gives an example of how to set up an optimization dot-blot assay.
1. Prepare the sample for alkaline transfer to the membrane. The sample is either
100 ng of control DNA (positive HPV type of control amplicons or negative
control placental DNA) or 10 µL of test PCR reaction (unknown amplicons to
be typed). Mix the sample with 10  µL of 10X denaturing solution and bring
to a final volume of 100  µL with sterile ddH2O. Incubate for 10 min at room
temperature.
2. Wet the nylon membrane in a dish of water, and assemble the nylon membrane in
the dot-blot apparatus according to the manufacturer’s instructions.
Table 3
DIG-Labeled Probe Synthesis PCR Mix
Reagent Final concentration Volume per reaction (µL)
10X PCR buffer 1X 10
25 mM MgCl2 2.5 mM 10
DIG dNTP mix 0.2 mM 10
50 µM MY09 0.5 µM 1
50 µM MY11 0.5 µM 1
Taq polymerase 2.5 U 0.5
H2O   — 62.5
Total volume 100
Detection of HPVs by PCR and ISH 285
3. Transfer the denatured DNA solution to the appropriate well in the assembled
apparatus. Use gentle vacuum to suck the solution through the membrane. Turn
the vacuum off after the solution is through to avoid drying the membrane.
4. Add 500 µL of 0.4 M NaOH to each well to rinse, and use gentle vacuum to suck
the solution through the membrane.
5. While maintaining the vacuum, disassemble the apparatus and mark the columns
with a waterproof ink pen for ease in cutting at a later point. Maintaining the
vacuum allows the wells to be visualized as depressions.
6. After the membrane is marked, release the vacuum and remove the membrane.
Rinse the membrane in 2X SSC and UV-crosslink the DNA to the membrane.
7. Use a ruler and pencil to draw a line between the columns of dots and cut the
strips as illustrated in Fig. 3A,B. Each strip will be approx 0.5 × 15 cm.
8. Prehybridize the membranes for 1 h at 70°C with 2 mL of DIG-Easy Hyb per
strip. The membrane strips can be hybridized in glass trays, small roller bottles,
or hybridization bags, but care should be taken to ensure that the strips do not
stick to one another.
9. Dilute each probe to a final concentration of 5 ng/mL of DIG-Easy Hyb, boil for
10 min, and place on ice.
10. Remove and discard the prehybridization fluid, and replace with the same vol-ume of hybridization solution containing the probe. Hybridize overnight at 70°C.
11. Remove the hybridization solution and save at –20°C. (We have found that these
probe solutions can be used at least one more time without significant loss of
signal.)
12. Remove the strips from the hybridization vessel, and rinse in excess 2X SSC
0.1% SDS to remove adherent hybridization cocktail. Add the strips into a plastic
dish containing 200 mL of 2X SSC and 0.1% SDS and wash twice for 5 min at
room temperature with gentle rocking. All the strips can be put into the same
container for washes.
13. Wash twice with 200 mL of 0.5X SSC and 0.1% SDS for 15 min each at 68°C
with gentle rocking.
14. Proceed to the chemiluminescent detection (see Subheading 3.1.7.).
3.1.7. Chemiluminescent Detection
Chemiluminescent detection is extremely sensitive, but requires careful
technique to keep the background reproducibly low. Membranes need to be
handled gently with forceps. Gloves should be worn for all steps. The dishes
used for washing, blocking, and detection steps should be rinsed with 0.1 N
HCl, washed with a soap solution, and rinsed with water. The acid wash inac-tivates any “environmental” or carryover alkaline phosphatase that could con-tribute to high background. Several membranes may be handled in the same
dish provided that they are not allowed to overlap (see Note 2).
1. Equilibrate the membrane in wash buffer (supplied in the DIG Wash and Block
Buffer Set) for 1 min.
286 Unger and Vernon
2. Transfer the membrane into a clean plastic dish and add sufficient blocking solu-tion to soak and cover the membrane completely. Cover and incubate at 37°C for
30–60 min.
3. During the blocking step, prepare the conjugate. Centrifuge the tube of conjugate
in a microcentrifuge at top speed for 5 min (this step pellets any aggregates that
may have formed).
4. Dilute the conjugate 1 10,000 in blocking solution (i.e., 5  µL of anti-DIG-alkaline phosphatase in 50 mL of 1X blocking solution) and mix gently by inver-sion. Pour the conjugate solution into a “dedicated” conjugate dish (this will help
minimize the alkaline phosphatase carryover). Make enough diluted conjugate to
cover the membrane (e.g., a 10 × 14 cm nylon membrane requires about 50 mL of
diluted conjugate).
5. Drain the blocking solution from the membrane and transfer the membrane to the
dish with diluted conjugate solution. Ensure that the membrane is covered evenly
with conjugate, cover the dish, and incubate at 37°C for 30 min.
6. Pour off the conjugate solution from the membrane and rinse the membrane with
wash buffer. Transfer the membrane to a fresh dish, and wash twice in 200 mL of
wash buffer at room temperature with gentle agitation, 15 min each wash.
7. Drain the final wash and transfer the membrane to a fresh dish with detection
buffer. Equilibrate for 5 min at room temperature.
8. Prepare 10 mL of a 1 100 dilution of CDP-Star in detection buffer.
9. Drain the detection buffer, add the 1 100 CDP-Star (chemiluminescent substrate
for alkaline phosphatase), and incubate for 5 min at room temperature.
10. Remove the membrane from the CDP-Star, and blot off excess CDP-Star by sand-wiching the membrane between blot paper.
11. Place the membrane into a plastic bag and expose to film. It is likely that a 1-min
exposure or shorter will be sufficient to determine the efficiency of DIG-dUTP
incorporation during the probe synthesis. Longer times will be required for
detection of dot-blot results.
3.1.8. Interpretation of L1 Consensus PCR Assay
All samples, except the water blank (contamination control), should amplify
the 286-bp region of  `-globin and the DNA product of that size should be
visible in each lane. The HPV positive control should amplify the 450-bp HPV
consensus product, and the negative control (human placental DNA) should
not produce any product. If the positive and negative controls give the
expected results, the assay may interpreted. Test samples that amplify `-globin
and have no HPV product visible are considered HPV negative. Those with an
HPV product are considered HPV positive, whether or not `-globin is ampli-fied. Test samples failing to amplify both `-globin and HPV cannot be inter-preted. These samples require further optimization or purification as described
in Notes 3–6. Figure 4 is a photograph of an EtBr-stained gel and shows the
results of an HPV consensus and `-globin PCR.
Detection of HPVs by PCR and ISH 287
3.1.9. Interpretation of Typing of HPV Consensus Amplicons
After chemiluminescent detection, the dot-blot optimization experiment
should result in specific hybridization to the positive control position dots and
to the HPV type-specific dot in the column (Fig. 5). There should be no hybrid-ization signal detected in the negative control and little to no cross hybridiza-tion to the other types of HPV. If conditions are satisfactory, dot blots can be
made for testing clinical samples using template as shown in Fig. 3B.
3.2. ISH Methods
3.2.1. Preparation of Cell Block
1. Harvest cells and wash two times with D-PBS. Check cell viability and adjust
cell concentration to 5 × 106 cells/mL by resuspending in 10% neutral buffered
formalin.
2. Allow the cell suspension to fix at room temperature for 1 h.
3. Using a labeling pencil, label the cassette with the name of the cell line, block
number, and year. Place one piece of Histowrap with each cassette.
Fig. 4. EtBr gel. Ten microliters of each PCR product is analyzed by gel electro-phoresis. The MY09/MY11 products are in the top half of the gel, and the correspond-ing `-globin PCR product for each sample is in the bottom half of the gel. Samples are
placed to correspond to the worksheet. Lanes 1–12 and 17–28 are clinical samples.
Lanes 13 and 29 are amplimers from Caski DNA, lanes 14 and 30 are amplimers from
placental DNA, and lanes 15 and 31 are water in place of DNA template. Lanes 16 and
32 contain molecular size markers. In this example, HPV-positive products are visible
in samples 2, 4, 10, 11, 12, as well as in the Caski control (lane 13). The water blank is
negative. All samples have amplifiable DNA except sample 8 (lane 24).
288 Unger and Vernon
4. Near the end of the fixation period, prepare collodion bags in a chemical fume
hood. Completely fill a clean 15-mL glass conical centrifuge tube with undiluted
collodion. Immediately pour the collodion back into the reagent bottle, rotating
the tube while pouring (collodion is reusable). Place the tube upside down in the
test tube rack and allow to dry approx 10–15 min. Touch the inside of the tube
to determine whether the bag is dry. Once prepared, the bag must be used within
10 min.
5. Pour the formalin-cell suspension into the bag and obtain the pellet by centrifug-ing for 5 min at 400–500g in a tabletop centrifuge.
6. Carefully decant the supernatant. Remove the bag from the tube by cutting around
the top edge with sharp-nosed tweezers. Twist the bag gently using the tweezers
and carefully lift and pull the bag away from the bottom of the tube.
7. Cut off the excess bag and fold down the top of the bag close to the surface of the
pellet.
Fig. 5. Chemiluminescent detection of digoxigenin-labeled HPV probes hybridized to
strips dotted with specific types of HPV. The format of the strips is exactly as shown in Fig.
3A (i.e., 9 dots per strip). There is no cross hybridization between types of HPV. This result
indicates satisfactory optimization of probe labeling and hybridization conditions.
Detection of HPVs by PCR and ISH 289
8. Place the folded bag in the center of the Histowrap. Wrap the Histowrap around
the pellet and bag to form a closed envelope. Place the wrapped pellet in the
cassette and close the lid.
9. Place the cassette in a 10% formalin container and take to the histology labora-tory for processing into paraffin block.
3.2.2. Probe Labeling
1. On ice, thaw the DNA to be labeled, Bio-11-dUTP, and one aliquot each of nucle-otide buffer solution and DNase activation buffer.
2. Prepare an appropriate dilution of DNase I. (Lot-to-lot variation will require
adjustment of this dilution, but a final dilution of 80 pg/µL is a good starting
point.) Remove the DNase I from the freezer and place on ice. Using a sterile
micropipet tip, remove 2 µL and place in a sterile microcentrifuge tube. Replace
the DNase I stock solution into the freezer. Add 250  µL of DNase activation
buffer to the 2-µL aliquot and mix gently. Transfer 2 µL of this dilution to a fresh
sterile tube using a clean sterile micropiet tip. Add 200 µL of DNase activation
buffer and mix gently. This last tube is the working dilution of DNase I. Discard
the other tubes.
3. For each DNA to be labeled, calculate the volume that equals 1 µg. Label one
sterile microcentrifuge tube for each reaction with identification of DNA, date,
and concentration and place on ice.
4. To the labeled tubes on ice add the following: 1–2 µL of diluted DNase I, __ µL
of DNA to be labeled (volume to equal 1 µg), 10.0 µL of nucleotide buffer solu-tion, 1.0  µL of Bio-11-dUTP, 5.0  µL of DNA Polymerase I/DNase I mixture,
and __ µL of sterile distilled water (to yield a final reaction volume of 50 µL).
Return the enzymes and Bio-11-dUTP to the freezer.
5. Close the lids of the tubes and mix gently. Briefly centrifuge to concentrate the
reagents in the bottom of the tube. Incubate in a 14°C ice/water bath for 1 h and
15 min.
6. Stop the reaction by transferring the tubes to a 70°C water bath for 10 min. Store
the labeled probes at –20°C until size and incorporation of label is verified (see
Subheading 3.2.3.). Once the size is verified, add 5 µL of EDTA solution to each
50-µL reaction and mix. Probes are stable at –20°C for at least 2 yr.
3.2.3. Verification of Probe Size and Labeling
Alkaline gel electrophoresis of nick-translated probes, followed by transfer
to nitrocellulose membrane and colorimetric detection, allows monitoring of
the incorporation of label and evaluation of the final size of the products. For
efficient tissue penetration, probes should be less than 300 bases. Because of
the strong alkaline environment, this procedure will not work for probes with
alkali-labile linkages of the affinity label.
1. Prepare a 2% agarose gel in 1X gel buffer in a minihorizontal gel electrophoresis
apparatus. Fill the buffer reservoirs and cover the gel with 1X running buffer.
290 Unger and Vernon
2. For each sample, label a microcentrifuge tube and add 7 µL of 1X running buffer,
1 µL of Ficoll dye solution and 1–3 µL of DNA (6–180 ng). Treat end-labeled
markers as sample. Mix and briefly centrifuge to collect liquid.
3. Run electrophoresis at 25 V for about 20 min to allow the samples to enter the
gel. Then increase to 75 V and run until the bromophenol blue dye front is about
three fourths of the way down the gel (approx 2.5 h).
4. Soak the gel in neutralization solution until the pH is <9.0. Monitor the pH of the
gel by touching the surface with pH paper. Neutralization will require about four
changes of 30 mL of buffer, 10 min/change. Rinse the gel in ddH2O.
5. Set up Southern blot with nitrocellulose filter using 20X SSC. Blot at room tem-perature for 2 h. (Overnight blotting can be done if more convenient.)
6. Wash the filter in 2X SSC for 5 min, air-dry, and UV-crosslink (Stratalinker).
Bake in an 80°C oven between filter paper for 1 h to overnight (at this point the
filter may be stored in the refrigerator).
7. Float the nitrocellulose membrane in a clean plastic tray filled with 1% BSA/TS
Brij 7.5. Once the membrane is wet, incubate with 1% BSA/TS Brij 7.5 for
10–30 min in a 37°C water bath.
8. Add 40 µL of avidin-alkaline phosphatase conjugate to 20 mL of 1% BSA/TS
Brij 7.5 and mix gently. Using forceps, transfer the nitrocellulose to a fresh plas-tic dish and cover with diluted conjugate. Incubate at room temperature for
10 min.
9. Remove the filter from the conjugate and transfer to a clean plastic dish with TS
Brij 7.5. Wash the nitrocellulose filter four times in TS Brij 7.5, 3 min/wash.
10. Wash the filter in TS Brij 9.5 and transfer the membrane to a clean plastic dish
with McGadey reagent. Incubate at 37°C until the signal reaches the desired
intensity. Remove the nitrocellulose, rinse in water, and air-dry. Keep the mem-brane as a record of the results.
3.2.4. ISH Assay
Formalin-fixed paraffin-embedded tissue sections for ISH should be cut at
5 µm and mounted from a protein-free tap water histobath on silanized or posi-tively charged glass (such as Fisher Plus). Tissue should be mounted within
0.2 mm of the end of the slide opposite the label. Slides are air-dried and stored
unmelted in a clean dust-free box until ready for use (see Note 7).
1. Select test and control slides for the assay: five slides of each specimen and con-trol are required for the complete probe set (HPV-6/11, HPV-16/18, HPV-31/33/
35, positive control [HG], negative control [pBR322]). Complete the work sheet
(see Fig. 6) with the proper pairing of slides to ensure that each tissue is matched
with respect to the digestion conditions and probe that will be used. Label the
frosted or painted end of each slide with alcohol/xylene stable marker, identify-ing the position it occupies in the MicroProbe slide holder and verify that the
work sheet matches this position.
Detection of HPVs by PCR and ISH 291
Fig. 6. An example ISH work sheet that allows for appropriate pairing of slides to
receive the same reagents, and directing of appropriate placement of digestion reagent
and probes for each slide. Positions of the holder correspond to positions in the isolons
(individual wells) in the staining station of the MicroProbe system.
292 Unger and Vernon
2. Place the test and control slides vertically (label at top, tissue at bottom) in a
TissueTek slide carrier and place the carrier upright in a 65–80°C convection
oven. Allow the slides to melt a minimum of 20 min. Air-dry the slides in the
carrier under the chemical fume hood.
3. Prepare a Tissue-Tek staining rack under the chemical fume hood for dewaxing
of slides. Fill three green solvent-resistant buckets with fresh xylene and two
white buckets with fresh absolute alcohol. Dewax the slides in TissueTek slide
carrier at room temperature. Perform three 10-min incubations in xylene and two
5-min incubations in absolute alcohol. Remove the slide carrier and slides from
the last alcohol wash and air-dry under the fume hood. Discard the xylene in a
xylene waste container (see Notes 8–9).
4. Match up the slide position number on the slide with the position on the work
sheet. Form capillary gaps with slide pairs and a 40 × 22 mm no. 2 cover slip by
sandwiching the cover slip between the frosted (label) ends of the slides with the
tissues facing each other (Fig. 7). Load the slides into the proper position in the
MicroProbe slide holder. Make sure the slide pairs are flush against the base of
the holder.
5. Add water to the reservoir of the incubator and set the temperature to 37°C.
Fill the MicroProbe reagent buckets with appropriate buffers as designated on
the work sheet. Insert the reagent isolons and blotting pad in the MicroProbe
workstation.
Fig. 7. Placement of slides to produce capillary gap. A “sandwich” is made using
two slides and a no. 2 cover slip. The cover slip keeps a narrow gap between the slides.
Slides are placed so that the tissues face the inside of the gap. The gap will draw fluid
up to cover the inside surface of the glass. Fluid is removed from the gap by blotting
onto absorbent material (pad).
Detection of HPVs by PCR and ISH 293
6. Prepare the digestion reagent. Weigh 8 mg of pepsin and transfer into a clean test
tube. Add 2 mL of 0.01 N HCl/Brij (final concentration of 4 mg/mL). The diges-tion reagent should be made up fresh prior to each run and placed in a 37°C water
bath for 15 min to ensure that the enzyme has dissolved. Make less concentrated
solutions of digestion reagent as needed by dilution with 0.01 N HCl/Brij. Diges-tion reagent is varied empirically to achieve optimal signal with endogenous posi-tive control probe. Suggested starting concentrations are as follows: 4 mg/mL for
cell blocks, 2 mg/mL for biopsy tissue, 3 mg/mL for surgical resection, 4 mg/mL
for autopsy tissue.
7. Fill the reagent isolons with the appropriate concentration of protease. Isolons
may be filled with a transfer pipet; approximately 200 µL is required for each
position. The position of the well determines which slide pair will receive the
reagent. Verify reagent placement by consulting the work sheet.
8. Wash the slides with 95% alcohol, by placing the holder in the reagent bucket
with 95% alcohol and allowing the reagent to completely fill the gaps between
the slides by capillary action (see Note 10). Remove the holder and transfer to a
blotting pad to remove the reagent from the gaps by absorption of fluid. Make
sure the entire surface of each tissue is covered with each reagent at each step of
the assay and that the blotting steps completely drain each capillary gap. This
cycle of reagent pickup and removal by capillary action is repeated throughout
the assay. The term gap refers to the capillary gap between the slides. Washing
the gaps, or adding reagent to the gaps, is synonymous with adding reagent or
washing the tissues on the slides.
9. Wash the gaps with ddH2O Brij and blot. Pick up the digestion reagent from the
isolon, aligning the well and slide gaps. Transfer the holder to a 37°C incubator.
Set the timer for 30 min.
10. While the slides are in the digestion step, remove the probe set from the freezer
and allow them to reach room temperature.
11. Fill the appropriate wells of the glass isolon with 155 µL of probe, as indicated
on the work sheet.
12. At the end of 30 min of digestion, remove the holder and change the temperature
of the incubator to 105°C. Blot to remove protease reagent and wash the gaps
four times with TS Brij 7.5. For each wash, allow the reagent to cover the area of
gap briefly and then blot to completely remove all the reagent.
13. Wash the gaps twice with 95% alcohol and twice with 100% alcohol. Allow the
gaps to air-dry briefly.
14. Pick up the probe from the glass isolon. Make sure the holder and isolon are
properly aligned so that each gap receives the appropriate reagent. The probe
enters the gap slowly because of the viscosity of the hybridization cocktail. Watch
carefully during this process to be sure that the probes do not mix. Make sure
each capillary gap is filled at least 2 mm above the height of each tissue. Add
additional probe if needed.
15. After the incubator has reached 105°C, transfer the holder to the incubator and
set the timer for 20 min. After 20 min, change the incubator temperature to 37°C.
294 Unger and Vernon
Add water to the reservoir to ensure that the chamber is moist, and let the slides
hybridize for 2 h.
16. After the hybridization time is complete, remove the holder from the incubator
and change the incubator temperature to 42°C. Remove the cocktail from the
gaps by blotting. The solution is usually difficult to remove, so after some has
come out in the blotting pad, transfer the slides to 2X SSC SDS/Brij to dilute the
cocktail. Repeat this procedure several times until the gaps are completely emp-tied of cocktail. Then start stringency washes.
17. Wash the gaps three times with 2X SSC SDS/Brij and three times with 0.2X SSC
SDS/Brij. Pick up the 0.1X SSC SDS/Brij and transfer the holder to the incubator
for 5 min (42°C). Repeat three times. Change the incubator temperature to 37°C.
18. Fill the gaps with blocking reagent and remove by blotting. Repeat addition of
block to the gaps and transfer the holder to the incubator (37°C) for 5 min.
19. Make up the detection reagent using a 1 400 dilution of Dako avidin-alkaline
phosphatase in 1% BSA TS7.5/Brij. Fill each isolon well with 200  µL of the
diluted detection reagent.
20. Remove the block by blotting and pick up the conjugate. Transfer the holder to
the incubator (37°C) for 20 min.
21. Remove the conjugate by blotting and wash the gaps four times with TS Brij 7.5
and then twice with TS Brij 9.5.
22. Fill two reagent wells with 5 mL of freshly prepared McGadey reagent.
23. Wash the gaps with McGadey reagent and blot. Then refill the gaps with
McGadey reagent and transfer the holder to incubator (37°C) for 1 h.
24. Toward the end of the incubation, fill the reagent isolons with approx 200 µL of
Nuclear Fast Red counterstain.
25. At the end of incubation, remove the McGadey reagent by blotting and wash the
gaps twice with TS Brij 7.5 and three times with ddH2O Brij.
26. Fill the gaps with stain and keep in place for 2 min. Then remove the stain by
blotting and wash the gaps twice with ddH2O Brij.
27. Carefully remove the slides from the holder and place them flat, tissue side up,
on a metal tray. Put a small thin layer of CrystalMount over each tissue section
and bake for 10 min at 60°C, to harden the CrystalMount. (Alternatively, the
slides can be air-dried overnight.) Allow the slides to cool to room temperature
and cover slip each tissue with Cytoseal 60.
27. Label the slides with the date and the type of probe. Review the slides with the
pathologist and record the quality control data.
3.2.5. Interpretation of ISH
3.2.5.1. PROBE LABELING AND VERIFICATION
Color development should result in dark purple where biotinylated DNA
is localized. DNA in the marker lane should be visible as distinct bands
with good resolution. If the DNA ladder is not resolved, the procedure must be
repeated.
Detection of HPVs by PCR and ISH 295
Nick-translated probes appear as a smear in the lane as the probes are ran-domly sized. To be acceptable for use, the probe should be less than 300 bases
and average about 200 bases. Probes that are not clearly visible after this pro-cess or that are too large cannot be used. If probes are too large, attempts can
be made to renick them by incubation with DNase. Once probe size is verified,
EDTA is added to stabilize the probes and inhibit any residual DNase. If EDTA
is added prior to verifying the size, renicking cannot be achieved because of
inhibition of DNase by EDTA.
3.2.5.2. IN SITU HYBRIDIZATION
The control blocks Caski and Hela should be positive with HPV 16/18 and
negative with the other probe groups. SiHa cells are at the limit of sensitivity of
the assay and should be weakly positive with HPV 16/18. Assays in which
Caski or Hela controls do not give appropriate results must be repeated. A
negative result with SiHa is acceptable but indicates that one or more compo-nents of the assay could be failing.
The positive control probe (human placental DNA; HG) demonstrates
whether the assay conditions allow hybridization of DNA in the test tissue.
Tissues hybridized with the HG probe should have an even dark blue-black
signal over every nucleus. Hybridization with the negative control probe
(pBR322) should result in no detectable signal. If these two conditions are met,
the hybridization with the HPV probes may be interpreted (Fig. 8). Signal with
the HPV probe is seen as blue-black primarily over the nuclei of infected cells
containing the target DNA. Some samples may demonstrate reaction with more
than one group of HPV probes. This is most often owing to cross hybridization
among the probe groups, and the sample should be typed as the group giving
the strongest signal. In some instances, the intensity of multiple probe groups
is identical, indicating the presence of more than one type of HPV. The assay
may be repeated with more stringent washes if the type of HPV is questionable.
The ISH assay must be interpreted within the context of the histologic lesion
and the clinical setting. It is not intended to replace routine histopathologic
diagnosis, but to provide additional information in a manner analogous to
immunohistochemical assays. Interpretation must be made by a pathologist.
The H&E slide of the original lesion must be reviewed by the pathologist to
verify that the recut sections assayed by ISH are representative of the lesion.
4. Notes
1. This protocol uses a final MgCl2 concentration of 2.5 mM, found to be optimal
for the HPV MY09/MY11 consensus PCR in our hands. The optimal MgCl2 con-centration may vary depending on the primer synthesis and should be tested by
assaying control samples with low (1.5 mM), moderate (2.5–3.5 mM), and high
(5 mM) MgCl2 concentrations.
296 Unger and Vernon
Fig. 8. Appropriate controls for ISH. Probe labeling, hybridization and detection are as
described in the text. Results are for serial sections of the same tissue (nuclear fast red
counterstain, BCIP/NBT substrate, bar = 5 µm). (A) Human placental DNA. The positive
control probe results in a dark even signal over all nuclei, indicating that the tissue DNA
is preserved and that tissue is adequately treated to allow hybridization  (B)  pBR322.
The negative control probe shows no signal, indicating no apparent problems with non-specific reactions of probe and detection reagents. (C) HPV probe. Results can be inter-preted as indicative of HPV. (Modified from ref. 3  with permission.)
Detection of HPVs by PCR and ISH 297
2. Boehringer Mannheim’s guide to the use of digoxigenin affinity labels in filter
hybridizations includes an extensive trouble-shooting guide.
3. Patient samples that fail to amplify globin may have insufficient intact DNA,
may require a different MgCl2 concentration, or may contain an inhibitor of
the Taq polymerase. Adjusting the input DNA by increasing or decreasing the
sample volume, or varying the MgCl2 concentration, may result in successful
amplification.
4. The heme component of blood is a potent inhibitor of Taq polymerase. Visible
blood in clinical samples should be recorded in the notes during purification,
because failure to amplify the globin control in these samples may be owing to
residual heme. Although the digestion and extraction of DNA described in Sub-headings 3.1.1. and 3.1.2. is frequently sufficient for DNA amplification, occa-sionally further steps must be taken to remove blood components. Several
commercially available DNA purification kits are effective in purifying DNA
from blood or blood-containing samples (e.g., from QIAGEN, Santa Clarita, CA).
Alternatively, hydrogen peroxide treatment of blood-containing samples has been
shown to be effective in decomposing the heme compound (17) .
5. Degradation of template DNA can affect amplification of larger amplicons. The
450-base MY09/MY11 amplicon is nearing the size limit of 500 bp recommended
for formalin-fixed paraffin-embedded archival tissues. If degradation of sample
DNA is a problem, other HPV consensus primers that amplify smaller fragments
may be tried; for example, the GP5+-GP6+ amplifies a 155-bp region of HPV
DNA (6) .
6. Contamination is detected when the “no template reaction” has an amplicon. The
fastest remedy is to discard all open reagents and begin again with fresh reagents.
All PCR locations should be thoroughly cleaned with a DNA contaminant-removal solution, such as DNA Erase. The exterior of all micropipets should be
wiped with a DNA contaminant-removal solution and exposed to UV light for
15 min on each side.
7. The importance of good histologic sectioning is often overlooked. Sections must
be of uniform thickness without folds or tears, and the thickness must be uniform
from section to section to allow even and reproducible penetration of reagents.
Tissue adherence to glass is also crucial. Conditions for the hybridization assay
are drastic, and the loss of tissue may occur during the assay if steps are not taken
to improve tissue adherence. Treatment of the glass slides with 3-amino-propyltriethoxysilane has largely replaced other techniques such as coating the
glass with poly-L-lysine or glue. When tissue sections are cut and floated on a
protein-free tap water bath, the silane-treated glass will form a covalent bond
with the tissue section. This “silanized glass” is available commercially and
greatly minimizes the problem of tissue adherence. The quality of silanization
may be monitored by the behavior of tissue sections during cutting. If the glass is
properly treated, the tissue will not be able to be “refloated” or moved around on
the slide once it has been lifted from the surface of the water. Difficulties with the
procedure can be attributed to poorly treated glass, dirty glass, or protein in the
298 Unger and Vernon
water bath. Once the section is picked up on the microscope slide, the slide is
placed vertically and allowed to air-dry. The paraffin should be retained on the
section (i.e., section not melted) until just before the assay is begun.
8. The paraffin must be completely removed for efficient reagent penetration.
Xylene is the most efficient clearing agent, but xylene substitutes may be used if
care is taken to ensure that dewaxing steps are efficient. Several changes of
dewaxing agents are recommended. Deparaffinization conditions that are satis-factory for routine staining may not be adequate for ISH. Overtreatment with
dewaxing agents has not been observed.
9. Ideally, the time and temperature of fixation should be standardized and when
processing to the paraffin block fresh reagents should be used. These ideal condi-tions are seldom met. Significant variations in fixation and processing that dra-matically influence nucleic acid preservation may go unnoticed by standard light
microscopic examination. Until histology laboratories develop methods to
ensure standardization of fixation and processing, ISH assays on diagnostic
samples must include controls that will detect and adjust for variations in tissue
preservation. The purpose of controls is to ensure that tissues without signal are
devoid of the target nucleic acid and that tissues with signal do contain the target.
Interpretation of a precipitated product as evidence of a particular nucleic acid
Table 4
Controls for ISHa
Control Requirement Purpose
Positive control 1. Processed in manner identical Positive results verify
tissue (handle as     to test tissue.     reaction of probe and
additional tissue 2. Known to contain target     detection  reagents.
sample)     that hybridizes to
test probe.
Positive control 1. Hybridizes with a target present Positive result verifies
probe (use on     in all tissues.     preservation of nucleic
each tissue) 2. Labeled in similar manner     acid and availability
to test probe and used at     to probe.
similar concentration.
Negative control 1. Probe of similar base pair Negative  results
probe (use on     composition to test probe that     monitor specificity
each tissue)     should not hybridize to test     of hybridization
and control tissues.     and detection.
2. Labeled in similar manner
to test probe and used at
similar concentration.
aModified from ref. 18 . Used with permission.
Detection of HPVs by PCR and ISH 299
sequence requires that all other explanations be eliminated. Controls also allow
problems in any assay to be detected so that appropriate corrective measures can
be instituted. There are many different kinds of controls that can be included, but
the three essential controls are listed in Table 4. These three are the minimum
required by the College of American Pathologists’ Checklist for Molecular
Pathology.
10. Brij 35 is added to most buffers used in the ISH protocol to lower surface tension
and improve flow through the capillary gap. Flow of reagents must be efficient.
Occasional variations in glass thickness or the use of a cover slip of different
thickness may cause problems with flow. When slides are first assembled in the
holder, flow of the reagent should be evaluated for each slide pair. Readjusting
the position of the slide or changing the cover slip usually corrects problems
with flow.
Acknowledgments
We gratefully acknowledge Donna L. Miller and Daisy R. Lee for their
expert technical contributions to standardization of the protocols.
References
1. Schiffman, M. H. and Burk, R. D. (1998) Human papillomaviruses, in Viral Infec-tions of Humans: Epidemiology and Control, 4th ed. (Evans, A. S. and Kaslow,
R. A., eds.), Plenum, New York, pp. 983–1022.
2. Jenkins, D., Sherlaw-Johnson, C., and Gallivan, S. (1998) Assessing the role of
HPV testing in cervical cancer screening. Papillomavirus Rep. 9, 89–101.
3. Unger, E. R., Vernon, S. D., Lee, D. R., Miller, D. L., and Reeves, W. C. (1998)
Detection of human papillomavirus in archival tissues: comparison of in situ
hybridization and polymerase chain reaction.  J. Histochem. Cytochem. 46,
535–540.
4. De Villiers, E. M. (1994) Human pathogenic papillomaviruses: an update. Curr.
Top. Microbiol. Immunol. 186, 1–12.
5. Ting, Y. and Manos, M. (1990) Detection and typing of genital human
papillomaviruses, in PCR Protocols: A Guide to Methods and Applications (Innis,
M. A., Gelfand, D. H., Sninsky, J. J., and White, T. J., eds.), Academic, New
York, pp. 356–367.
6. De Roda Husman, A. M., Walboomers, J. M., Van Den Brule, A. J. C., Meijer,
C. J., and Snijders, P. J. (1995) The use of general primers GP5 and GP6 elon-gated at their 3v ends with adjacent highly conserved sequences improves human
papillomavirus detection by PCR. J. Gen. Virol. 76, 1057–1062.
7. Qu, W., Jiang, G., Yvette, C., Chang, C. J., Ho, G. Y. F., Klein, R. S., and Burk,
R. D. (1997) PCR detection of human papillomavirus: comparison between
MY09/MY11 and GP5+/GP6+ primer systems. J. Clin. Microbiol. 35, 1304–1310.
8. Zehbe, I. and Wilander, E. (1996) Two consensus primer systems and nested poly-merase chain reaction for human papillomavirus detection in cervical biopsies:
a study of sensitivity. Hum. Pathol. 27, 812–815.
300 Unger and Vernon
9. Baay, M., Quint, W., Koudstaal, J., Hollema, H., Duk, J. M., Burger, M., Stolz,
E., and Herbrink, P. (1996) Comprehensive study of several general and type-specific primer pairs for detection of human papillomavirus DNA by PCR in par-affin-embedded cervical carcinomas. J. Clin. Microbiol. 34, 745–747.
10. Cooper, K., Herrington, C. S., Strickland, J. E., Evans, M. F., and McGee, J. O.
(1991) Episomal and integrated human papillomavirus in cervical neoplasia
shown by non-isotopic in situ hybridization. J. Clin. Pathol. 44, 990–996.
11. Unger, E. R., Hammer, M. L., and Chenggis, M. L. (1991) Comparison of 35S and
biotin labels for in-situ hybridization: use of an HPV model system. J. Histochem.
Cytochem. 39, 145–150.
12. Unger, E. R., Brigati, D. J., Chenggis, M. L., Budgeon, L. R., Koebler, D., Cuomo,
C., and Kennedy, T. (1988) Automation of in-situ hybridization: application of
the capillary action robotic workstation. J. Histotechnol. 11, 253–258.
13. Chenggis, M. L. and Unger, E. R. (1993) Application of a manual capillary action
workstation to colorimetric in-situ hybridization. J. Histotechnol. 16, 33–38.
14. Lungu, O., Wright, T. C., and Silverstein, S. (1992) Typing of human
papillomaviruses by polymerase chain reaction amplification with L1 consensus
primers and RFLP analysis. Mol. Cell. Probes 6, 145–152.
15. Rubben, A., Traidl, C., Baron, J. M., and Grussendorf-Conen, E. I. (1995) Evalu-ation of non-radioactive temperature gradient SSCP analysis and of temperature
gradient gel electrophoresis for the detection of HPV 6-variants in condylomata
acuminata and Buschke-Loewenstein tumors. Eur. J. Epidemiol. 11, 501–506.
16. Gravitt, P. E., Peyton, C. L., Apple, R. J., and Wheeler, C. M. (1998) Genotyping
of 27 human papillomavirus types by using L1 consensus PCR products by a
single-hybridization, reverse line blot detection method. J. Clin. Microbiol. 36,
3020–3027.
17. Akane, A. (1996) Hydrogen peroxide decomposes the heme compound in foren-sic specimens and improves the efficiency of PCR. Biotechniques 21, 392–394.
18. Unger, E. R. and Lee, D. R. (1995) In situ hybridization: principles and diagnostic
applications in infection. J. Histotechnol. 18, 203–209.
Detection of Epstein-Barr Virus 301
22
Molecular Methods for Detecting
Epstein-Barr Virus (Part I)
In Situ Hybridization to Epstein-Barr
Virus–Encoded RNA (EBER) Transcripts
Hongxin Fan and Margaret L. Gulley
1. Introduction
In situ hybridization (ISH) to Epstein-Barr virus (EBV)–encoded RNA
(EBER) is considered the “gold standard” for detecting and localizing latent
EBV in biopsy samples. Transcripts from the EBER1 and EBER2 genes are an
appropriate target because they are the most abundant viral RNA in latently
infected cells, exceeding 1 million transcripts per cell. Furthermore, EBERs
are consistently expressed in virtually all EBV-infected tumors and in the lym-phoid tissues of infectious mononucleosis (1–5) . As a result, EBERs represent
a naturally amplified target for detecting and localizing latent EBV in histo-logic samples. The only EBV-related disorder in which EBER is consistently
absent is oral hairy leukoplakia, an infection of squamous epithelial cells in
which the virus undergoes lytic viral replication rather than latent infection (6) .
EBER ISH relies on the use of riboprobes (7,8) , oligonucleotide probes (9) ,
or peptide nucleic acid probes (Dako, Carpinteria, CA) to identify EBER1 or
EBER2 transcripts in histologic samples. Even though EBER transcripts are
abundantly expressed, these and other species of RNA are subject to degrada-tion by ubiquitous RNase enzymes, potentially causing loss of signal in all or
part of the tissue. To avoid false-negative EBER results, and to properly inter-pret the morphologic and cytologic distribution of EBER in human tissues, it is
imperative that a control stain be run alongside every EBER stain to ensure
that RNA is preserved and available for hybridization. Some investigators
301
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
302 Fan and Gulley
target poly-A tails of mRNA as a control, whereas others target cellular U6
transcripts because they are similar in copy number to EBERs and are likewise
localized to the nucleus.
There are several clinical situations in which EBER ISH imparts diagnostic
or prognostic information. In transplant recipients with lymphoproliferative
lesions, the assay is used to help distinguish EBV-driven posttransplant
lymphoproliferative disorder from organ rejection or other inflammatory con-ditions (10) . The assay is also helpful in confirming a diagnosis of infectious
mononucleosis or nasopharyngeal carcinoma. EBERs are sometimes detected
within tumor cells of Hodgkin disease, AIDS-related lymphoma, and lymphoe-pithelioma-like carcinomas arising in various anatomic sites such as the stom-ach; however, the clinical utility of EBER testing is limited because only a
fraction of these tumors harbor EBV, and clinical management does not
depend on whether EBV is present.
The procedure described herein is a 1-d ISH assay targeting EBER1 and U6
control RNA using digoxigenin (DIG)-labeled riboprobes, a procedure adapted
from one first introduced by Wu et al.  (1) and Barletta et al.  (7) . Types of
samples amenable to EBER ISH include paraffin-embedded tissues, frozen
sections, and cytology preparations.
2. Materials
2.1. Preparation of Probes
1. HindIII, EcoRI, BamHI restriction endonuclease.
2. RNase-free TE buffer: 10 mM Tris-HCl, pH 8.0, 0.1 mM EDTA. (see Note 1).
3. Diethylpyrocarbonate (DEPC)-treated H2O (see Note 1).
4. 25X labeling reaction buffer: 1  M Tris and 150 mM MgCl2 made with
DEPC-H2O.
5. 100 mM Spermidine.
6. 100 mM Dithiothreitol (DTT).
7. Acetylated bovine serum albumin (BSA) (10 mg/mL).
8. RNasin® Ribonuclease inhibitor (40 U/µL) (Promega, Madison, WI).
9. 100 mM each of r-ATP, r-GTP, r-CTP, and r-UTP.
10. Digoxigenin-11-UTP (250 nmol/25  µL) (Boehringer-Mannheim, Mannheim,
Germany).
11. T3 and T7 RNA polymerase (Promega).
12. RNase-free DNase (Promega).
13. 0.4 M EDTA, pH 8.0.
14. RNase-free tRNA (10 mg/mL).
15. 4 M LiCl.
16. 100% Ethanol.
17. 70% Ethanol made with DEPC-H2O.
Detection of Epstein-Barr Virus 303
18. Buffer 1: 100 mM Tris-HCl, 150 mM NaCl, pH 7.5.
19. Blocking reagent (Boehringer-Mannheim).
20. Blocking buffer: 1% (w/v) blocking reagent, dissolved in buffer 1.
21. Anti-DIG-alkaline phosphatase (Boehringer-Mannheim), stored at 4°C.
22. Buffer 2: 100 mM Tris-HCl, 100 mM NaCl, 50 mM MgCl2, pH 9.5. (The proper
pH is critical to the color reaction. Filter before each use.)
23. Buffer 3: 10 mM Tris-HCl, 1 mM EDTA, pH 8.0.
24. Labeled control RNA (Boehringer Mannheim).
25. Template DNA for EBER1 and U6 (plasmid available from Richard Ambinder,
MD, PhD, Johns Hopkins University).
26. Nitroblue tetrazolium chloride (NBT) solution: 75 mg/mL in 70% N,N, dimethyl-formamide (DMF) (see Note 2).
27. 5-Bromo-4-chloro-3-indolyl phosphate p-toluidine salt (BCIP) solution: 50 mg/mL
in 100% DMF (see Note 3).
2.2. In Situ Hybridization
1. Xylene.
2. Graded ethanol solutions: 50, 70, 80, and 95% made with DEPC-H2O.
3. 10X phosphate-buffered saline (PBS) made with DEPC-H2O.
4. Proteinase K (10 mg/mL) in RNase-free 100 mM Tris-HCl, 50 mM EDTA,
pH 7.5.
5. Digestion buffer: 100 mM Tris-HCl, 50 mM EDTA, pH 8.0, 0.3% Triton X-100.
6. Formamide (molecular biology grade); store at 4°C in the dark.
7. 20X saline sodium citrate (SSC): 3 M NaCl, 0.3 M sodium citrate, pH 7.4.
8. 100X Denhardt’s reagent made with DEPC-H2O: 2% Ficoll 400 (type 400), 2%
polyvinylpyrrolidon), 2% BSA (Fraction V).
9. 20% sodium dodecyl sulfate (SDS) solution.
10. 0.4 M EDTA, pH 7.5.
11. Sonicated salmon sperm DNA (10 mg/mL); boil before use.
12. Yeast tRNA, RNase-free (10 mg/mL) (Sigma, St. Louis, MO).
13. 75% Dextran sulfate made with DEPC-H2O.
14. Hybridization solution: 50% formamide, 5X SSC, pH 7.4, 5X Denhardt’s
reagent, 1% SDS, 1 mM EDTA, pH 7.5, 100 µg/mL of boiled sonicated salmon
sperm DNA, 460 µg/mL of tRNA, 7% dextran sulfate, made using RNase-free
stock solutions.
15. RNase A (Sigma; concentration varies by lot number).
16. Sheep serum.
17. Triton X-100.
18. Wash solution: 2X SSC + 0.1% SDS.
19. Wash solution: 0.1X SSC + 0.1% SDS.
20. Methyl green counterstain: 1% in H2O.
21. Permount.
22. Rubber cement.
304 Fan and Gulley
2.3. Equipment
1. Silane-coated slides (or slides otherwise treated to promote tissue adherence).
2. 55°C Water bath.
3. 80°C Vacuum oven.
4. Vacuum centrifuge.
5. Microscope.
6. Cover slips.
7. PAP PEN™ (Daido Sangyo, Japan).
8. Moist chamber (RNase-free).
9. Microcentrifuge.
10. Slide-staining baths (Tissue-Tek®II, Sakura Finetek).
3. Methods
3.1. Preparation of Probes
Single-stranded RNA probes (riboprobes) complementary to target RNA
transcripts are generated by transcribing cloned DNA sequences using T3 or
T7 RNA polymerases in the presence of DIG-labeled UTP. Transcription and
labeling are accomplished using commercial kits or by the following method.
3.1.1. General Comments
Gloves should be worn during probe preparation procedures to avoid RNase
contamination. All solutions should be RNase-free (see Note 1) as well as all
glassware and plasticware (see Note 4).
3.1.2. Preparation of DNA Templates
The DNA constructs that serve as templates for probe production are RA386
(EBER1) and RA390 (U6). The RA386 construct represents the full-length
458-bp EBER1 gene in a 2746-bp M13BS + pBS vector (Strategene, La Jolla,
CA), and the U6 construct represents U6 sequences in the BS-SK vector.
1. Digest each construct with an appropriate restriction enzyme (see Note 5).
2. Purify the digested construct by phenol/chloroform extraction and ethanol pre-cipitation, and then resuspend in 20 µL of RNase-free TE buffer at a concentra-tion of 1  µg/µL. This purified DNA is used as a template for producing
DIG-labeled riboprobes.
3.1.3. Labeling of Probes
1. For each labeling reaction, prepare 41.25 µL of labeling reaction mixture on ice
by combining the following reagents: 15.25  µL of DEPC-H2O, 2  µL of 25X
labeling reaction buffer; 1 µL of 100 mM spermidine; 0.5 µL of 100 mM DTT;
2.5 µL of 10 mg/mL acetylated BSA; 2  µL of RNasin; 5  µL of each 10 mM
r-ATP, r-GTP, r-CTP; 3 µL of 10 mM r-UTP.
Detection of Epstein-Barr Virus 305
2. To each reaction tube, add 1.75  µL of DIG-UTP, 1  µg of linearized template
DNA brought up to 5 µL with DEPC-H2O, and 2 µL of RNA polymerase (either
T3 or T7 RNA polymerase per Note 5) for a total reaction volume of 50 µL.
3. Incubate at 37°C for 2 h, adding another 1  µL of the appropriate RNA poly-merase after the first hour.
4. Digest the residual DNA template by adding 1 µL of 1 U/µL RNase-free DNase
and incubating at 37°C for 10 min.
5. Stop the reaction by adding 1 µL of 0.4 M EDTA, pH 8.0.
6. To precipitate the labeled RNA probe, add 2 µL of 10 mg/mL tRNA, mix, and
then add 0.1 vol of 4 M LiCl and 2.5–3.0 vol of chilled 100% ethanol. Mix and
incubate at –70°C overnight.
7. Remove the reaction from the freezer and centrifuge at 12,000g at 4°C for
15 min.
8. Decant the ethanol supernatant and wash the pellet with 100  µL of 70% cold
ethanol, centrifuge at 12,000g at 4°C for 5 min, and then remove as much ethanol
supernatant as possible.
9. Spin the tube in a vacuum centrifuge until the pellet is completely dry. Add
50 µL of DEPC-H2O and mix well by pipetting up and down a few times. Take
out 2 µL to test for adequacy of labeling (see Subheading 3.1.4.), and immedi-ately store the rest of labeled RNA probe at –70°C. (Note: Minimize freeze/thaw
cycles and always use appropriate tissue controls to ensure probe integrity.)
3.1.4. Probe-Labeling Test
Estimate the incorporation of DIG label into the RNA probe using the fol-lowing “spot test” protocol, which is essentially the same as that described in
the Boehringer-Mannheim package inserts accompanying the probe labeling
and detection reagents.
1. Pipette 1 µL of each reaction and dilutions thereof (1 10, 1 100, and 1 1000 in
DEPC-H2O) onto a small nylon membrane, along with equivalent dilutions of
control-labeled RNA.
2. Bake the membrane at 80°C for 20 min in a vacuum oven.
3. Wash the membrane 1 min in buffer 1 with shaking.
4. Incubate the membrane for 30 min in blocking buffer with shaking at room
temperature.
5. Wash the membrane for 1 min in buffer 1 with shaking.
6. Dilute the antibody conjugate 1 5000 in buffer 1 (20 mL of buffer 1, 4  µL of
anti-DIG-alkaline phosphatase) and use immediately.
7. Incubate the membrane in antibody diluent for 30 min with shaking.
8. Wash two times in 100 mL of buffer 1 for 15 min with shaking. Equilibrate in
20 mL of buffer 2 for 2 min with shaking.
9. Place the membrane in a bag and prepare the color reaction buffer in a darkened
room (10 mL of filtered buffer 2, 45  µL of NBT solution, 35  µL of BCIP
solution).
306 Fan and Gulley
10. Add the color reaction buffer to the bag and seal; do not shake. Store in the dark
for about 20–40 min or until the color is adequate.
11. Wash the membrane for 5 min with shaking in 50 mL of buffer 3 to stop the
reaction. If desired, store in a sealed bag containing buffer 3 to preserve color.
12. To interpret probe labeling, each probe should be labeled at least as strongly as a
1 10 dilution of the control probe, as judged by comparing the intensity of spot
colors. If not, the probe labeling reaction should be rerun using half or twice the
volume of linearized template DNA. Probe specificity and adequacy of labeling
are further verified by actual ISH to control tissues.
3.2. In Situ Hybridization
3.2.1. General Comments
All steps through the hybridization procedure must be RNase free (see Notes
1 and 4). The following incubations are at room temperature unless otherwise
stated. Before starting, prepare eight slide baths, each 200 mL in volume,
containing xylene; graded ethanols (100, 95, 80, 70, and 50%); 1X PBS;
and 20 µg/mL of proteinase K solution (200 mL of digestion buffer prewarmed
to 37°C; when ready for use, add 400 µL of proteinase K stock solution and
mix well).
3.2.2. Preparation of Specimens for Hybridization
1. From a fixed, paraffin-embedded tissue block, cut 4-µm sections onto silane-coated
slides using a sterile water bath and gloves to reduce RNA degradation. (Tissues
fixed in 10% buffered formalin more consistently retain RNA than do those fixed
in B5 or subjected to decalcification.) To prepare cytology specimens, see Note 6
and then go directly to step 4. For frozen sections, go directly to step 4.
2. Place paraffin section slides in an RNase-free staining rack for steps 2–4. Each
experiment should include appropriate controls including positive and negative
tissue controls, and sense as well as antisense probes (see Note 7). Bake the slides
at 80°C for 40 min under a vacuum if recently cut, or 20 min if thoroughly dry.
Immediately (while the paraffin is still hot) immerse the slides in fresh xylene for
6 min to deparaffinize.
3. Rehydrate the slides for 1 min each through graded ethanols (100, 95, 80, 70, and
50%) and then DEPC-PBS.
4. Digest the protein by immersing the slides in 20 µg/mL of proteinase K in diges-tion buffer at 37°C for 6 min (see Note 8).
5. Dip in 1X DEPC-PBS for 1 min to diminish proteinase K activity. Dab the slides
on Kimwipes to remove excess proteinase K, and lay them flat in an RNase-free
chamber in preparation for hybridization.
3.2.3. Hybridization
1. Add DIG-labeled EBER1 or U6 riboprobe to the hybridization solution at a 1:20
dilution. (The precise dilution may vary with each new batch of probe and is
Detection of Epstein-Barr Virus 307
determined empirically by running known control tissues and examining stain
intensity. It is recommended that EBER stains be slightly more intense than U6
stains to avoid false-negative EBER results.)
2. Use a pipet to dispense the probe hybridization solution over each tissue section,
using about 20 µL/slide or enough to cover the tissue completely. The pipet tip
can be used to help spread the solution over the section, but avoid scraping the
tissue. Cover slip with parafilm squares that are cut larger than the tissue dimen-sions, and remove any bubbles from under the parafilm using a clean pipet tip. If
using more than one probe on a slide, rubber cement can be applied as a dam to
prevent carryover from one area to another.
3. Lay the slides flat and allow them to hybridize in a covered, barely humid RNase-free chamber at 55°C for 3 h. Longer hybridization times produce stronger sig-nals but will also increase background. Beyond this step, there is no need to use
RNase-free solutions or glassware.
3.2.4. Washing and Detection of Antibody
1. Prewarm the 2X SSC to 37°C in preparation for step 4, and prewarm the 0.1X
SSC + 0.1% SDS to 55°C in preparation for step 5.
2. Remove the parafilm cover slip using a scalpel blade to facilitate lifting but
taking care not to scratch the tissue. To remove excess probe, blot the slides on
clean Kimwipes. Insert the slides into a rack, in which they will remain for the
next several wash steps. Dip the slides in a bath containing 200 mL of 2X SSC +
0.1% SDS.
3. Wash the slides in a bath of 2X SSC + 0.1% SDS for 10 min.
4. To remove excess unbound probe, incubate the slides at 37°C for 5 min in 2X
SSC into which you have freshly mixed 0.3 U/mL of RNase A. (If high back-ground is a problem, the RNase concentration may be doubled.) Caution: All
reagents and plasticware used in this and subsequent steps should be labeled
“RNase-contaminated,” and should not be allowed to come in contact with any
materials used in the prehybridization steps.
5. Dab the slides on Kimwipes, and wash in 0.1X SSC + 0.1% SDS for 5 min at
55°C. (It is in this step that the tissue is most susceptible to falling off the slide;
the likelihood of this occurring is diminished by blotting the slide on dry
Kimwipes prior to this wash.)
6. Rinse in 2X SSC for 1 min.
7. Prepare the anti-DIG antibody solution. This solution represents a 1 500 dilution
of anti-DIG antibody linked to alkaline phosphatase, 1% sheep serum, and 0.3%
Triton X-100 in buffer 1. (To make this solution, combine 2.8 mL of buffer 1,
28 µL of sheep serum, 84  µL of 1 10 Triton X-100, and 5.6  µL of anti-DIG-alkaline phosphatase.)
8. Dab the slides on Kimwipes and place in a horizontal staining tray. Circle each
section using a PAP PEN to form a dam. Apply only enough antibody solution to
each slide to cover the tissue completely, but not to overrun the dam, usually
around 100 µL per section.
308 Fan and Gulley
9. Incubate for 1 h at room temperature in a humid chamber. Add extra antibody
solution as needed to prevent the sections from drying out; dried antibody results
in nonspecific staining.
10. Dab the slides on Kimwipes, and then transfer slides to a staining rack for the
duration of washes.
11. Immediately wash in buffer 1 for 1 min with shaking.
12. Wash in filtered buffer 2 for 1 min with shaking.
13. Immerse the slides into freshly prepared color reaction buffer (180 mL of filtered
buffer 2, 810 µL of NBT solution, 630 µL of BCIP solution), and incubate in the
dark for 90 min or until the positive control is strongly blue.
14. Wash the slides in a bath of buffer 3 for 5 min to stop the reaction.
3.2.5. Counterstaining and Cover Slipping
1. Dip the slides for 1 min in methyl green (1% in H2O), dehydrate by dipping for
2 s each through four baths of 100% ethanol, followed by two dips in xylene. (Do
not use the same ethanol baths that are used for prehybridization because of the
risk of RNase contamination.) Let the slides sit in the xylene while you proceed
with cover slipping the individual slides.
2. Remove each slide from the xylene and place onto absorbent paper towels to
allow excess xylene to run off, but do not let the xylene dry completely. Mount
with Permount under a 50-mm cover slip.
3.2.6. Interpretation of Results
A pathologist should examine the slides microscopically in conjunction with
routine hematoxylin and eosin (H&E) stains to evaluate tissue architecture and
cytomorphology. EBV EBER1 and U6 transcripts are visualized as multiple
and often confluent blue specks localized to the nucleus, except in dividing
cells in which cytoplasmic extension of the signal is seen (see Fig. 1).
Examine the U6 stain to determine whether RNA is well preserved in the
target cells. Then proceed to look for EBER1 signal in these cells. If both are
positive, then the target cells are interpreted as positive for latent EBV infec-tion. (Because U6 contamination of EBER1 slides is feasible, be wary of posi-tivity in inappropriate cell types.) If U6 is negative but EBER1 is positive in a
particular case, use your discretion regarding the appropriate cytomorphology
of the EBER1-positive cells, and consider whether U6 might be below the
threshold for detection.
If EBER1 and U6 stains are both negative in a particular case, repeat with a
shorter (i.e., 2 min) proteinase K incubation, or reduce the RNase concentra-tion and duration of treatment. If both stains are still negative, then the results
are not interpretable owing to insufficient RNA preservation. Lack of adequate
RNA is especially prevalent in salivary gland and in any small biopsy that was
not fixed immediately on collection.
Detection of Epstein-Barr Virus 309
309
Fig. 1. (A) ISH to EBER1 transcripts in a paraffin section of PTLD reveals EBER1 localization to the nucleus of many
lymphoid cells. (B) The presence of U6 transcripts in virtually all cells within the tissue confirms that RNA is preserved and
available for hybridization. (C) The same tissue stained with standard H&E reveals normal squamous epithelium overlying a
dense atypical lymphoid infiltrate.
310 Fan and Gulley
4. Notes
1. To make a solution RNase-free, add 0.1% DEPC, mix well, incubate at 37°C
overnight (or stir vigorously for at least 2 h), and then autoclave for 20 min on
liquid cycle. Be aware that DEPC cannot be relied on to eliminate RNase activity
in solutions containing Tris, because Tris inactivates DEPC. Instead, crystalline
Tris can be dissolved in RNase-free H2O. We have not tested rapid commercial
RNase inactivators.
2. NBT is reconstituted from powder (Boehringer-Mannheim) to a concentration of
75 mg/mL in 70% DMF/water. Make the 70% DMF in a polypropylene tube before
adding the NBT. (Polystyrene tubes and pipets react with DMF.) The resulting solu-tion is stable when stored in the dark at 4°C for 1 mo, or at –20°C for at least 4 mo.
3. BCIP (previously called X-phos) is made from powder (Boehringer-Mannheim).
Bring the BCIP to a concentration of 50 mg/mL in 100% DMF, avoiding polysty-rene tubes and pipets.
4. All glassware and plasticware used in the steps leading to hybridization should
be RNase free. To make them RNase free, bake glassware at 250°C for at least
4 h, or soak glassware and plasticware in 0.1% DEPC for a few minutes, wrap in
aluminum foil, and autoclave for 20 min. Plastic items can be soaked in
0.5 M NaOH for 10 min, rinsed thoroughly with DEPC water, and autoclaved.
Disposable pipets are preferred, and they need not be treated prior to onetime
use. Be aware that some laboratories claim that extensive precautions against
RNA degradation are unnecessary if proper controls for RNA degradation are
included. We recommend a cautious approach in which RNase-free reagents are
used in addition to appropriate controls.
5. The RA386 and RA390 plasmids can be transfected into Escherichia coli and propa-gated in ampicillin-containing broth. Abundant plasmid is then isolated, cut by the
appropriate restriction enzyme to linearize the DNA template, and transcribed to pro-duce riboprobes. To make antisense EBER1 riboprobe, cut the RA386 construct with
HindIII and use T7 polymerase. To make sense EBER1 probe, cut RA386 with EcoRI
and use T3 polymerase. To make antisense U6 probe, cut RA390 with EcoRI and use
T3 polymerase. To make sense U6 probe, cut RA390 with BamHI and use T7 poly-merase. Antisense EBER1 probe recognizes a nonpolyadenylated pol III transcript
characteristic of latent EBV infection that is not translated to protein, whereas sense
EBER1 probe serves as a control of nonspecific hybridization. Antisense U6 probe
recognizes an ubiquitously transcribed human small nuclear pol III transcript that
serves as an indicator of RNA preservation.
6. Cytospins can be prepared from suspensions of mononuclear cells including
blood, body fluids, or cell cultures. Wash cultured cells twice in RNase-free PBS
before suspending about 2  × 106 cells in 400  µL of RNase-free PBS. Add an
equal volume of 10% buffered formalin (Polysciences, Warrington, PA), vortex
gently, and incubate at 4°C for 30 min. Prepare several cytospins on silane-coated
slides, using about 2.5  × 105 cells per slide. Air-dry at room temperature and
hybridize as soon as possible, or store in an RNase-free box at 4°C for up to 2 d
before hybridization.
Detection of Epstein-Barr Virus 311
7. EBV-related Hodgkin disease makes a good positive control because Hodgkin
cells are easily distinguished from uninfected small background lymphocytes on
microscopic examination.
8. Proteinase K digestion facilitates probe penetration into paraffin-embedded tis-sues. Less digestion produces a weaker hybridization signal, whereas over-digestion destroys tissue morphology. Optimal digestion time can be determined
empirically for each tissue.
References
1. Wu, T. C., Mann, R. B., Epstein, J. I., MacMahon, E., Lee, W. A., Charache, P.,
Hayward, S. D., Kurman, R. J., Hayward, G. S., and Ambinder, R. F. (1991) Abun-dant expression of EBER1 small nuclear RNA in nasopharyngeal carcinoma: a
morphologically distinctive target for detection of Epstein-Barr virus in formalin-fixed paraffin-embedded carcinoma specimens. Am. J. Pathol. 138, 1461–1469.
2. Wu, T. C. and Kuo, T. T. (1993) Study of Epstein-Barr virus early RNA-1
(EBER1) expression by in situ hybridization in thymic epithelial tumors of Chi-nese patients in Taiwan. Hum. Pathol. 24, 235–238.
3. Gulley, M. L., Eagan, P. A., Quintanilla-Martinez, L., Picado, A. L., Smir, B. N.,
Childs, C., Dunn, C. D., Craig, F. E., Williams, J. W., and Banks, P. M. (1994)
Epstein-Barr Virus DNA is abundant and monoclonal in the Reed-Sternberg cells
of Hodgkin’s disease: association with mixed cellularity subtype and Hispanic
American ethnicity. Blood 83, 1595–1602.
4. Reynolds, D. J., Banks, P. M., and Gulley, M. L. (1995) New characterization of
infectious mononucleosis and a phenotypic comparison with Hodgkin’s disease.
Am. J. Pathol. 146, 379–388.
5. Ambinder, R. F. and Mann, R. B. (1994) Epstein-Barr encoded RNA in situ
hybridization: diagnostic applications. Hum. Pathol. 25, 602–605.
6. Gilligan, K., Rajadurai, P., Resnick, L., and Raab-Traub, N. (1990) Epstein-Barr
virus small nuclear RNAs are not expressed in permissively infected cells in
AIDS-associated leukoplakia. Proc. Natl. Acad. Sci. USA 87, 8790–8794.
7. Barletta, J. M., Kingma, D. W., Ling, Y., Charache, P., Mann, R. B., and
Ambinder, R. F. (1993) Rapid in situ hybridization for the diagnosis of latent
Epstein-Barr virus infection. Mol. Cell. Probes 7, 105–109.
8. Ambinder, R. F., Browning, P. J., Lorenzana, I., et al. (1993) Epstein-Barr virus
and childhood Hodgkin’s disease in Honduras and the United States. Blood 81,
462–467.
9. Randhawa, P. S., Jaffe, R., Demetris, A. J., Nalesnik, M., Starzl, T. E., Chen,
Y. Y., and Weiss, L. M. (1992) Expression of Epstein-Barr virus-encoded small
RNA in liver specimens from transplant recipients with post-transplantation
lymphoproliferative disease. N. Engl. J. Med. 327, 1710–1714.
10. Montone, K. T., Friedman, H., Hodinka, R. L., Hicks, D. G., Kant, J. A., and
Tomaszewski, J. E. (1992) In situ hybridization for Epstein-Barr virus NotI
repeats in posttransplant lymphoproliferative disorder. Mod. Pathol. 5, 292–302.
Detection of Epstein-Barr Virus 313
23
Molecular Methods for Detecting
Epstein-Barr Virus (Part II)
Structural Analysis of Epstein-Barr Virus DNA
as a Marker of Clonality
Hongxin Fan and Margaret L. Gulley
1. Introduction
The Southern blot technique can be used to determine the clonality of
Epstein-Barr virus (EBV) infected cells (1 ,2) . This clonality assay capitalizes
on measurable polymorphisms in EBV genomic structure, namely, the vari-able number of tandem repeats lying at either end of the linear viral genome.
Thus, the size of genome varies from virion to virion depending on the number
of terminal repeat sequences. When a particular virion infects a cell, its linear
genome circularizes by fusing the terminal repeats to form an episome contain-ing from 1–20 tandem repeat sequences. If an infected cell undergoes malig-nant transformation, the viral DNA replicates along with cell DNA during
mitosis, and the same terminal repeat structure is inherited by all tumor cell
progeny. Therefore, Southern blot analysis of the EBV terminal restriction frag-ment serves as a marker of cellular clonality.
The EBV clonality assay presented herein, which is adapted from a proce-dure first described by Raab-Traub and Flynn (1) , begins by preparing a South-ern blot using BamHI-restricted DNA extracted from patient tissue. The blot is
hybridized to a probe targeting the terminal restriction fragment, such as the
Xho1a probe that recognizes unique DNA adjacent to the terminal repeat
sequences on the right end of the EBV genome. The resultant band pattern
reflects the clonality of the lesion with respect to the structure of EBV DNA
(Fig. 1).
313
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
314 Fan and Gulley
Distinguishing polyclonal from monoclonal lymphoid tumors may have
prognostic and therapeutic implications in immunosuppressed patients. In
addition, latent (episomal) viral DNA may be distinguished from replicative
(linear) viral DNA in this assay; only linear viral DNA is inhibited by acyclovir
or related nucleoside analogs.
2. Materials
2.1. DNA Digestion and Southern Blot
1. BamHI restriction endonuclease.
2. 10X Restriction enzyme BamHI buffer.
3. 10X Gel-loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol, 15%
Ficoll (type 400) in 10X TAE buffer.
4. Agarose.
5. 1X TAE buffer: 40 mM Tris-acetate, 1 mM EDTA.
6. Ethidium bromide (10 mg/mL).
7. 20X Saline sodium citrate (SSC): 3 M NaCl, 0.3 M sodium citrate, pH 7.4.
Fig. 1. Southern blot analysis of EBV DNA structure can be used to determine
whether viral DNA is derived from monoclonal or oligoclonal cells, or whether it is a
product of infectious viral replication. First, DNA is cut with the BamHI restriction
enzyme (arrows), size fractionated by gel electrophoresis, and transferred to a mem-brane by the Southern blot method. The probe most commonly used to detect the EBV
terminal restriction fragment is the 1.9-kb  Xho1a probe (bar), which targets unique
sequences adjacent to the right terminal repeats. Infectious virions have variably sized
terminal restriction fragments resulting in a ladder array of bands on the Southern blot.
In monoclonal tumors, a single fused terminal restriction fragment is seen. In oligo-clonal tumors, several discrete restriction fragments are seen, reflecting the presence
of multiple separate clones.
Detection of Epstein-Barr Virus 315
8. Nylon membrane.
9. Whatman 3MM paper.
10. Paper towels.
2.2. Preparation of RNA Probe from DNA Template
Preparation and use of RNA probes requires that all solutions be RNase free
(see Note 1).
1. HindIII, EcoRI restriction endonuclease.
2. 10X Restriction enzyme HindIII, EcoRI buffer.
3. RNase-free TE buffer: 10 mM Tris-HCl, pH 8.0, 0.1 mM EDTA.
4. 100 mM Dithiothreitol (DTT), RNase free.
5. RNasin® ribonuclease inhibitor (40 U/µL) (Promega, Madison, WI).
6. Bovine serum albumin (BSA), nuclease-free (1 mg/mL).
7. 10 mM NTP stock.
8. 32P-UTP (10 mCi/mL) (3000 Ci/mmol) (New England Nuclear, Boston, MA).
9. SP6 or T7 RNA polymerase (Promega).
10. 5X Transcription buffer (Promega).
11. RNase-free DNase (Promega).
12. G-50 Sephadex (RNA) Quick Spin™ columns (Boehringer Mannheim,
Mannheim, Germany).
2.3. Hybridization
1. Formamide (molecular biology grade); store at 4°C in the dark.
2. Diethylpyrocarbonate (DEPC)-treated H2O.
3. 20X SSC: 3 M NaCl, 0.3 M sodium citrate, pH 7.4.
4. Sonicated salmon sperm DNA (10 mg/mL).
5. 20% sodium dodecyl sulfate (SDS).
6. 1 M PIPES, (piperazine-1,4-diethanesulfonic acid), pH 6.5.
7. 100X Denhardt’s reagent made with DEPC-H2O: 2% Ficoll 400 (type 400),
2% PVP (polyvinylpyrrolidone), 2% BSA (Fraction V).
8. Hybridization solution made from RNase-free stock solutions: 50% formamide,
5X SSC, pH 7.4, 1X Denhardt’s reagent, 0.2% SDS, 100 µg/mL of boiled soni-cated salmon sperm DNA, 50 mM PIPES, pH 6.5.
9. Washing solutions (no. 1, 2, 3, and 4): 0.2% SDS, 0.01% sodium pyrophosphate,
and sequential dilutions of 2X SSC (2, 1, 0.5, and 0.25X SSC).
10. RNase A (Sigma, St. Louis, MO; concentration varies by lot number).
11. RNase buffer: 2X SSC in 20 mM Tris, 5 mM EDTA, pH 8.0.
2.4. Equipment
1. Vacuum oven.
2. Hybridization oven.
3. Centrifuge.
4. Water bath.
5. Scintillation counter.
316 Fan and Gulley
3. Methods
3.1. Preparation of DNA
Biopsy tissue is snap-frozen and stored at –70°C until analysis. Intact
genomic DNA is isolated by SDS-proteinase K lysis and phenol-chloroform
extraction (see Chapter 2).
3.2. DNA Digestion and Southern Blot
1. Restriction endonuclease digestion: Mix 10 µg of genomic DNA, 100 µ of BamHI
restriction enzyme, and 5  µL of 10X  BamHI reaction buffer. Add double-distilled H2O (ddH2O) to a total volume of 50 µL, mix well, and incubate at 37°C
for 4 h to overnight. Add 5 µL of gel-loading buffer to stop the reaction.
2. Load the sample onto a 0.7% agarose gel in 1X TAE buffer containing 0.5 µg/mL
of ethidium bromide. Electrophorese for about 16 h at 1.5 V/cm until the dye
reaches the end of the gel.
3. Denature the DNA and transfer to a nylon membrane using 10X SSC, pH 7.4, by
the Southern blot method.
4. To immobilize the DNA, place the nylon membrane between two sheets of
Whatman 3MM paper and bake under vacuum at 80°C for 1 to 2 h. Keep the
membrane dry until hybridization, or put it in a sealed plastic bag at –20°C for
long-term storage.
3.3. Preparation of RNA Probes from DNA Template
3.3.1. Linearizing Plasmid Vector Containing Template DNA
1. Set up a restriction endonuclease digestion reaction using one of the following
vectors containing template DNA (see Note 2):
a. Xho1a probe in pGEM2 vector: To 20 µg of vector DNA, add 20 µL of 10X
HindIII reaction buffer, 100 U of HindIII restriction endonuclease, and ddH2O
to 200 µL.
b. EcoR1I probe in pGEM2 vector: To 20 µg of vector DNA, add 20 µL of 10X
EcoRI reaction buffer, 100 U of EcoRI restriction endonuclease, and ddH2O
to 200 µL.
2. Incubate the digests at 37°C for 4 h to overnight.
3. Electrophorese a small aliquot of undigested vector/insert alongside the digested
vector/insert in an agarose gel to check for completeness of digestion. Only one
band of the appropriate size should be evident in the linearized template.
4. Purify the linearized DNA by phenol/chloroform extraction and ethanol precipi-tation, and resuspend it in 20  µL of RNase-free TE buffer at a concentration
of 1  µg/µL. This purified linear DNA is used as a template for producing
32P-labeled riboprobes.
3.3.2. Labeling of RNA Probes with 32 P-UTP
Preparation of probes must be carried out in an RNase-free environment.
Gloves should be worn during all steps to avoid RNase contamination (see
Detection of Epstein-Barr Virus 317
Note 1). RNA is transcribed in the presence of radionucleotides to produce a
labeled riboprobe. Transcription initiates in vector sequences containing RNA
polymerase recognition sites (3) .
1. For each 20-µL transcription reaction, mix 4 µL of 5X transcription buffer; 2 µL
of 100 mM DTT; 2 µL of 1 mg/mL BSA; 0.8 µL of 40 U/µL RNasin; 1 µL each
of 10 mM rATP, rGTP, and rCTP; 1.5  µL of freshly made 1 100 dilution of
10 mM rUTP; 5 µL of 10 mCi/mL 32P-UTP; 1 µL of linearized template DNA;
and 1 µL of appropriate RNA polymerase (SP6 for Xho1a, T7 for EcoR1I).
2. Incubate the reaction at 37°C for 30 min, add another 1 µL of the appropriate
RNA polymerase, and incubate at 37°C for another 30 min.
3. Add 1.25  µL of RNasin to stablilize the product, and 1  µL of 1 U/µL DNase
(RNase free) to degrade the template DNA. Incubate the reaction at 37°C for
10 min.
4. Isolate the 32P-labeled RNA probe on a G-50 Sephadex (RNA) Quick Spin col-umn according to the manufacturer’s protocol. Collect the probe in a microfuge
tube for hybridization.
5. Measure the labeling activity of the RNA probe by assaying 1 µL in a scintilla-tion counter. Counts should be between 1–3 million cpm/µL.
3.4. Hybridization
Southern blot hybridization to either the  Xho1a or the  EcoR1I riboprobe
yields visible bands on autoradiographs. Nonspecific hybridization is avoided
by pretreating the membrane with prehybridization solution, and by removing
excessive unbound probe by RNase treatment. DNA from the Raji Burkitt’s
lymphoma cell line (American Type Culture Collection, Rockville, MD) is
recommended as a positive control. Any EBV-negative tissue can serve as a
negative control.
1. Prepare 40 mL of fresh hybridization solution at room temperature; then prewarm
to 45°C.
2. To prehybridize, pour half of the hybridization solution (20 mL) over the mem-brane and incubate at 45°C while shaking for at least 2 h. Then discard the
prehybridization solution.
3. Add about 30 µL of 32P-UTP-labeled Xho1a or EcoR1I RNA probe to 20 mL of
fresh hybridization solution and mix well. Incubate with the membrane at 45°C
while shaking overnight. (Because the probe is single stranded, it need not be
boiled prior to use.)
4. Prepare the four wash solutions (no. 1, 2, 3, and 4) and warm them to 68°C for at
least 1 h or overnight.
5. Wash at 68°C for 30 min each in wash solution no. 1 and wash solution no. 2.
6. Rinse the membrane several times in ddH2O, then once with RNase buffer.
7. Digest the unbound probe by incubating the membrane while shaking it in 20 mL
of prewarmed RNase buffer containing 0.3 U/mL of RNase A at 37°C for exactly
318 Fan and Gulley
5 min. Then rinse the blot several times with ddH2O to remove excess RNase. Be
careful to contain the RNase solution so that it does not contaminate RNase-free
materials in the laboratory.
8. Wash at 68°C for 30 min each in wash solution no. 3 and wash solution no. 4.
9. Place the membrane in an envelope of Whatman 3MM paper or air-dry for at
least 30 min.
10. Set up autoradiography by exposing the blot to X-ray film overnight.
3.5. Interpretation of Results
In this procedure, total DNA was extracted from patient tissues, digested
with BamHI restriction endonuclease, electrophoresed through an agarose gel
to size fractionate the fragments, and transferred to a nylon membrane. The
immobilized DNA fragments were hybridized to a 32P-radiolabeled RNA probe
targeting the terminal restriction fragment of the EBV genome. Unbound probe
was washed away, and the membrane was exposed to X-ray film to detect
hybridized bands. Positive and negative control samples, and a molecular
weight size marker, should be run in adjacent lanes.
Raji control DNA produces a single band of about 23 kb, confirming that
monoclonal EBV DNA is present. In patient samples, the presence of a single
band larger than 6 kb is interpreted as monoclonal EBV DNA, implying the
presence of a monoclonal population of infected cells. Two bands larger than
6 kb represent biclonal EBV DNA, and additional bands over 6 kb represent
oligoclonal EBV DNA (see Fig. 1). A ladder of bands smaller than 6 kb is
indicative of linear EBV DNA, which is the product of viral replication. Viral
replication may coexist in the same tissue as latent infection, potentially yield-ing both small and large bands.
4. Notes
1. To make any solution RNase free, add 0.1% DEPC, mix well, incubate at 37°C
overnight (or stir vigorously for at least 2 h), and then autoclave for 20 min on
liquid cycle. Be aware that DEPC cannot be relied on to eliminate RNase activity
in solutions containing Tris, because Tris inactivates DEPC. Instead, Tris solu-tions are made using RNase-free H2O. We have not tested commercial RNase
inactivators. Glassware and plasticware used in all steps through hybridization
should be RNase free. To make them RNase free, bake glassware at 250°C for at
least 4 h, or soak glassware and plasticware in 0.1% DEPC for a few minutes,
wrap in aluminum foil, and autoclave for 20 min. Plastic items can also be soaked
in 0.5 M NaOH for 10 min, rinsed thoroughly with water, and autoclaved. Dis-posable pipets are preferred, and they need not be treated prior to onetime use.
2. The probes most commonly used to detect the terminal restriction fragment of
EBV DNA are the 1.9-kb Xho1a probe or the 4.0-kb BamH1 subfragment of the
EcoR1I fragment (1 ,4) . The templates for making these probes are available in
Detection of Epstein-Barr Virus 319
plasmid vectors from Nancy Raab-Traub, PhD, University of North Carolina at
Chapel Hill. The plasmid can be transfected into Escherichia coli and propagated
in ampicillin-containing LB broth. Then abundant plasmid can be isolated and
cut with an appropriate restriction enzyme to linearize the DNA immediately
downstream of the cloned insert. The insert can then serve as a template for RNA
transcription in the presence of radionucleotides to produce “riboprobes” labeled
with 32P. Instead of making RNA probe as described in this procedure, an alter-native approach is to make a labeled DNA probe from the same plasmid insert
previously mentioned, or to use another probe such as the 5.2-kb BamHI/EcoRI
fused terminal repeat probe labeled by random priming (5) .
References
1. Raab-Traub, N. and Flynn, K. (1986) The structure of the termini of the Epstein-Barr virus as a marker of clonal cellular proliferation. Cell 47, 883–889.
2. Gulley, M. L. and Raab-Traub, N. (1993) Detection of Epstein-Barr virus in
human tissues by molecular genetic techniques.  Arch. Pathol. Lab. Med. 117,
1115–1120 (review).
3. McCracken, S. (1985) Preparation of RNA transcripts using SP6 RNA poly-merase. Focus (Bethesda Research Laboratories) 7, 5.
4. Gulley, M. L., Raphael, M., Lutz, C. T., Ross, D. W., and Raab-Traub, N. (1992)
Epstein-Barr virus integration in human lymphomas and lymphoid cell lines.
Cancer 70, 185–191.
5. Weiss, L. M., Strickler, J. G., Warnke, R. A., Purtilo, D. T., and Sklar, J. (1987)
Epstein-Barr viral DNA in tissues of Hodgkin’s disease.  Am. J. Pathol. 129,
86–91.
Detection of Epstein-Barr Virus 321
24
Molecular Methods for Detecting
Epstein-Barr Virus (Part III)
EBV Viral Load by Competitive Polymerase Chain Reaction
Hongxin Fan and Margaret L. Gulley
1. Introduction
Epstein-Barr virus (EBV) viral load testing is rapidly gaining acceptance in
the diagnosis and monitoring of patients with EBV-related neoplasia, includ-ing transplant recipients, autoimmune deficiency syndrome (AIDS) patients
with brain lymphoma, and patients with nasopharyngeal carcinoma. In trans-plant recipients, several studies have shown that EBV viral load in blood is a
useful marker of EBV-driven posttransplant lymphoproliferative disease
(PTLD), both in terms of predicting disease and in monitoring efficacy of
therapy (1–4) . In AIDS patients, high EBV viral load in cerebrospinal fluid
(CSF) is so characteristic of brain lymphoma that the assay has been touted as
sufficient for making that diagnosis without the need for brain biopsy, assum-ing that there is also clinical and radiographic support for the diagnosis (5) . In
nasopharyngeal carcinoma patients, a recent study showed that plasma
EBV viral load is nearly always elevated, and the degree of elevation is higher
in those with distant metastases  (6) . Additional studies are needed to more
completely define the utility of EBV viral load measurement in monitoring
residual disease following therapy, and to evaluate clinical utility in other EBV-associated diseases.
EBV viral load can be measured by several methods including quantitative
culture, counting the proportion of EBV-encoded RNA (EBER)-stained cells
in tissue specimens, and quantitative polymerase chain reaction (PCR) (7–10) .
Quantitative PCR is recommended because it is sensitive, specific, quantita-321
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
322 Fan and Gulley
tive across a wide dynamic range, and semiautomated to reduce labor costs.
Furthermore, several studies have suggested that blood or CSF is amenable for
testing, thus avoiding invasive biopsy procedures.
Several protocols have been described for quantitative PCR, including real-time measurement of PCR products in a thermocycler (6 ,11 ,12) . or post-PCR
quantitation of products using commercial detection kits (DiaSorin, Stillwater,
MN; Digene Diagnostics, Silver Spring, MD; BioSource International,
Camarillo, CA). The EBV viral load protocol presented here is a competitive
PCR assay modified from a commercial kit (Viral Quant™ EBV kit, BioSource
International). This assay relies on competition between EBV genomic
sequences in the patient sample and an internal calibration standard (ICS) that
is spiked into the sample prior to extraction. Following coamplification using
biotinylated primers, PCR products are detected in an automated enzyme-linked immunosorbent assay (ELISA) plate system using a 96-well plate
washer and reader. The spiked standard contains the same primer recognition
sequences as the EBV genomic target (the EBER gene), but has different
interim sequences that are distinguished by specific internal probes coating the
inside of the microwells. Comparison between the amount of EBV product and
the amount of internal standard product permits calculation of EBV viral load
in the patient specimen.
In our modified procedure, DNA is extracted using a commercial kit
(QIAmp Viral RNA kit, Qiagen, Valencia, CA), and dUTP replaces dTTP in
the PCR reaction so that uracil N-glycosylase (UNG) enzyme can be used to
diminish the risk of amplicon contamination  (13) . Sample types include
plasma, serum, blood mononuclear cells, or CSF.
2. Materials
2.1. Reagents
1. QIAamp Viral RNA kit (Qiagen) (see Note 1).
2. AE buffer (Qiagen).
3. 100% Ethanol.
4. GeneAmp dNTPs with dUTP (PE Applied Biosystems, Foster City, CA).
5. Taq DNA polymerase (Gibco-BRL, Gaithersburg, MD).
6. Uracil-DNA glycosylase, heat labile (heat labile UNG; Boehringer Mannheim,
Mannheim, Germany).
7. Viral Quant EBV kit (BioSource International).
8. 1X TAE buffer: 40 mM Tris-acetate, 1 mM EDTA.
9. Agarose.
10. Ethidium bromide (10 mg/mL).
11. 10X Gel-loading buffer: 0.25% bromophenol blue, 0.25% xylene cyanol, 15%
Ficoll (type 400) in 10X TAE buffer.
12. Nuclease-free H2O.
Detection of Epstein-Barr Virus 323
2.2. Equipment
1. Centrifuge.
2. Microcentrifuge.
3. Thermocycler (GeneAmp PCR system 9700; PE Applied Biosystems).
4. ELP-40 Microplate Strip Washer (Bio-Tek, Winooski, VT).
5. Elx 800 Automated Microplate Reader (Bio-Tek).
3. Methods
3.1. General PCR Precautions
Precautions are necessary to prevent contamination of PCR reactions. Clean
work areas, physical separation of pre- and postamplification laboratories, and
appropriate controls are essential. Most important is meticulous care by well-trained personnel. UNG is used to further ensure that amplicons from previous
reactions are not amplified.
3.2. Preparation of Samples
Plasma, serum, and blood mononuclear cells are all potential sources of
virus in transplant recipients. Plasma and serum are recommended in nasopha-ryngeal carcinoma patients, whereas blood mononuclear cells have not yet been
studied. CSF is the preferred sample type in AIDS patients with suspected brain
lymphoma. For comparability purposes, serial samples from patients followed
over time should always be analyzed using the same sample type and labora-tory procedure.
1. Plasma: Collect whole blood in EDTA anticoagulant. Centrifuge at 1800g for 10
min. Recover the plasma, aliquot, and freeze at –20°C.
2. Serum: Recover the serum from the whole blood following centrifugation at
1800g for 10 min. Aliquot and store at –20°C.
3. CSF: Collect 1 mL of fresh specimen, aliquot, and store at –20°C. (If the CSF is
reddish or cloudy, indicating high probability of red cells or leukocytes, centri-fuge it and recover the supernatant for testing.)
4. Peripheral blood mononuclear cells: Isolate the DNA using your standard proto-col, such as the Puregene DNA isolation kit (Gentra Systems, Minneapolis, MN).
Quantitate the DNA, and then proceed directly to to PCR, or store the DNA at
–20°C.
3.3. Isolation of Viral DNA
Viral DNA is isolated from plasma, serum, or CSF using the QIAamp Viral
RNA kit. Although the name of this kit implies that its purpose is to isolate
RNA, we found it effective in isolating EBV DNA from patient samples. The
kit is supplied with carrier RNA to improve sample recovery. (Alternatively,
the QIAmp Blood kit can be used as described in Note 1. )
324 Fan and Gulley
1. Pipet 140 µL of plasma, serum, or CSF into a 1.5-mL microfuge tube.
2. Add 5000 copies of ICS (supplied in the Viral Quant EBV kit) to each tube,
including the positive and negative controls.
3. Follow the manufacturer’s instructions provided in the QIAmp viral RNA kit,
and at the last step elute the DNA into 50 µL of AE buffer or nuclease-free H2O.
Of this, only 5 µL is typically used in subsequent PCR reactions.
3.4. Polymerase Chain Reaction
PCR is performed using primers targeting the EBV EBER gene. One of
these primers is biotinylated, and both are provided in the Viral Quant EBV kit
(5v-CCCGCCTACACACCAACTAT-3v; 5v-AGTCTGGGAAGACAACCA-CA-3v). The manufacturer’s procedure has been modified to accommodate the
UNG system of hindering amplicon contamination (see Note 2).
1. Prepare the PCR master mix in a 1.5-mL microfuge tube on ice, and aliquot
95 µL into each 0.2-mL thin-walled tube for PCR. Per reaction, the master mix
should include 69.5 µL of nuclease-free H2O, 10 µL of 10X PCR buffer, 2 µL of
25 pmol/µL primer set, 1  µL of heat-labile UNG, 8  µL of dNTPs with dUTP
(2.5 mM of each dATP, dCTP, and dGTP; 5.0 mM of dUTP), 4 µL of 50 mM
MgCl2, and 0.5 µL of 5 U/µL of Taq polymerase (see Note 3).
2. For DNA isolated from plasma, serum, or CSF, add 5 µL of template DNA (which
contains 500 copies of spiked ICS) to the aliquot of master mix. For DNA
extracted from blood mononuclear cells, add 1 µg of template DNA to the aliquot
of master mix, and since ICS has not already been spiked into this sample, add
500 copies of ICS directly to the PCR mixture, adjusting the total PCR volume to
100 µL.
3. Controls: Use the plasmid construct containing EBER1 sequences and the carrier
DNA in the Viral Quant EBV kit to serve as a positive and a negative control,
respectively. In addition, it is wise to include high and low patient controls in
each run, and additional negative controls for every 10 patient samples.
4. Run the thermocycler program as follows: 20°C for 10 min (to allow heat-labile
UNG to act); 95°C for 3 min (to inactivate heat-labile UNG); and then 34 PCR
cycles consisting of denaturation at 94°C for 30 s, annealing at 60°C for 30 s, and
extension at 72°C for 1 min. Run the final extension step at 72°C for 15 min, and
then hold briefly at 72°C until the PCR tubes are removed for immediate detec-tion or storage at –20°C (see Note 4). According to the Viral Quant EBV kit
protocol, these thermocycler conditions allow detection of about 25 copies per
amplification reaction. To detect lower copy numbers, see Note 5.
3.5. Confirmation of PCR Products
by Agarose Gel Electrophoresis
3.5.1. Agarose Gel Electrophoresis
Viewing the size and relative proportion of ICS and EBV products is useful
for quality assurance purposes and for troubleshooting. It is also used to con-
Detection of Epstein-Barr Virus 325
firm PCR productivity prior to proceeding with the relatively expensive ELISA
plate detection steps.
1. Prepare a 2% agarose gel in 1X TAE buffer containing 0.5 µg/mL of ethidium
bromide.
2. Mix 25 µL of the PCR product with 3 µL of 10X gel-loading buffer and load into
the wells. Load one lane with a molecular weight marker.
3. Electrophorese at 5 V/cm until the dye has migrated about three fourths the length
of the gel.
4. Photograph the gel under a UV transilluminator.
3.5.2. Interpretation of Gel Electrophoresis
Amplified ICS produces a 260-bp product, whereas amplified EBV produces
a 210-bp product. Bands at 210 and 260 bp indicate coamplification of EBV
and ICS. The relative amount of EBV and ICS in each sample reflects the
proportion of starting templates in the PCR reaction.
ICS at 260 bp should be visible in all patient samples. If only the ICS band is
seen, then EBV DNA is below the threshold for visualization, and generally
will also be near or below the threshold for reporting following the ELISA
plate detection steps. If the ICS band is absent but a strong EBV 210-bp band is
seen, this probably indicates that abundant EBV DNA is present and has over-whelmed the ICS amplification. In this circumstance, dilute an aliquot of the
original patient sample 1 10 with H2O, then spike with ICS, extract the DNA,
and amplify again. Be sure to document the dilution so that the final results
will not underestimate the amount of EBV.
If no bands are detected in the patient lane but the controls are as expected,
a PCR inhibitor may be the culprit. If this is the case, consider repeating the
PCR reaction using less template DNA, and document this so that calculations
can be adjusted accordingly. Alternatively, further purify the original template
DNA by phenol/chloroform extraction and ethanol precipitation before repeat-ing the assay.
3.6. ELISA Plate Detection System
3.6.1. Microplate Detection
Microtiter plates precoated with ICS or EBV capture oligonucleotide probes
are provided in the Viral Quant EBV kit. All detection reagents are prepared
according to the manufacturer’s instructions.
1. If frozen, thaw the PCR product immediately before use. Make serial dilutions
and follow the Viral Quant EBV kit instructions for detection of products (see
Note 6).
2. Proceed with the detection procedure and wash steps according to the Viral Quant
EBV kit instructions. Measure the OD450 on an ELISA plate reader.
326 Fan and Gulley
3.6.2. Calculation of Results
For each sample, calculate the EBV viral load as instructed in the Viral
Quant EBV kit protocol (see Note 7). Remember to adjust the copy number by
any dilution factor used prior to amplification (see Note 8). Assay results
are objective; however, medical use of these results requires clinicopathologic
correlation.
3.6.3. Clinicopathologic Interpretation of Results
The “normal” range for this assay depends on the clinical status. Each labo-ratory is encouraged to validate the assay on its own patient populations.
Healthy nonimmunosuppressed patients generally have undetectable viral
loads, i.e., less than about 1000 copies/mL of plasma or serum. For clinical
evaluation of immunosuppressed transplant recipients, serial samples are
encouraged rather than a onetime test. In serial samples analyzed over time,
10-fold differences in viral load are considered to be clinically significant.
Baseline values are encouraged in healthy transplant recipients so that future
values may be judged in comparison. Baseline values may be higher in patients
who were seronegative at the time of transplant, compared with seropositive
recipients who harbor prior immunity. In general, patients with EBV-driven
PTLD have plasma or serum values higher than 5000 copies/mL. Undetectable
EBV viral loads are consistent with the absence of EBV-driven PTLD. Values
in the gray zone, as well as onetime values at any level, should be interpreted in
the context of clinicopathologic findings.
Although few studies of nasopharyngeal carcinoma have been conducted,
our own experience suggests that these patients have relatively low EBV viral
loads until the disease becomes widely metastatic.
4. Notes
1. An alternate kit for isolating DNA from these samples is the QIAamp Blood kit
(Qiagen). It has the advantage of slightly higher sensitivity since it samples 200
rather than 140  µL of patient specimen; however, the kit is not supplied with
carrier DNA. Therefore, carrier nucleic acid, such as Poly-deoxy-adenylic-thymidylic acid (Boehringer Mannheim), must be added to the AE buffer at a
final concentration of 20 µg/mL. The amount of spiked ICS remains the same at
5000 copies/sample, and target DNA is eluted off the column into 50 µL of AE
buffer. Calculation of viral load must be adjusted to accommodate the higher
initial sample volume.
2. We recommend heat-labile rather than conventional UNG. The problem with
conventional UNG is that it may not be completely inactivated by heating for
10 min at 95°C, and therefore it may degrade newly formed PCR products (14) .
By contrast, heat-labile UNG is more rapidly and efficiently inactivated (15) , and
we have confirmed that it works better in this protocol. Be aware that some
Detection of Epstein-Barr Virus 327
laboratories use an alternate strategy: UNG is not used on a routine basis, but
dUTP is routinely incorporated into PCR products so that UNG will be effective
should a contamination problem occur.
3. Although the PCR conditions have been carefully optimized in this protocol, it
might be necessary to further optimize this protocol in your own laboratory’s
thermocycler, considering the dUTP and magnesium concentrations, and verify-ing that UNG activity is ablated during the 95°C preincubation.
4. Following amplification, immediately proceed to the detection procedure or
freeze the products at –20°C. Attention to timeliness will minimize the possibil-ity of residual UNG activity destroying the PCR products (14) .
5. Lower EBV copy numbers can be detected by using 40 rather than 34 cycles;
however, if 40 cycles are used, begin by spiking with 100 rather than 500 copies
of ICS per reaction. For routine clinical use in transplant recipients, we have
found 34 cycles to be adequate.
6. The manufacturer’s instructions recommend making two- or fivefold dilutions of
the PCR products prior to detection. These dilution factors can be adjusted
according to your experience with the patient samples in your own facility. In our
laboratory, we found that fourfold dilutions produced more definitive values.  To
make fourfold serial dilutions in wells B–D, add 100 µL of hybridization buffer
to wells B, C, and D of both the ICS and EBV strips. Add 133.3  µL of the
suggested starting dilution (1 20) of EBV or ICS to well A. Transfer 33.3 µL
from row A to B and mix up and down six times by pipeting with plugged tips.
Continue this process through row D. After mixing row D, remove and discard
33.3 µL.
7. The manufacturer’s instructions state that an OD450 of 0.3–1.5 is the acceptable
range for proceeding, but we often visualize an electrophoretic EBV band in
samples with an OD450 as low as 0.1. Therefore, we recommend that if a weak
EBV band is visualized on the gel, and the OD450 is between 0.1 and 0.3, you
may still calculate a result.
8. To clarify the adjustments that must be made to accommodate dilution factors,
remember that the starting volume of patient plasma, serum, or CSF is typically
140 µL (QIAamp Viral kit) or 200 µL (QIAamp Blood kit). This gets eluted into
50 µL of diluent following column extraction. Then 5 µL of that is used in PCR.
Therefore, to report EBV copies/mL of patient specimen, the raw result should
be adjusted as follows. For 140-µL samples, multiply the raw result by 71.4. For
200-µL patient samples, multiply the raw result by 50. For DNA extracted from
blood mononuclear cells and later spiked with ICS, results are reported in copies/
µg of DNA, so raw results need not be adjusted, as 1 µg of template DNA was
used in the PCR reaction. Finally, round off all results to the first two digits (e.g.,
5397 becomes 5400; 236,000 becomes 240,000).
References
1. Savoie, A., Perpete, C., Carpentier, L., Joncas, J., and Alfieri, C. (1994) Direct
correlation between the load of Epstein-Barr virus-infected lymphocytes in the
328 Fan and Gulley
peripheral blood of pediatric transplant patients and risk of lymphoproliferative
disease. Blood 83, 2715–2722.
2. Rooney, C. M., Loftin, S. K., Holladay, M. S., Brenner, M. K., Krance, R. A., and
Heslop, H. E. (1995) Early identification of Epstein-Barr virus-associated post-transplantation lymphoproliferative disease. Br. J. Haematol. 89, 98–103.
3. Rowe, D. T., Qu, L., Reyes, J., Jabbour, N., Yunis, E., Putnam, P., Todo, S., and
Green, M. (1997) Use of quantitative competitive PCR to measure Epstein-Barr
virus genome load in the peripheral blood of pediatric transplant patients with
lymphoproliferative disease. J. Clin. Microbiol. 35, 1612–1615.
4. Riddler, S. A., Breinig, M. C., and McKnight, J. L. (1994) Increased levels of
circulating Epstein-Barr virus (EBV)-infected lymphocytes and decreased EBV
nuclear antigen antibody responses are associated with the development of
posttransplant lymphoproliferative disease in solid-organ transplant recipients.
Blood 84, 972–984.
5. Cingolani, A., De Luca, A., Larocca, L. M., Ammassari, A., Scerrati, M., Antinori,
A., and Ortona, L. (1998) Minimally invasive diagnosis of acquired immuno-deficiency syndrome-related primary central nervous system lymphoma. J. Natl.
Cancer Inst. 90, 364–369.
6. Lo, Y. M., Chan, L. Y., Lo, K. W., Leung, S. F., Zhang, J., Chan, A. T., Lee, J. C.,
Hjelm, N. M., Johnson, P. J., and Huang, D. P. (1999) Quantitative analysis of
cell-free Epstein-Barr virus DNA in plasma of patients with nasopharyngeal car-cinoma. Cancer Res. 59, 1188–1191.
7. Randhawa, P. S., Jaffe, R., Demetris, A. J., Nalesnik, M., Starzl, T. E., Chen,
Y. Y., and Weiss, L. M. (1992) Expression of Epstein-Barr virus-encoded small
RNA (by the EBER-1 gene) in liver specimens from transplant recipients with
post-transplantation lymphoproliferative disease.  N. Engl. J. Med. 327,
1710–1714.
8. Stevens, S. J., Vervoort, M. B., van den Brule, A. J., Meenhorst, P. L., Meijer,
C. J., and Middeldorp, J. M. (1999) Monitoring of Epstein-Barr virus DNA load
in peripheral blood by quantitaive competitive PCR.  J. Clin. Microbiol. 37,
2852–2857.
9. Laroche, C., Drouet, E. B., Brousset, P., Pain, C., Boibieux, A., Biron, F., Icart, J.,
Denoyel, G. A., and Niveleau, A. (1995) Measurement by the polymerase chain
reaction of the Epstein-Barr virus load in infectious mononucleosis and AIDS-related non-Hodgkin’s lymphomas. J. Med. Virol. 46, 66–74.
10. Bai, X., Hosler, G., Rogers, B. B., Dawson, D. B., and Scheuermann, R. H. (1997)
Quantitative polymerase chain reaction for human herpesvirus diagnosis and mea-surement of Epstein-Barr virus burden in posttransplant lymphoproliferative dis-order. Clin. Chem. 43, 1843–1849.
11. Swinnen, L. J., Gulley, M. L., Hamilton, E., and Schichman, S. A. (1998) EBV
DNA quantitiation in serum is highly correlated with the development and regres-sion of post-transplant lymphoproliferative disorder (PTLD) in solid organ trans-plant recipients. Blood 92(Suppl. 1), 314a,315a.
Detection of Epstein-Barr Virus 329
12. Kimura, H., Morita, M., Yabuta, Y., Kuzushima, K., Kato, K., Kojima, S.,
Matsuyama, T., and Morishima, T. (1999) Quantitative analysis of Epstein-Barr
virus load by using a real-time PCR assay. J. Clin. Microbiol. 37, 132–136.
13. Longo, M. C., Berninger, M. S., and Hartley, J. L. (1990) Use of uracil DNA
glycosylase to control carry-over contamination in polymerase chain reactions.
Gene 93, 125–128.
14. Thornton, C. G., Hartley, J. L., and Rashtchian, A. (1992) Utilizing uracil DNA
glycosylase to control carryover contamination in PCR: characterization of
residual UDG activity following thermal cycling. Biotechniques 13, 180–184.
15. Sobek, H., Schmidt, M., Frey, B., and Kaluza, K. (1996) Heat-labile uracil-DNA
glycosylase: purification and characterization. FEBS Lett. 388, 1–4.
Molecular Detection of KSHV/HHV-8 331
25
Molecular Detection of Kaposi’s
Sarcoma–Associated Herpesvirus/
Human Herpesvirus-8
Ethel Cesarman
1. Introduction
Kaposi’s sarcoma–associated herpesvirus (KSHV), also called human her-pesvirus-8 (HHV-8), is the most recently identified human herpesvirus (1) . It
has been found to be invariably present in Kaposi’s sarcoma (KS) lesions,
whether these are associated with AIDS (epidemic KS), therapeutic immuno-suppression (iatrogenic KS), or high-incidence regions in Africa (endemic KS),
or in its “classic” form (sporadic KS) (for reviews see refs. 2 and 3 ). By con-trast, with few reported exceptions, it has not been found to be present in a
variety of other vascular tumors and reactive conditions. A seroepidemiologic
association of this virus and KS has been well documented, and it is currently
accepted that KSHV plays a necessary, although not sufficient, role in the
development of KS. Although diagnosis of KS is usually not difficult based on
clinical and histologic features, some cases may have unusual morphology,
with features overlapping those of other vascular and spindle cells prolifera-tions. In these instances, molecular detection is useful to confirm or rule out a
diagnosis of KS.
This virus has also been found to be present in a subset of malignant lym-phomas called primary effusion lymphomas (PELs) or body cavity–based
lymphomas (4). These lymphomas have an unusual set of features that in con-junction with the presence of KSHV suggest that they represent a distinct dis-ease entity  (5 ,6) . Among these features are a morphology bridging that of
anaplastic large-cell lymphoma and immunoblastic lymphoma, and the fre-quent lack of expression of B-cell-associated antigens in spite of a B-cell geno-331
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
332 Cesarman
type. However, other types of lymphomas can involve body cavities as malig-nant effusions, either primarily or secondarily, and do not contain this virus.
This is particularly frequent in the case of Burkitt lymphomas. Furthermore,
PELs can have an “extracavitary” presentation. In these cases, the detection of
KSHV may be important for their subclassification.
A third disease in which the presence of KSHV has been convincingly dem-onstrated is multicentric Castleman’s disease (MCD). When occurring in asso-ciation with AIDS, KSHV is present in the vast majority of cases, whereas it is
seen in close to 50% of cases of MCD in human immunodeficiency virus
(HIV)-negative individuals. Thus far no studies have suggested that the KSHV-positive cases of MCD are different from the negative ones in terms of disease
behavior, outcome, or potential treatment. However, it is possible that as more
information is gathered, assessment of the status of KSHV in MCD may be
useful clinically.
The easiest and by far the most common method used so far for the detection
of KSHV is polymerase chain reaction (PCR). However, extreme care must be
taken, as always with this method, to avoid contamination. Many published
studies have suggested that this virus is present in a variety of conditions or in
the general population, creating extensive controversy in the literature, and
mostly being unconfirmed. Most of these studies have used techniques that
increase the sensitivity of detection, such as nested PCR. In our hands a single-step PCR, followed by hybridization with a radiolabeled internal oligonucle-otide, detects between 1 and 10 copies of the virus in 100 ng of DNA, and has
been found to be sensitive and specific (see Note 1). Using this method, we can
detect every case of KS and PEL, 50% of cases of MCD, close to 30% of
tissues from HIV-positive patients, and <10% of non-KS tissues. However, to
exclude the possibility of contamination, we strongly recommend performing
PCR with at least three independent sets of primers. A wide variety of primer
sets have been published. In this chapter, we provide the sequences and proto-col for three sets with which we have the most extensive experience in our
laboratory. One of these sets, called KS330233, is the first reported method for
detection of KSHV, and the most extensively used (1) . The other two sets were
developed in our laboratory, and we have found the sequences they recognized
to be well conserved among the various KSHV isolates we have examined and
those reported in the literature.
A second method, which we believe to be important in the diagnosis of PEL,
is Southern blot analysis on genomic DNA. Because these lymphomas occur
most frequently in HIV-positive patients, detection by PCR may mean sys-temic infection with KSHV, rather than a specific association with this lym-phoma. Furthermore, PELs contain 40–80 copies of the viral genome per cell,
making its detection using this method quite easy. A full protocol for Southern
Molecular Detection of KSHV/HHV-8 333
blotting is not provided, because any standard method can be used, but the
details specific for KSHV detection are given in Note 2. If Southern blot analy-sis is not possible, approximate quantitation to document high viral copy num-bers is necessary before a diagnosis of PEL is made. This can be performed by
serially diluting the DNA and comparing to known standards.
2. Materials
2.1. Oligonucleotide Primers and Probes
Primers and internal oligonucleotide probes for open reading frame 26
(ORF 26) (Capsid protein), K9 (vIRF1), and ORF 74 (vGPCR). Sequences are
provided in Table 1. Any primer set can be used as a control for integrity of
DNA and absence of inhibitors. One of these control primers sets, shown in
Table 1, is for the `-actin gene.
2.2. Controls
As a positive control, any PEL cell line can be used. One of these is BC-3,
which contains KSHV but lacks EBV. This cell line can be obtained from the
American Type Culture Collection (ATCC) (Rockville, MD; ATCC designa-tion CRL-2277). This cell line contains approx 30 copies of the viral genome/
cell, so it can be diluted 100- to 1000-fold to be used as a control for PCR.
Alternatively, DNA extracted from a KS lesion can be used as a positive
control. DNA from a KS cell line should not be used, because these “lose” the
Table 1
Oligonucleotides for PCR detection of KSHV
Size of
Region amplicon (bp) Primer Primer
ORF 26 233 Forward 5v-AGCCGAAAGGATTCCACCAT-3v
(KS330233) Reverse 5v-TCCGTGTTGTCTACGTCCAG-3v
Probe 5v-TGCAGCAGCTGTTGGTGTACCACATC-3v
K9 184 Forward 5v-CCCTTTCGCGGATATACACA-3v
(vIRF1) Reverse 5v-AGTGAGGGGAAAGCGTCAAT-3v
Probe 5v-GTCTCTGCGCCATTCAAAAC-3v
ORF 74 492 Forward 5v-CCGTGGTGCCTTACACGTGG-3v
(vGPCR) Reverse 5v-CAGTCTGCAGTCATGTTTCC-3v
Probe 5v-TGTGTGCGTCAGTCTAGTGAG-3v
`-Actin 540 Forward 5v-GTGGGGCGCCCCAGGCACCA-3v
Reverse 5v-CTCCTTAATGTCACGCACGATTTC-3v
334 Cesarman
KSHV genome after a few passages in culture. As a negative control, DNA
from peripheral blood mononuclear cells of healthy individuals, or a KSHV-negative cell line (any lymphoblastoid or myeloid cell line), can be used.
2.2. Polymerase Chain Reaction
1. 10X PCR buffer containing 15 mM MgCl2 (Perkin-Elmer, Norwalk, CT).
2. Taq DNA polymerase (5 U/µL) (Perkin-Elmer).
3. Primers (10 pmol/µL).
4. dNTP mixture of 10 pmol/µL of each nucleotide (Boehringer Mannheim, Indian-apolis, IN).
2.3. Analysis of PCR Products
1. 2% Agarose gel in 1X TAE buffer: 40 mM Tris-acetate, 1 mM EDTA, 0.0001%
ethidium bromide.
2.4 Transfer and Hybridization
1. Oligonucleotide probe at 10 pmol/µL (Table 1).
2. [a-32P]ATP (3000 Ci/mmol; NEN).
3. T4 polynucleotide kinase (10 U/µL) (Boehringer Mannheim).
4. 10X T4 polynucleotide kinase buffer (Boehringer Mannheim).
5. Sephadex G25 spin column (Amersham-Pharmacia).
6. Salmon sperm DNA (10 mg/mL) (Boehringer Mannheim).
7. 20X SSPE: 3 M NaCl, 0.2 M NaH2PO4, 0.5 M EDTA, pH 7.4.
8. 20% Sodium dodecyl sulfate (SDS).
9. 50X Denhardt’s solution: 1% Ficoll 400 (Sigma, St. Louis, MO), 1% poly-vinylpyrrolidone (Sigma), and 1% bovine serum albumin, (Pentax fraction V;
Sigma).
10. X-ray film (Kodak).
3. Methods
3.1. Isolation of Nucleic Acid
PCR can be performed on DNA extracted from fresh, frozen, or paraffin-embedded tissue using standard methodology. It is important to extract the
DNA in a physical area that is separate from that used for amplification and
manipulation of PCR products.
3.2. Polymerase Chain Reaction
1. Prepare a master mix in a plastic microfuge tube. This mix should be prepared in
a “clean” area that is separate from those where the DNA is extracted and where
the amplification is performed and PCR products are analyzed. Calculate the
amount of solution to be used by multiplying by the number of samples (n) to be
Molecular Detection of KSHV/HHV-8 335
analyzed plus one (n + 1). Per sample add and mix the following: 14.05 mL of
H2O, 1.25 µL of 2mM dNTP, 2.5 µL of 10X buffer, 1 µL of each forward and
reverse primer at a concentration of 10 pmol/µL, and 0.2 µL of Taq polymerase.
2. Aliquot 20 µL of master mix in each sample tube.
3. Add 100 ng of DNA previously diluted in a total volume of 5 µL.
4. Perform PCR on a thermocycler starting with a denaturation step at 94°C for
3.5 min, followed by 30 cycles of denaturation (1 min at 94°C), annealing (1 min
at 58°C), and extension (1.5 min at 72°C).
3.3. Analysis of PCR Products
1. Withdraw 10 µL from each sample tube. Add 1 µL of 10X loading dye contain-ing bromophenol and xylene-cyanol blue, and mix.
2. Load onto a 2% agarose gel and electrophorese for approx 2 h at 80 V.
3. Visualize in a UV transilluminator and photograph the gel.
3.4. Transfer and Hybridization
1. Transfer of PCR products onto a nitrocellulose filter or nylon membrane should be
performed according to standard procedures as first described by Southern (7) .
2. Prehybridize the filters for 2 h at 37°C in a hybridization solution containing 5X
SSPE, 5X Denhardt’s solution, and 0.5% SDS.
3. Label the internal oligonucleotide probe as follows: In a plastic tube mix 2.5 µL
of probe (25 pmol), 5 µL of [a- 32P]ATP, 1 µL of 10X kinase buffer, and 1 µL of
T4 polynucleotide kinase. Incubate for 90 min at 37°C. Stop the reaction by add-ing 90 µL of TE (10 mM Tris-HCl/1 mM EDTA) and 1 µL 0.5 M EDTA. Remove
free label by centrifugation in a G25 spin column.
4. Hybridize the filters for 5 h to overnight at 37°C in hybridization solution
containing 100 µg/mL of salmon sperm DNA and radiolabeled probe at approx
2 × 106 cpm/mL.
5. Wash the filters twice for 15 min at room temperature in 2X SSPE and 1% SDS,
followed by washing for 10 min at 60°C (KS330233), 55°C (vIRF1), or 57°C
(vGPCR), using a preheated solution of 5X SSPE, and 0.1% SDS. Rinse the fil-ters in 2X saline sodium citrate.
6. Expose on an autoradiograph film.
3.5. Interpretation of Results
Cases should be considered positive only when at least two sets of primers
detect a clearly positive signal. In negative cases, it is important to make sure
that the DNA is amplifiable using a control set of primers.
4. Notes
1. In our hands, the PCR products can be visualized without the need for hybridiza-tion, even when very low copy numbers (10–100) are present. With the vIRF1
primers we can detect 1–5 copies without the need for hybridization. However,
hybridization is useful to confirm the specificity of the amplified products.
336 Cesarman
2. Southern blot using genomic DNA for detection of KSHV should be performed
for distinction of PEL and circumstantial presence of KSHV owing to dissemi-nated viremia in an AIDS-associated lymphoma. For this, 5 µg of DNA may be
digested with BamHI, electrophoresed, and transferred using standard procedures
(7) . We recommend hybridization with a probe to ORF 26 as described (1) . If a
plasmid containing this insert is not available, this probe can be made by PCR
amplification of the ORF 26 (KS330233) fragment, using a PEL positive control
as template. A hybridization band of 330 bp will be obtained in positive cases.
This method also detects approx 75% of cases of KS.
References
1. Chang, Y., Cesarman, E., Pessin, M. S., Lee, F., Culpepper, J., Knowles, D. M.,
and Moore, P. S. (1994) Identification of herpesvirus-like DNA sequences in
AIDS-associated Kaposi’s sarcoma. Science 266, 1865–1869.
2. Cesarman, E. and Knowles, D. M. (1997) Kaposi’s sarcoma-associated herpesvi-rus: a lymphotropic human herpesvirus associated with Kaposi’s sarcoma, pri-mary effusion lymphoma, and multicentric Castleman’s disease. Semin. Diagn.
Pathol. 14, 54–66.
3. Ganem, D. (1997) KSHV and Kaposi’s sacroma: the end of the beginning? Cell
91,157–160.
4. Cesarman, E., Chang, Y., Moore, P. S., Said, J. W., and Knowles, D. M. (1995)
Kaposi’s Sarcoma-associated Herpesvirus-like DNA sequences in AIDS-related
body cavity-based lymphomas. N. Engl. J. Med. 332, 1186–1191.
5. Nador, R. G., Cesarman, E., Chadburn, A., Dawson, D. B., Ansari, M. Q., Said, J.,
and Knowles, D. M. (1996) Primary effusion lymphoma: a distinct clinicopatho-logic entity associated with the Kaposi’s sarcoma-associated herpesvirus. Blood
88, 645–656.
6. Cesarman, E. and Knowles, D. M. (1999) The role of Kaposi’s sarcoma-associated herpesvirus (KSHV/HHV-8) in lymphoproliferative diseases. Semin.
Cancer Biol. 9, 165–174.
7. Southern, E. M. (1975) Detection of specific sequences among DNA fragments
separated by gel electrophoresis. J. Mol. Biol. 98, 503–517.
Quantitative PCR for Cytomegalovirus 337
26
Diagnostic Applications of Quantitative
Polymerase Chain Reaction for Cytomegalovirus
Richard H. Scheuermann and Xin Bai
1. Introduction
Eight viruses in the herpes family have been identified that infect humans:
herpes simplex viruses 1 and 2, varicella-zoster virus, Epstein-Barr virus,
cytomegalovirus (CMV), human herpesviruses 6 and 7 and the Kaposi sarcoma–
associated herpesvirus  (1) . In immunocompetent individuals, primary infec-tions are usually handled effectively by the host immune system without
therapeutic intervention. However, these viruses are never completely eradi-cated by the immune response, probably because these viruses have the capac-ity to enter a latent state in a subset of infected cells. However, this does not
normally pose a problem since the host immune system has been primed to
handle any subsequent reactivation. Thus, human herpesviruses rarely cause
serious problems in immunocompetent individuals.
By contrast, individuals who are immunocompromised owing to congenital
abnormalities, therapeutic immunosupression, or human immunodeficiency
virus infection frequently experience life-threatening complications due to primary
infection or reactivation by these viruses. For example, bone marrow allograft
recipients receiving immunosupressive therapy have a high risk of developing
several complications related to CMV infection, including encephalitis, esoph-agitis, hepatitis, and pneumonitis; the latter is associated with a high mortality
rate (2 ,3) .
In normal patients, diagnosis of viral infection is usually made based on
serology, i.e., the detection of antibodies to the virus in question in the blood
(e.g. [4] ). However, the interpretation of serology for herpesviruses in immuno-337
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
338 Scheuermann and Bai
compromised patients is difficult owing to the high seroprevalence of herpes-viruses in the normal population, and the variable effects of immunosuppres-sion on antibody titers. For CMV, viral culture methods have been
developed for the direct detection of infectious viral particles. However, these
methods suffer from relatively low sensitivity and are very sensitive to sample-handling conditions. These culture techniques have largely been replaced by
an antigenemia test for the detection of CMV infection in the management of
immunosuppressed patients (5–8) .
More recently, the polymerase chain reaction (PCR) has been applied to the
detection of CMV in these patients. However, it appears that simple PCR is not
useful in the detection of clinically relevant disease since CMV can be
detected in a significant subset of normal, healthy individuals, presumably
owing to the presence of latently infected cells in the diagnostic sample (9–15) .
Therefore, this simple PCR procedure has been adapted to include an internal
calibration standard (ICS) in such a way as to provide clinically useful infor-mation (16) . ICS-PCR has two major advantages over simple PCR. First, it
provides quantitative determination of CMV levels that allows the distinction
between latent and active infection. Second, the use of an internal control helps
rule out false-negative results occasionally seen in PCR reactions when using
DNA isolated from patient samples. Thus, ICS-PCR provides an accurate,
objective, and sensitive technique for diagnosing and monitoring CMV dis-ease in immunosuppressed patients.
2. Materials
2.1. Isolation of DNA
QIAamp Blood Kit (PN 29104; QIAGEN, Santa Clara, CA) (see Note 13 for
an important update). For 50 isolations, the kit includes the following components:
1. Lyophilized QIAGEN Protease (28 mg).
2. Reagent AL1 (12 mL).
3. Reagent AL2 (3 mL).
4. Buffer AW concentrate (17 mL).
5. Buffer AE (12 mL).
6. Fifty QIAamp spin columns.
7. One hundred fifty 2-mL Microfuge collection tubes.
2.1.1. Preparation of Reagents from Kit
1. Protease solution: Add 1.4 mL of double-distilled H2O (ddH2O) to 28 mg of
QIAGEN protease and dissolve. Store in aliquots at –20°C.
2. AL buffer: Mix 12 mL of reagent AL1 with 3 mL of reagent AL2. Store in the
dark at room temperature.
Quantitative PCR for Cytomegalovirus 339
3. AW buffer: Add 40 mL of 100% ethanol to 17 mL of buffer AW concentrate.
Store at room temperature.
4. Sterile ddH20.
5. Microfuge tubes (1.5 mL).
6. 100% Ethanol.
7. Extra AL buffer (PN 19075; QIAGEN).
8. TE: 10 mM Tris-HCl, 1 mM EDTA, pH 8.0.
2.2. Polymerase Chain Reaction (see Note 1)
1. 10X AmpliTaq Buffer (PN N808-0006; Perkin-Elmer, Branchburg, NJ).
2. 10 mM each of dATP, dCTP, and dGTP, and 20 mM dUTP (see Note 2)
(PN N808-0095; Perkin-Elmer).
3. AmpliTaq DNA polymerase (5 U/µL) (PN N801-0060; Perkin-Elmer).
4. 5v CMV primer resuspended in ddH2O at 100 pmol/µL (TACCCCTATCG-CGTGTGTTC) (16) .
5. 3v CMV primer resuspended in ddH2O at 100 pmol/µL (ATAGGAGGCGC-CACGTATTC) (16) .
6. HHVQ-1 Internal Calibration Standard (available from authors; [16] ) or another
ICS incorporating CMV-specific primer sequences (10 molecules/µL TE). See
Note 3 for the characteristics of an effective ICS.
7. Microamp reaction tubes and caps.
8. Thermocycler with heated lid, e.g., Perkin-Elmer GeneAmp PCR System 9600
(see Note 1).
2.3. Detection of Product
1. 5X DNA sample buffer: 200 mM Tris-HCl acetate, 10 mM EDTA, 50% glycerol,
0.25% (w/v) bromophenol blue.
2. Agarose (molecular biology grade, e.g., Seakem GTG Agarose, FMC, Rock-land, ME).
3. 50X TAE buffer: 2.0 M Tris-HCl acetate, 50 mM EDTA. Dissolve 242 g of Tris
base in 600 mL of ddH2O while adding 57.1 mL of glacial acetic acid and
100 mL of 0.5 M EDTA, pH 8.0. Bring up the volume to 1.0 L with ddH2O.
4. DNA molecular weight marker, e.g., 123-bp DNA ladder (PN 15613-011; Gibco-BRL, Gaithersburg, MD) at 0.1 µg/µL in 1X DNA sample buffer.
5. Sybr Gold nucleic acid gel stain (PN S-11494; Molecular Probes, Eugene, OR)
(see Note 4), or ethidium bromide (EtBr) (PN E8751; Sigma, St. Louis, MO) at
10 mg/mL in ddH2O.
6. Gel-casting tray, electrophoresis apparatus, and DC power supply.
7. Imaging hardware (see Note 5): For analysis of gels stained with SYBR Gold,
illumination with a 488-nm laser and a fluorescence imaging system is used
(e.g., Fluorimager SI; Molecular Dynamics, Sunnyvale, CA). For analysis of gels
stained with EtBr, illumination with UV light at a 254-nm wavelength and analy-sis using a UV imaging system is used (e.g., AlphaImager 2000; Alpha Innotech,
San Leandro, CA).
340 Scheuermann and Bai
3. Methods
3.1. Isolation of DNA
In the early days of molecular analysis, DNA was isolated from cell sources
using organic extraction methods. The cells were first lysed with detergent,
proteins removed by phenol extraction, and the DNA purified and concentrated
by alcohol precipitation. Although this technique is effective, it is also some-what time-consuming and involves the use of hazardous chemicals. More
recently, several companies have developed DNA isolation kits based on a
number of nonorganic methods for DNA purification. These kits provide a
cost-effective means of isolating high-quality DNA without the use of organic
solvents. We have evaluated many of these kits for their suitability in the isola-tion of viral DNA from patient samples, especially whole blood, and found that
three give high-quality material suitable for most PCR applications: the
QIAamp Blood Kit (QIAGEN), the Puregene DNA Isolation Kit (GENTRA
Systems, Minneapolis, MN), and the Isoquick Nucleic Acid Extraction Kit
(ORCA Research, Bothell, WA). In this section, we describe a modification of
the QIAamp Blood Kit that we use for the analysis of CMV levels in patient
samples. Regardless of the method used, the key point is that the DNA material
isolated should allow sensitive and specific target amplification. In Subhead-ing 3.4.1., we provide an example of how one can evaluate any DNA isolation
procedure to determine whether the performance characteristics are optimal.
The isolation method employed in the QIAamp Blood Kit relies on the pref-erential binding of nucleic acids to silica gel surfaces in the presence of
chaotropic salts. The bound nucleic acids are washed to remove contaminants
and eluted in small volumes using a low-salt buffer (AE buffer). The silica gel
matrices are provided in a spin column format to facilitate rapid isolation. This
procedure can be applied to a wide variety of patient samples. Here we
describe the isolation of DNA from whole blood; for isolation from other
sample sources, one should refer to the product guide for specifics (see Note
13 for an important update):
1. Place 200 µL of blood in a sterile 1.5-mL microfuge tube.
2. Add 25 µL of protease solution.
3. Add 200 µL of AL buffer. Mix immediately by vortexing for 15 s.
4. Incubate the cell lysate at 70°C for 10 min.
5. Add 210 µL of 100% ethanol and mix by vortexing.
6. Apply the mixture to a QIAamp spin column seated in a 2-mL microfuge tube.
7. Centrifuge at 6000g (8000 rpm in a standard Eppendorf microfuge) for 1 min.
8. Discard the flowthrough.
9. Wash the column twice with 500 µL of AL buffer. (This step is a modification of the
original procedure that was developed in our laboratory to further improve the qual-ity of the isolated DNA specifically for PCR applications; see Subheading 3.4.1.)
Quantitative PCR for Cytomegalovirus 341
10. Wash the column twice with 500 µL of AW buffer. For the last wash, centrifuge
the column for 3 min at 20,000g (14,000 rpm) to remove as much of the wash
solution as possible.
11. Place the column in a clean 1.5-mL microfuge tube.
12. Elute DNA by adding 200 µL of AE buffer that has been preheated to 70°C to the
column. Incubate at 70°C for 5 min. Centrifuge the column 1 min to collect eluted
DNA. At this stage the purified DNA can be used directly in PCR reactions, further
concentrated by alcohol precipitation (see Note 6), or stored at –20°C for later use.
3.2. Polymerase Chain Reaction
The PCR reaction involves a variety of components and conditions that can
vary depending on the specific primer-template combination used. Rather than
trying to optimize these factors for every target we wish to analyze, we have
taken a different approach to the issue of optimization. When we are develop-ing a new procedure to amplify a new target, we evaluate a panel of primers
and select only the ones that work well under standard PCR reaction condi-tions. This approach has greatly simplified all PCR protocols in our laboratory
because they all work well under essentially the same amplification condi-tions. For a detailed discussion on primer selection in general and the primers
used here for detection and quantification of CMV, see Bai et al.  (16) . The
following protocol is for 50-µL PCR reactions:
1. Place purified DNA into individual microfuge tubes. For maximum sensitivity
(i.e., low limits of detection), we routinely use DNA isolated from 10 µl of blood
in 10 µL of AE elution buffer (see Note 7).
2. Add 2 µL of HHVQ-1 standard (10 molecules/µL) to each tube (see Note 8).
3. Determine the number of samples to be analyzed in a given experiment.
4. Design a master mix containing all the PCR components except the purified
sample DNA (see Table 1). A mix is prepared for 10% more than the required
number of samples. For example, for 10 samples prepare enough master mix for
11 samples; for 20 samples, prepare a master mix for 22 samples. In this
example, the analysis of 10 samples is described.
5. In a 1.5-mL microfuge tube prepare the PCR master mix as described in Table 1.
Mix by vortexing.
6. Add 38 µL of master mix to each tube containing purified blood DNA/standard
mixture. Vortex briefly.
7. Seal the tubes carefully with caps and place in a thermocycler.
8. Amplify under the following cycling conditions: 1 cycle of 94°C for 2 min;
34 cycles of 94°C for 0.4 min, 60°C for 0.4 min, 72°C for 1.0 min; 1 cycle of
72°C for 9.0 min; and hold at 8°C. After amplification is complete, the samples
can be stored for several hours at 4°C. For longer storage, –20 or –70°C is recom-mended. (N.B. The optimal conditions may vary depending on the thermocycler
used. See Note 1 for a discussion of how this can be evaluated. The conditions
described here work well for the Perkin-Elmer GeneAmp PCR System 9600.)
342 Scheuermann and Bai
3.3. Detection of PCR Product
Two methods are commonly used to detect and quantify specific PCR products
following amplification: staining following gel electrophoresis or product capture
and detection on a solid support (e.g., microtiter plate wells). The advantages to
these two detection methods are that each offers an additional layer of specificity to
the PCR reaction. In gel electrophoresis, the size of the DNA product is evaluated
and should correspond to the size predicted for the DNA target in question. In the
microtiter plate approach, capture and/or detection involves the hybridization of a
probe to specific sequences contained in the specific PCR product. A method for
the detection and quantification of CMV using the microtiter/capture approach has
been developed by BioSource International, Inc. (Camarillo, CA) (17). Here we
describe detection and quantification of PCR products following gel electrophoresis.
Although we give specific details of a procedure that works well for this analysis, many
other systems can also be used, including polyacrylamide gels and real-time PCR.
Table 1
Preparation of PCR Master Mix
Volume for 11 Volume/ Final
Component  reactions (µL) reaction (µL) concentration
ddH2O 310.2a 28.2a —
10X PCR buffer 55 5 1X
10 mM dATP 11 1 200 µM
10 mM dCTP 11 1 200 µM
10 mM dGTP 11 1 200 µM
20 mM dUTP 11 (see Note 2)1 400 µM
5v CMV primer 2.2 0.2 20 pmol
(100 pmol/µL)
3v CMV primer 2.2 0.2 20 pmol
(100 pmol/µL)
AmpliTaq 4.4 0.4 2 U
polymerase
5 U/µL)
a The amount of ddH2O to include in the reaction depends on the volume of sample to be
analyzed. In this example, we are analyzing a sample containing 10 µL of blood DNA and 2 µL
of internal standard in a final reaction volume of 50 µL. This means that 38 µL of the master mix
will need to be added to the DNA mixture. To calculate the amount of ddH2O to include in the
master mix, multiply 38 µL × 11 samples to give a final volume of 418 µL for the total master
mix. Then subtract the volumes of all the other master mix components to give the volume of
ddH2O required (418 – 55 – 11 – 11 – 11 – 11 – 2.2 – 2.2 – 4.4 = 310.2 µL). The volumes of the
other components are determined by the concentrations of the stock solutions used and the final
concentrations required in the PCR reaction.
Quantitative PCR for Cytomegalovirus 343
1. Add 2 g of agarose to 100 mL of 1X TAE buffer in a 500-mL Erlenmeyer flask.
Heat in a microwave on high setting for 2 to 3 min until all of the agarose is
dissolved. (The amount of agarose solution may be adjusted based on the size of
the gel to be used.)
2. Pour molten agarose into a gel-casting tray with a well comb in position. Allow it
to cool until solidified (~1 h at room temperature).
3. Place the gel in an electrophoresis apparatus. Cover with 1X TAE buffer.
4. Add 12 µL of 5X DNA sample buffer to each 50-µL PCR reaction sample. Mix
by vortexing.
5. Carefully load 20–30 µL of each sample into separate wells of the gel using a
micropipettor.
6. After all the samples are loaded, load an appropriate molecular weight marker
(e.g., 123-bp DNA ladder) into the next adjacent well.
7. Close the electrophoresis apparatus. Connect the cables to the power supply and
apparatus with the positive cathode positioned at the bottom end of the gel in the
direction of migration.
8. Electrophorese for 1.5 h at 100 V or until the bromophenol blue tracking dye is
within 1 cm of the end of the gel.
9. Turn off the power supply, remove the gel, and place in a staining solution con-taining either ethidium bromide (1 µg/mL) or Sybr Gold (1 10,000 dilution of
stock) for 30–60 min.
10. Examine EtBr-stained gels using UV wavelength illumination; examine Sybr
Gold–stained gels using 488 nm illumination.
3.4. Experimental Results
3.4.1. Quality of DNA Isolation Procedure
The quality of the DNA for PCR amplification can be evaluated in reactions that
contain a relatively large amount of material with a small number of specific targets. The
experiment presented in Fig. 1 shows an example of this kind of analysis. In this case, a
constant amount of ICS has been added to each PCR reaction with increasing amounts of
DNA isolated from whole blood using the standard and modified QIAamp procedure.
(The standard procedure lacks step 9 in Subheading 3.1.) In reactions containing DNA
isolated by the standard procedure, the addition of 0.1 or 1.0 µL of this DNA had no
effect on the ability to amplify the ICS (compare lanes 9 and 10 with lane 8). However,
if 10 or 20 µL of this DNA was added (lanes 11 and 12), inhibition of ICS amplification
was observed. On the other hand, even 20 µL of blood DNA isolated by the modified
procedure could be added without affecting amplification efficiency of the ICS (lane 6).
This indicates that DNA isolated by the standard procedure contains a PCR inhibitor
that reaches significant levels when 10 µL of this DNA is used under these conditions.
However, with a simple modification, these inhibitors can be largely removed and
more DNA included in the reaction. (See Note 13 for an important update on the DNA
isolation procedure.)
344 Scheuermann and Bai
The use of a standard molecule as an amplification target allows one to
assess amplification efficiency under the conditions used. In this case, we used
this approach to evaluate the quality of the DNA isolation procedure used.
However, a similar approach can be used to evaluate any of the components of
the PCR reaction. For example, if one is considering the use of an alternative
polymerase for amplification, it should be able to generate product when a
limiting amount of target is mixed with DNA isolated from a patient sample.
3.4.2. Determination of CMV Viral Burden in Patient Samples
CMV viral burden can be determined by using serial dilutions of patient
sample DNA amplified with a constant quantity of ICS. For analysis of whole
blood, DNA is isolated as described in Subheading 3.1. from 200 µL of blood
collected in acid citrate dextrose (ACD), EDTA, or even heparin tubes (see
Note 9). To achieve low limits of detection, as much of this purified DNA is
included in the first PCR reaction. In our experience, 10-µL samples can be
reproducibly used in these PCR reactions without adverse effects of inhibitors.
Additional reactions are also prepared using 1.0 µL and 1.0 µL of a 1/10 dilu-tion of this purified DNA. Each reaction contains a constant amount of ICS;
when conditions are optimal, 20 molecules of ICS can be routinely amplified.
Thus, for each patient sample, three PCR reactions are prepared with 10-fold
serial dilutions of purified patient DNA and a constant amount of ICS.
Fig. 1. Efficient PCR amplification from blood samples using a modified DNA
isolation protocol. DNA was isolated from 200  µL of whole blood from a healthy
volunteer using the standard QIAamp Blood Kit protocol (lanes 8–12) that does not
include step 9 in Subheading 3.1. or by a modification of this procedure (lanes 2–6)
that includes  step 9. PCR reactions were set up with CMV-specific primers, 300
molecules of the HHVQ-1 ICS, and varying amounts of this purified DNA (from 0 to
20 µL) as indicated. Reactions were amplified under standard conditions and products
analyzed following agarose gel electrophoresis and Sybr Gold staining. A 123-bp DNA
ladder (M) is included to determine product sizes (lane 1).
Quantitative PCR for Cytomegalovirus 345
Figure 2 presents results of this type of analysis for three different patient
samples. Using 10 µL of DNA isolated from patient 1, a single intense band is
observed derived from the CMV viral DNA (lane 2). In this case, no ICS-specific band is found owing to competition from the large amount of viral
target present in the reaction. Even with 1.0 µL of DNA, the ICS-specific band
is very faint when compared with the CMV-specific band (lane 3). Only with
0.1 µL of blood DNA from this patient can both bands be observed clearly
(lane 4). Quantification of the intensities of these two bands gives a virus ICS
ratio of 2 (see Note 10). Since 20 molecules of ICS were included in this reac-tion, this indicates that 40 viral targets were present in 0.1 µL of blood, or a
viral burden of 400,000 viral genomes/mL of blood.
For patient 2, clear bands from both the ICS and the CMV targets were
observed only when 10 µL of blood DNA was included in the PCR reaction
(lane 5). Quantification of the band intensities gives a virus ICS ratio of 1.5, in
a reaction containing 20 molecules of ICS. Thus, patient 2 has a viral burden of
3000 viral genomes/mL of blood.
Finally, no CMV-specific bands were observed in any of the samples with
DNA from patient 3. Using this procedure, we can estimate the limits of detec-Fig. 2. Analysis of CMV viral burden in whole blood from three different patient
samples. DNA was isolated from whole blood of three patient samples by the modi-fied QIAamp procedure. PCR reactions were set up with CMV-specific primers,
20 molecules of the HHVQ-1 ICS, and varying amounts of this purified DNA (from
0.1 to 10 µL) as indicated. Samples were also prepared that contained 67 molecules of
purified CMV DNA (lane 11) or no DNA (lane 12) as target. Reactions were amplified
under standard conditions and products analyzed following agarose gel electrophore-sis and Sybr Gold staining. The upper band derived from the ICS standard and the
lower band derived from the CMV viral genome are indicated with arrows. A 123-bp
DNA ladder (M) is included to determine product sizes (lane 1).
346 Scheuermann and Bai
tion in this experiment because we could have expected to see a CMV band
one fourth the intensity of the ICS-specific band in the 10  µL DNA sample
(lane 8). This would give a viral burden of <500 viral genomes/mL of blood for
patient 3. In addition, the importance of detecting an ICS-specific band in the
reactions using DNA from patient 3 is discussed in Note 11.
Lanes 11 and 12 contain positive and negative control samples, respectively,
which should always be included at the end of each set of reactions. The posi-tive sample contained 67 copies of CMV, and the ratio of intensities is approx
3, as expected. The absence of a CMV-specific band in the negative control
sample indicates that contamination of these PCR reactions during reaction
setup has not occurred in this experiment.
3.4.3. Conclusion
CMV viral monitoring can be used to improve the management of solid
organ and bone marrow transplant recipients. Sensitive techniques to identify
patients with high CMV levels before clinical manifestations of viral disease
allow the application of preemptive therapeutic approaches rather than pro-phylactic therapy of high-risk patients. This reduces the risk of clinical compli-cations associated with the antiviral therapies used and promises to reduce the
overall patient cost associated with these therapies (18–22) .
One of the challenges in using a sensitive method such as PCR for the detec-tion of CMV in the transplant population is to distinguish between clinically
relevant and latent infection. Growing evidence indicates that this distinction
can be made based on the differences in CMV levels in these two populations
( [23] ; our unpublished data). The use of ICSs allows quantitative measure-ment of CMV levels in patient samples.
From our studies, it appears that the cutoffs for clinically relevant detection are
levels above 10,000 viral targets/mL of blood, at least for cmv in the pediatric solid
organ transplant population. The numbers for the adult solid-organ transplant popu-lation appear to be similar. With this information in hand, it may be possible to
simplify the analysis described in  Subheading 3.4.2. so that only a single
PCR reaction would need to be evaluated for each patient sample (see Note 12).
In addition to its ability to facilitate quantification, ICS-PCR also provides
another important use in clinical diagnostic laboratories. By serving as an
internal positive control in every reaction, amplification of the ICS helps rule
out false-negative results.
4. Notes
1. We have found that the source of reagents and polymerase used in the PCR reac-tion and the thermocycler used for amplification can have dramatic effects on the
Quantitative PCR for Cytomegalovirus 347
results when low limits of detection are sought in a complex nucleic acid mix-ture. Using an evaluation strategy similar to the one described in Fig. 1, the suit-ability of other reagents and cycling parameters can be evaluated easily by
performing PCR reactions containing small amounts of ICS target (e.g., 20–50
molecules) in DNA isolated from relatively large amounts of blood (5–10 µL). If
the reagents give strong, specific product without nonspecific amplification, they
can be substituted for the ones recommended here.
2. In clinical laboratories, dUTP is routinely used in place of dTTP in the PCR
reactions. This is done to facilitate the elimination of PCR product carryover
from one experiment to another. Because PCR generates a large amount of spe-cific product and because it is also able to detect a small number of targets, prod-uct carryover is an important problem. To avoid this problem, it is important to
physically separate different steps in the amplification process, i.e., DNA isola-tion from PCR reaction setup from postamplification sample handling. In addi-tion, if dUTP is incorporated into the PCR products during amplification, these
molecules can be eliminated during subsequent reaction setup using uracil
N-glycosylase (PN N808-0068; Perkin-Elmer) which hydrolyzes DNA contain-ing uracil. Although we do not routinely include this step in our PCR analyses,
we include a negative control at the end of each experiment (see Fig. 2, lane 12)
to determine whether contamination has occurred. If contamination is evident,
the glycosylase step can then be included to allow clean results.
3. Because reaction components can have dramatic effects on amplification effi-ciency, it is essential to use an internal standard for accurate quantification. The
most critical characteristic of this internal standard is that it is amplified with the
same PCR primers as the target in the sample. The ICS is designed to give prod-uct that differs in size from product derived from the specific target in order to
distinguish them following gel electrophoresis. Here we describe the use of
HHVQ-1, which can be used as an ICS for all HHVs (16) . However, any DNA
molecule that incorporates the specific primers to be used for amplification and
can be distinguished from the specific target can be used. The advantage of using
HHVQ-1 is that the CMV primers incorporated into this ICS were selected after
screening more than 20 different primer pairs for specificity and low limits of
detection, and quantification using this ICS has been extensively validated.
4. The advantage of SYBR Gold over other DNA-binding dyes is that it gives
extremely low background fluorescence in agarose when it is not bound to DNA.
5. The key aspect of the imaging hardware used is that it allows the accurate quan-tification of the amount of PCR product generated following gel electrophoresis.
6. Purified DNA can be concentrated by ethanol precipitation. Add 20 µL of 3 M
sodium acetate to 200  µL of eluted DNA. Then add 600  µL of 100% ethanol.
Vortex briefly. Centrifuge at maximum speed in a microfuge for 10 min at room
temperature. Decant the supernatant. Briefly air-dry the DNA pellet and resus-pend in 10–20 µL of TE. However, keep in mind that this procedure may also
concentrate PCR inhibitors carried over during DNA isolation.
348 Scheuermann and Bai
7. Using DNA isolated by the modified version of the QIAamp procedure, we find
that we can routinely use 10  µL of blood DNA in PCR reactions without evi-dence of inhibition. Although blood collected in EDTA is preferable for maxi-mum sample stability, ACD and heparin blood samples can also be used with
satisfactory results. See Subheading 3.4.1. for further discussion.
8. Under the conditions described here, 20 ICS molecules can be routinely ampli-fied to give enough product to be visualized by standard gel electrophoresis and
staining protocols. However, for diagnostic use it appears that this level of detec-tion is probably not necessary. The most important criteria for determining the
amount of ICS to use is that it can be amplified reproducibly when mixed with
patient DNA. A discussion of the clinically relevant limits of detection is in
Subheading 3.4.2.
9. Many groups have found that DNA isolated from heparin blood samples is diffi-cult to amplify. We have found that by using the modified QIAamp isolation
procedure, these samples can be amplified with only small effects on amplifica-tion efficiency.
10. For reasons that are not entirely clear, quantification is most accurate when the
intensity ratio of the two products is close to 1, i.e., within a factor of 3 of a 1:1
ratio. Using three 10-fold serial dilutions, one of the reactions usually gives
results within these parameters.
11. It is difficult to overemphasize the value of using internal standards for identify-ing false-negative results in clinical PCR analysis. Many factors can have
adverse effects on amplification efficiency, not only inhibitors carried over dur-ing DNA isolation, but even effects of lot-to-lot difference in reagent composi-tions. Although the effects may be small, their impact is amplified with each
PCR cycle. For example, a reduction of 20% in amplification efficiency results
in a 36-fold reduction in the amount of PCR product after 34 cycles. The use of
an ICS is critical to verify that amplification was achieved with acceptable effi-ciency in each sample analyzed.
12. Although this procedure was established to maximize the limits of detection, it
may not be necessary to detect clinically relevant disease. For example, if one uses
10,000 viral targets/mL of blood for the clinically relevant cutoff, a single PCR reac-tion containing 2.0 µL of blood DNA and 20 molecules of ICS standard could be
used. A clinically relevant positive result would be indicated if the CMV-specific
band were more intense than the ICS-specific band, regardless of the exact quan-tification. This approach could be used for routine patient monitoring. For posi-tive samples, more accurate quantification using a series of DNA dilutions would
be useful to help monitor therapeutic responses.
13. Since this chapter was written, QIAGEN has replaced the QIAamp Blood Kit
with a second-generation kit (same part number) that incorporates the modifica-tions described in this chapter based on these findings. The new kit and protocol
can now be used as suggested by the manufacturer to give DNA with low levels
of PCR inhibitors without further modification.
Quantitative PCR for Cytomegalovirus 349
References
1. Zuckerman, A. J., Banatvala, J. E., and Pattison, J. R., eds. (1994) Principles and
Practice of Clinical Virology, John Wiley & Sons, West Sussex, England.
2. Zaia, J. A. (1994) Cytomegalovirus infection, in Bone Marrow Transplantation
(Forman, S. J., Blume, K. G., and Thomas, E. D., eds.), Blackwell Scientific,
Boston, MA, pp. 376–403.
3. Couriel, D., Canosa, J., Engler, H., Collins, A., Dunbar, C., and Barrett, A. J.
(1996) Early reactivation of cytomegalovirus and high risk of interstitial pneu-monitis following T-depleted BMT for adults with hematological malignancies.
Bone Marrow Transplant. 18, 347–353.
4. Arvin, A. M. (1992) Human cytomegalovirus, in Laboratory Diagnosis of Viral
Infections (Lennette, E. H., ed.), Marcel Dekker, New York, pp. 333–350.
5. Niubo, J., Perez, J. L., Martinez-Lacasa, J. T., Garcia, A., Roca, J., Fabregat, J.,
Gil-Vernet, S., and Martin, R. (1996) Association of quantitative cytomegalovi-rus antigenemia with symptomatic infection in solid organ transplant patients.
Diagn. Microbiol. Infect. Dis. 24, 19–24.
6. Salzberger, B., Franzen, C., Fatkenheuer, G., Cornely, O., Schwenk, A., Rasokat,
H., Diehl, V., and Schrappe, M. (1996) CMV-Antigenimia in peripheral blood for
the diagnosis of CMV disease in HIV-infected patients. J. Acquir. Immune Defic.
Syndr. Hum. Retrovirol. 11, 365–369.
7. Wetherill, P. E., Landry, M. L., Alcabes, P., and Friedland, G. (1996) Use of a
quantitative cytomegalovirus (CMV) antigenemia test in evaluating HIV+ patients
with and without CMV disease. J. Acquir. Immune Defic. Syndr. Hum. Retrovirol.
12, 33–37.
8. Murray, B. M., Amsterdam, D., Gray, V., Myers, J., Gerbasi, J., and Venuto, R.
(1997) Monitoring and diagnosis of cytomegalovirus infection in renal transplan-tation. J. Am. Soc. Nephrol. 8, 1448–1457.
9. Patel, R., Smith, T. F., Espy, M., Portela, D., Wiesner, R. H., Krom, R. A. F., and
Paya, C. V. (1995) A prospective comparison of molecular diagnostic techniques
for the early detection of cytomegalovirus in liver transplant recipients. J. Infect.
Dis. 171, 1010–1014.
10. Peiris, J. S. M., Taylor, C. E., Main, J., Graham, K., and Madeley, C. R. (1995)
Diagnosis of cytomegalovirus (CMV) disease in renal allograft recipients: the
role of semiquantitative polymerase chain reaction (PCR). Nephrol. Dial. Trans-plant. 10, 1198–1205.
11. Abecassis, M. M., Koffron, A. J., Kaplan, B., Buckingham, M., Muldoon, J. P.,
Cribbins, A. J., Kaufman, D. B., Fryer, J. P., Stuart, J., and Stuart, F. P. (1997)
The role of PCR in the diagnosis and management of CMV in solid organ recipi-ents; What is the predictive value for the development of disease and should PCR
be used to guide antiviral therapy? Transplantation 63, 275–279.
12. Lo, C. Y., Ho, K. N., Yuen, K. Y., Lui, S. L., Li, F. K., Chan, T. M., Lo, W. K.,
and Cheng, K. P. (1997) Diagnosing cytomegalovirus disease in CMV seroposi-
350 Scheuermann and Bai
tive renal allograft recipients: a comparison between the detection of CMV
DNAemia by polymerase chain reaction and antigenemia by CMV pp65 assay.
Clin. Transplant. 11, 286–293.
13. Imbert-Marcille, B.-M., Cantarovich, D., Ferre-Aubineau, V., Richet, B.,
Soulillou, J.-P., and Billaudel, S. (1997) Usefulness of DNA viral load quantifica-tion for cytomegalovirus disease monitoring in renal and pancreas/renal trans-plant recipients. Transplantation 63, 1476–1481.
14. Lao, W. C., Lee, D., Burroughs, A. K., Lanzini, G., Rolles, K., Emery, V. C., and
Griffiths, P. D. (1997) Use of polymerase chain reaction to provide prognostic
information on human cytomegalovirus disease after liver transplantation. J. Med.
Virol. 51, 152–158.
15. Stephan, F., Fajac, A., Grenet, D., Honderlick, P., Ricci, S., Frachon, I., Friard,
S., Caubarrere, I., Bernaudin, J.-F., and Stern, M. (1997) Predictive value
of cytomegalovirus DNA detection by polymerase chain reaction in blood
and broncoalveolar lavage in lung transplant patients.  Transplantation 63,
1430–1435.
16. Bai, X., Hosler, G., Rodgers, B. B., Dawson, D. B., and Scheuermann, R. H.
(1997) Quantitative PCR for human herpes viruses and measurement of Epstein
Barr virus burden in post-transplant lymphoproliferative disorder.  Clin. Chem.
43, 1843–1849.
17. Reagan, K. J., Cabradilla, C., Shuman, B., Stollar, N., Laudemann, J., Bai, X.,
Hosler, G., and Scheuermann, R. H. (1998) Analytical performance of a quantita-tive CMV DNA detection method, in  CMV-Related Immunopathology. Mono-graphs in Virology, vol. 21 (Scholz, M., Rabenau, H. F., Doerr, H. W., and Cinatl,
J. Jr., eds.), Karger, Basel, Switzerland, pp. 252–261.
18. Einsele, H., Ehninger, G., Hebart, H., Wittkowski, K. M., Schuler, U., Jahn, G.,
Mackes, P., Herter, M., Klingebiel, T., Loffler, J., Wagner, S., and Muller, C. A.
(1995) Polymerase chain reaction monitoring reduces the incidence of cytomega-lovirus disease and the duration and side effects of antiviral therapy after bone
marrow transplantation. Blood 86, 2815–2820.
19. Brennan, D. C., Garlock, K. A., Lippmann, B. A., Buller, R. S., Gaudreault-Keener, M., Lowell, J. A., Miller, S. B., Shenoy, S., Howard, T. K., and Storch,
G. A. (1997) Control of cytomegalovirus-associated morbidity in renal transplant
patients using intensive monitoring and either preemptive or deferred therapy.
J. Am. Soc. Nephrol. 8, 118–125.
20. Gotti, E., Suter, F., Baruzzo, S., Perani, V., Moioli, F., and Remuzzi, G. (1996)
Early ganciclovir therapy effectively controls viremia and avoids the need for
cytomegalovirus (CMV) prophylaxis in renal transplant patients with cytomega-lovirus antigenemia. Clin. Transplant. 10, 550–555.
21. Egan, J. J., Lomax, J., Barber, L., Lok, S. S., Martyszcuzuk, R., Yonan, N., Fox,
A., Deiraniya, A. K., Turner, A. J., and Woodcock, A. A. (1998) Preemptive treat-ment for the prevention of cytomegalovirus disease in lung and heart transplant
recipients. Transplantation 65, 747–752.
Quantitative PCR for Cytomegalovirus 351
22. Hebart, H., Kanz, L., Jahn G., and Einsele, H. (1998) Management of cytomega-lovirus infection after solid-organ or stem-cell transplantation; current guidelines
and future prospects. Drugs 55, 59–72.
23. Drouet, E., Colimon, R., Michelson, S., Fourcade, N., Neveleau, A., Ducerf, C.,
Boibieux, A., Chevallier, M., and Denoyel, G. (1995) Monitoring levels of human
cytomegalovirus DNA in blood after liver transplantation. J. Clin. Microbiol. 33,
389–394.
PCR Detection of HSV in CSF 353
27
A Colorimetric Microtiter Plate Polymerase
Chain Reaction System That Detects
Herpes Simplex Virus in Cerebrospinal
Fluid and Discriminates Genotypes 1 and 2
Yi-Wei Tang
1. Introduction
Herpes simplex virus (HSV) is an ubiquitous agent responsible for a wide
variety of human infections. In addition to epithelial infections such as gingi-vostomatitis, pharyngitis, genital herpes, whitlow, conjunctivitis, and keratitis,
HSV is an important cause of central nervous system (CNS) infections and
accounts for 2–19% of human encephalitis cases (1 ,2) . The clinical spectrum
of CNS diseases has been recently expanded; for example, most cases of
benign recurrent aseptic meningitis (Mollaret meningitis) are caused by HSV
(3) , especially HSV-2 (4) . Because specific antiviral therapy is available, the
rapid, definitive laboratory diagnosis of HSV is important to support clinical
findings. Moreover, in the setting of possible HSV encephalitis, patients are
often managed as inpatients while awaiting test results.
Although cell culture is considered the standard method for laboratory diag-nosis of ulcerative HSV infections, HSV is rarely recovered in cell cultures
inoculated with cerebrospinal fluid (CSF). Brain biopsy specimens may yield
culturable virus, but this invasive surgical procedure is controversial when per-formed solely to collect specimens for the laboratory diagnosis of infectious
disease. The sensitivity of HSV antigen and antibody assays for CNS infec-tions is very low  (5) . In addition, antibodies may appear in the CSF as the
consequence of a breakdown in the blood-brain barrier, leading to false-positive results (6) .
353
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
354 Tang
The diagnosis of HSV CNS infections has recently been facilitated by the
development of polymerase chain reaction (PCR) technology. As an alterna-tive to the aforementioned techniques, the detection of HSV DNA in the CSF
of patients with suspected HSV encephalitis or meningitis allows rapid and
noninvasive confirmation of the diagnosis. With the utilization of primers from
an HSV DNA sequence that was common to both HSV-1 and HSV-2, several
investigators (7–13) have reported successful identification of HSV DNA in
the CSF. Further studies have shown that PCR detection of HSV DNA in CSF
should be considered the new standard for laboratory diagnosis of CNS disease
caused by this virus (14–18) .
Although the technology underlying PCR is relatively rapid, the PCR prod-uct (amplicon) must be identified definitively as the sequence of interest to
provide adequate diagnostic specificity. The conventional technique for this is
the hybridization of a specific probe to a Southern blot, increasing both sensi-tivity and specificity of the test. This step, however, takes an additional
12–24 h to complete, delaying the use of test results for clinical intervention.
The ideal postamplification detection system would combine the increased sen-sitivity and specificity of the Southern blot with rapid turnaround time. For this
purpose, enzyme-linked adsorbent microtiter plate systems have been adapted
for amplicon identification (15 ,19–21) .
2. Materials
2.1. Special Instruments
1. Thermal cycler (PE 9600 or 9700 is preferred).
2. Enzyme-linked immunosorbent assay (ELISA) automatic washer and reader.
3. Shaking incubator.
4. Positive displacement pipettor.
5. Multichannel pipettor.
2.2. Extraction of DNA
1. RNase-free water.
2. Isopropanol.
3. Ethanol.
4. Pellet Paint Co-Precipitant (Novagen, Madison, WI).
5. IsoQuick DNA Extraction Kit (Orca Research, Bothell, WA).
2.3. PCR Amplification
1. RNase-free water.
2. GeneAmp 10X PCR Buffer II (Perkin-Elmer, Foster City, CA).
3. Deoxynucleotide mixture (dNTPs) (Roche Diagnostics, Indianapolis, IN).
PCR Detection of HSV in CSF 355
4. Primers for PCR, (polyacrylamide gel electrophorisis (PAGE) purified.
Sequences are listed in Table 1.
5. Digoxigenin-11-2vdeoxyuridine-5v-triphosphate (DIG-dUTP) (Roche Diagnostics).
6. Glycerol.
7. Ampli Taq polymerase (Perkin-Elmer).
8. Uracil N-glycosylase (UNG) (Epicentre Technologies, Madison, WI).
2.4. Reagents for Colorimetric Detection
1. PCR ELISA (DIG detection) Kit (Roche Diagnostics) (see Note 1).
2. Capture probe sequences are listed in  Table 1. These are PAGE purified and
5v biotinylated.
3. Methods
The protocol is divided into three stages: nucleic acid extraction, PCR
amplification, and amplicon identification.
3.1. Extraction of Nucleic Acid by IsoQuick Extraction Kit
A modified solvent-extraction procedure (IsoQuick) is used for DNA
extraction. The procedure described here is slightly modified from the instruc-tions provided by Ocra Research (see Note 2).
1. Prepare lysate by mixing an equal volume (200 µL) of reagent 1 (lysis solution)
to a CSF sample in a 2.0-mL microcentrifuge tube. Mix the tube.
2. Shake reagent 2 (extraction matrix) vigorously. Add 750 µL to sample lysate.
3. Add 500 µL of reagent 3 (extraction buffer) to the sample. Vortex thoroughly.
4. Centrifuge at 12,800g for 5 min.
Table 1
Characteristics of Primers Used to Detect HSV DNA in CSF
Genbank accession
Primer Description no. (reference) Sequence (5′ A 3′)a
TK-A Upstream X03764 (22) GAC MAG CGC CCA GAT AAC AA
PCR primer    and X01712 (23)
TK-B Downstream X03764 (22) MCA GCA TRG CCA GGT CAA GC
PCR primer    and X01712 (23)
TK-G HSV-1-specific X03764 (22) ACA AAC ATC GTG TTG GGG GC
probe
TK-H HSV-2-specific X01712 (23) ACG AAC CTG GTC CTG GGT GT
probe
a M = A or C; R = A or G.
356 Tang
5. Transfer the upper aqueous phase (about 500  µL) to a new 1.5-mL micro-centrifuge tube.
6. Add 0.1 vol (50 µL) of reagent 4 (sodium acetate) to the aqueous-phase sample.
7. Add 2.0 µL of Pellet Paint Co-Precipitant to the aqueous-phase sample.
8. Add an equal volume (500 µL) of isopropanol to the aqueous-phase sample. Mix
gently by inversion to precipitate nucleic acid. Precipitate for 30 min at –20°C. A
visible pink pellet should be seen on the bottom of the tube.
9. Centrifuge at 12,800g for 30 min.
10. Discard the supernatant by aspirating the alcohol without disturbing the pink
nucleic acid pellet.
11. Add 1 mL of 70% ethanol to the pellet. Mix gently by inverting the tube several
times.
12. Centrifuge at 12,800g for 5 min.
13. Discard the supernatant by aspirating the alcohol without disturbing the nucleic
acid pellet. Allow the DNA pellet to air-dry at room temperature.
14. Resuspend the DNA pellet in 25 µL of reagent 5 (RNase-free water). Allow the
pellet to dissolve at room temperature with occasional gentle mixing.
3.2. PCR amplification
A specific 335-bp DNA sequence encoding a portion of the thymidine
kinase (TK) gene (Fig. 1) is amplified by PCR (22 ,23)  (see Note 3).
1. Prepare the PCR master mix as follows: 1X PCR Buffer II; 1.5 mM MgCl2;
200 µM each dATP, dCTP, and dGTP; 100 µM dTTP; 90 µM dUTP; 10 µM DIG-dUTP; 1 µM each primer TK-A and TK-B; 10% glycerol; 0.01 U/µL of UNG;
0.025 U/µL Ampli Taq polymerase.
2. Aliquot 45 µL of master mix into 0.2-mL MicroAmp tubes using a positive dis-placement pipettor.
Fig. 1. Gene location and nucleotide base position of primers and probes designed
to detect HSV DNA and differentiate genotypes 1 and 2.
PCR Detection of HSV in CSF 357
3. Add 5 µL of sample extract to the master mix.
4. Transfer the tubes to the DNA thermal cycler.
5. Run the PCR thermal profile using the following parameters: 50°C for 5 min,
94°C for 3 min, 50 cycles of 94°C for 15 s and 60°C for 30 s, 72°C for 10 min,
4°C soak. Approximate run time is 2.2 h on the PE thermocycler 9600. PCR
reactions may be left in the thermocycler overnight at 4°C to soak.
6. After the profile is completed, remove the tubes from the thermocycler. If
amplicon identification is not to be completed the same day, store the tubes at
–20°C.
3.3. Identification of Amplicon
Identification of amplicon is performed using an ELISA microtiter format.
Denatured amplicon is mixed with a hybridization solution containing a
biotin-labeled DNA capture probe specific for HSV-1 or HSV-2. The probes
hybridize to the corresponding target DNA sequence if present, and the result-ing complexes are captured on the streptavidin-coated microtiter plate wells.
HSV-specific DNA complexes are detected by anti-DIG-peroxidase conjugate,
and the peroxidase substrate is added (Fig. 2). Determination of the presence
of HSV-1 or HSV-2 is then made based on the color production (21) .
The following procedure is slightly modified from the instructions provided
by Roche Diagnostics:
1. Complete the HSV PCR ELISA tray map using the work sheet provided in the
kit. Calculate the amount of each reagent required for the batch run. Prepare an
excess of each reagent to allow for multichannel pipetting losses. As a rule of
thumb, prepare enough reagent for at least eight extra wells.
2. Reconstitute the lyophilized anti-DIG-peroxidase stock (vial 7) with 250 µL of
sterile water.
3. Dispense the calculated volume of hybridization buffer (vial 4) to two 50-mL
conical tubes, and add the calculated volume of HSV capture probes TK-G and
TK-H (final concentration of 7.5 pmol/mL) to the appropriate tubes. Mix gently
and avoid foaming.
4. Prepare the anti-DIG-peroxidase working solution by diluting 1 vol of the recon-stituted solution from vial 7 (step 3) with 99 vol of conjugate dilution buffer (vial
6) in a 50-mL conical tube using the calculated values on the work sheet.
5. Prepare an adequate volume of 2,2v-azinobis (3-ethylbenzothizoline-6-sulfonic
acid), diammonium salt (ABTS) substrate solution for the number of wells being
detected. Dissolve one tablet of ABTS (vial 9) per 5 mL of substrate buffer (vial
8) in a 50-mL conical tube using the calculated values on the work sheet. Steps 4
and 5 should be done at least 1 h prior to use. Mix gently and avoid foaming.
Store the solution away from light at room temperature. Prepare a fresh reagent
each day of use.
6. Prepare a wash solution by dissolving one wash tablet (vial 5) in 2 L of distilled
water. Mix thoroughly prior to use.
358 Tang
7. Add 10 µL of reconstituted DIG-labeled Control PCR product to a 1.5-mL tube.
Add 40 µL of denaturation solution to the tube. Mix the contents up and down
five times, and allow the reaction to incubate 10 min at room temperature.
8. Add 40  µL of denaturation solution to each 50-µL PCR reaction tube using a
multichannel pipettor. Mix the contents up and down five times and allow the
reactions to incubate for 10 min at room temperature.
9. Just prior to the end of the 10-min incubation, add 180 µL of the two hybridiza-tion solutions containing 7.5 pmol/mL of capture probe (TK-G or TK-H) to the
Fig. 2. Detection of DIG-labeled PCR amplicon with the Roche PCR ELISA (DIG
detection) Kit. (A) During PCR amplification, Taq DNA polymerase incorporates DIG-dUTP into the target DNA.  (B) A biotin-labeled oligonucleotide probe “captures” the
DIG-labeled PCR amplicon.  (C) The probe-amplicon hybrid is immobilized on a
streptavidin-coated microtiter plate.  (D) The immobilized probe-amplicon hybrid is
detected with peroxidase-conjugated anti-DIG antibody and ABTS colorimetric substrate.
PCR Detection of HSV in CSF 359
alternating rows of the detection wells according to the tray map using a multi-channel pipettor. Well A1 is designated as the reagent control. Add 180 µL of
hybridization buffer (vial 4) to the well. All the reagents should be added with the
exception of target DNA throughout the procedure. Well B1 is designated as the
kit positive control well. Add 180 µL of the hybridization buffer/control capture
probe mixture prepared earlier. Add 20  µL of denatured DIG-labeled Control
PCR product to this well.
10. Add 20 µL of denatured amplicon to the appropriate wells. Mix up and down five
times using a multichannel pipettor. Seal the plate with a mylar sheet provided
in the kit. Each specimen is detected in duplicate; that is, 20  µL of denatured
amplicon is added to a well containing TK-G and to a well in the parallel row
containing probe TK-H.
11. Place the plate containing the microtiter strips into the 37°C rotary incubator for
3 h at 60 rpm in the dark.
12. Remove the plate and wash each well five times on the automatic plate washer
with the wash solution prepared previously.
13. Add 200 µL of freshly prepared anti-DIG-peroxidase working solution to each
well using a multichannel pipettor. Seal the plate and incubate the solution in the
37°C rotary incubator for 30 min at 60 rpm in the dark.
14. Remove the plate and wash each well five times on the automatic plate washer
with the wash solution prepared previously.
15. Add 200 µL of freshly prepared ABTS substrate to each well using a multichan-nel pipettor.
16. Seal the plate and incubate the solution in the 37°C rotary incubator for 30 min at
60 rpm in the dark.
17. Remove the mylar sheet cover and immediately read the plate on the plate reader
using dual wavelengths (405/490 nm) with blank correction.
3.4. Interpretation of Results
The results are strictly qualitative. The presence of HSV DNA is determined
by relating the absorbance of the specimen well to the intrinsic extinction of
the ABTS solution well. A clinical specimen with an OD405/490 *0.1 should be
interpreted as positive for the presence of HSV DNA. A clinical specimen with
an OD405/490 <0.1 should be interpreted as negative for the presence of HSV
DNA. If positive, the specimen is identified as being positive for HSV-1 or
HSV-2 or both. The results should be interpreted with consideration of other
clinical laboratory findings. A negative result does not eliminate the possibil-ity of infection. Reliable results depend on an adequate specimen collection
and the absence of inhibitory substances in the specimens.
4. Notes
1. Although only one colorimetric microtiter plate PCR system is described for the
detection of HSV in the CSF, several equivalent systems are commercially avail-
360 Tang
able. They include the PrimeCapture™ from ViroMed Laboratories, Quanti-PATH™ from CPG, and GEN-ETI-K™ from DiaSorin. Our evaluation indicated
that the majority of those systems, including the one described herein, have a
comparable sensitivity and specificity in comparison to the conventional South-ern blot followed by probe hybridization (21) .
2. Ideally, separate rooms are designed for nucleic acid extraction, PCR amplifica-tion, and amplicon identification in order to avoid cross contamination. How-ever, laboratories with limited space can separate these functions by established
separate workstations in two rooms, one for preamplification steps and another
for amplification and postamplification steps. In addition, a UNG-based inacti-vation system was adapted to control possible contamination by amplicon
carryover (21 ,24) .
3. Additional advantages of the described system include the recognition of poly-morphisms in the TK gene that may be responsible for drug resistance. Several
point mutations in the TK gene may also be responsible for acyclovir resistance
(25–27) . The determination of acyclovir resistance by detection of these point
mutations may be important in patients undergoing long-term therapy and in
immunocompromised hosts  (27 ,28) . Acyclovir resistance could be determined
by direct sequencing of the amplicon based on clinical management.
Acknowledgment
This chapter contains information presented in the Molecular Microbiology
Laboratory Procedure Manual used at the Mayo Clinic. I acknowledge the
contributions to this manual from Paul N. Rys, Mark J. Espy, Shawn P.
Mitchell, Dr. Thomas F. Smith, and Dr. David H. Persing.
References
1. Skoldenberg, B., Forsgren, M., Alestig, K., Bergstrom, T., Burman, L., Dahlqvist,
E., Forkman, A., Fryden, A., Lovgren, K., and Norlin, K. (1984) Acyclovir versus
vidarabine in herpes simplex encephalitis: randomised multicentre study in con-secutive Swedish patients. Lancet 2, 707–711.
2. Whitley, R. J., Alford, C. A., Hirsch, M. S., Schooley, R. T., Luby, J. P., Aoki,
F. Y., Hanley, D., Nahmias, A. J., and Soong, S. J. (1986) Vidarabine ver-sus acyclovir therapy in herpes simplex encephalitis.  N. Engl. J. Med. 314,
144–149.
3. Yamamoto, L. J., Tedder, D. G., Ashley, R., and Levin, M. J. (1991) Herpes sim-plex virus type 1 DNA in cerebrospinal fluid of a patient with Mollaret’s meningi-tis. N. Engl. J. Med. 325, 1082–1085.
4. Tedder, D. G., Ashley, R., Tyler, K. L., and Levin, M. J. (1994) Herpes simplex
virus infection as a cause of benign recurrent lymphocytic meningitis.  Ann.
Intern. Med. 121, 334–338.
5. Kohl, S. (1988) Herpes simplex virus encephalitis in children. Pediatr. Clin. North
Am. 35, 465–483.
PCR Detection of HSV in CSF 361
6. Nahmias, A. J., Whitley, R. J., Visintine, A. N., Takei, Y., and Alford, C. A. Jr.
(1982) Herpes simplex virus encephalitis: laboratory evaluations and their diag-nostic significance. J. Infect. Dis. 145, 829–836.
7. Powell, K. F., Anderson, N. E., Frith, R. W., and Croxson, M. C. (1990) Non-invasive diagnosis of herpes simplex encephalitis. Lancet 335, 357,358.
8. Rowley, A. H., Whitley, R. J., Lakeman, F. D., and Wolinsky, S. M. (1990) Rapid
detection of herpes-simplex-virus DNA in cerebrospinal fluid of patients with
herpes simplex encephalitis. Lancet 335, 440,441.
9. Puchhammer-Stockl, E., Popow-Kraupp, T., Heinz, F. X., Mandl, C. W., and
Kunz, C. (1990) Establishment of PCR for the early diagnosis of herpes simplex
encephalitis. J. Med. Virol. 32, 77–82.
10. Klapper, P. E., Cleator, G. M., Dennett, C., and Lewis, A. G. (1990) Diagnosis of
herpes encephalitis via Southern blotting of cerebrospinal fluid DNA amplified
by polymerase chain reaction. J. Med. Virol. 32, 261–264.
11. Kimura, H., Futamura, M., Kito, H., Ando, T., Goto, M., Kuzushima, K., Shibata,
M., and Morishima, T. (1991) Detection of viral DNA in neonatal herpes simplex
virus infections: frequent and prolonged presence in serum and cerebrospinal
fluid. J. Infect. Dis. 164, 289–293.
12. Aurelius, E., Johansson, B., Skoldenberg, B., Staland, A., and Forsgren, M. (1991)
Rapid diagnosis of herpes simplex encephalitis by nested polymerase chain reac-tion assay of cerebrospinal fluid. Lancet 337, 189–192.
13. Aslanzadeh, J., Osmon, D. R., Wilhelm, M. P., Espy, M. J., and Smith, T. F.
(1992) A prospective study of the polymerase chain reaction for detection of her-pes simplex virus in cerebrospinal fluid submitted to the clinical virology labora-tory. Mol. Cell. Probes 6, 367–373.
14. Lakeman, F. D. and Whitley, R. J. (1995) Diagnosis of herpes simplex encephali-tis: application of polymerase chain reaction to cerebrospinal fluid from brain-biopsied patients and correlation with disease. J. Infect. Dis. 171, 857–863.
15. Vesanen, M., Piiparinen, H., Kallio, A., and Vaheri, A. (1996) Detection of her-pes simplex virus DNA in cerebrospinal fluid samples using the polymerase chain
reaction and microplate hybridization. J. Virol. Methods 59, 1–11.
16. Tang, Y. W., Espy, M. J., Persing, D. H., and Smith, T. F. (1997) Molecular evi-dence and clinical significance of herpesvirus coinfection in central nervous sys-tem. J. Clin. Microbiol. 35, 2869–2872.
17. Mitchell, P. S., Espy, M. J., Smith, T. F., Toal, D. R., Rys, P. N., Berbari, E. F.,
Osmon, D. R., and Persing, D. H. (1997) Laboratory diagnosis of central nervous
system infections with herpes simplex virus by PCR performed with cerebrospi-nal fluid specimens. J. Clin. Microbiol. 35, 2873–2877.
18. Tang, Y. W., Mitchell, P. S., Espy, M. J., Smith, D. H., and Persing, D. H. (1999)
Molecular diagnosis of herpes simplex virus infections in the central nervous sys-tem [minireview]. J. Clin. Microbiol. 37, 2127–2136.
19. Mantero, G., Zonaro, A., Albertini, A., Bertolo, P., and Primi, D. (1991) DNA
enzyme immunoassay: general method for detecting products of polymerase chain
reaction. Clin. Chem. 37, 422–429.
362 Tang
20. Loeffelholz, M. J., Lewinski, C. A., Silver, S. R., Purohit, A. P., Herman, S. A.,
Buonagurio, D. A., and Dragon, E. A. (1992) Detection of Chlamydia trachomatis
in endocervical specimens by polymerase chain reaction. J. Clin. Microbiol. 30,
2847–2851.
21. Tang, Y. W., Rys, P. N., Rutledge, B. J., Mitchell, P. S., Smith, T. F., and Persing,
D. H. (1998) Comparative evaluation of colorimetric microtiter plate systems for
detection of herpes simplex virus in cerebrospinal fluid. J. Clin. Microbiol. 36,
2714–2717.
22. Graham, D., Larder, B. A., and Inglis, M. M. (1986) Evidence that the ‘active
centre’ of the herpes simplex virus thymidine kinase involves an interaction
between three distinct regions of the polypeptide. J. Gen. Virol. 67, 753–758.
23. Swain, M. A. and Galloway, D. A. (1983) Nucleotide sequence of the herpes sim-plex virus type 2 thymidine kinase gene. J. Virol. 46, 1045–1050.
24. Longo, M. C., Berninger, M. S., and Hartley, J. L. (1990) Use of uracil DNA
glycosylase to control carry-over contamination in polymerase chain reactions.
Gene 93, 125–128.
25. Englund, J. A., Zimmerman, M. E., Swierkosz, E. M., Goodman, J. L., Scholl,
D. R., and Balfour, H. H. Jr. (1990) Herpes simplex virus resistant to acyclovir.
A study in a tertiary care center. Ann. Intern. Med. 112, 416–422.
26. Palu, G., Gerna, G., Bevilacqua, F., and Marcello, A. (1992) A point mutation in
the thymidine kinase gene is responsible for acyclovir-resistance in herpes sim-plex virus type 2 sequential isolates. Virus Res. 25, 133–144.
27. Gaudreau, A., Hill, E., Balgour, H. H. Jr., Erice, A., and Boivin, G. (1998) Pheno-typic and genotypic characterization of acyclovir-resistant herpes simplex viruses
from immunocompromised patients. J. Infect. Dis. 178, 297–303.
28. Pease, A. C., Solas, D., Sullivan, E. J., Cronin, M. T., Holmes, C. P., and Fodor,
S. P. (1994) Light-generated oligonucleotide arrays for rapid DNA sequence
analysis. Proc. Natl. Acad. Sci. USA 91, 5022–5026.
Detection and Typing of HCV 363
28
Detection and Typing of Hepatitis C Virus
Frederick S. Nolte
1. Introduction
For more than two decades prior to the discovery of the hepatitis C virus
(HCV), posttransfusion non-A, non-B (NANB) hepatitis was thought to have a
viral etiology. In 1989, the virus was finally identified through a unique appli-cation of molecular cloning techniques by investigators at the Centers for Dis-ease Control, and the Chiron Corporation  (1) . In a reversal of the usual
sequence of events, HCV was identified and defined before its existence was
substantiated through tissue culture growth, electron microscopic observation,
or serologic detection. Molecular cloning of HCV preceded the use of any of
the conventional methods of viral identification and serologic detection then
evolved from the blind immunoscreening of millions of clones with serum from
a patient who had NANB hepatitis (2) . The peptide expressed in a single reac-tive clone (5-1-1) served as the basis for the first Food and Drug Administra-tion (FDA)–licensed diagnostic test for detection of antibodies to HCV (3) .
HCV is now recognized as the cause of most cases of posttransfusion NANB
hepatitis and more than half of sporadic NANB hepatitis. About half of all
infected individuals develop chronic hepatitis and about 20% of these may
develop cirrhosis. It is estimated that nearly 4 million people are chronically
infected with HCV in the United States. End-stage liver disease owing to
chronic HCV infection is the leading reason for liver transplantation in the
United States.
The entire genome of HCV has been cloned and sequenced. It is a single-stranded, positive sense RNA virus composed of approx 10,000 nucleotides
coding for 3000 amino acids. The genome consists of a single open reading
frame that is translated to yield a polyprotein from which viral proteins are
363
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
364 Nolte
derived by posttranslational cleavage. HCV exists as a heterogeneous group of
viruses showing about 70% homology overall with six distinct major geno-types and a number of subtypes (4). The virus is related to the Flaviviridae and
Pestiviridae and has a high spontaneous mutation rate (10–2 mutations/
[nucleotide · yr]) (5,6) . As a result of this high spontaneous mutation rate, indi-vidual patients are infected with a heterogeneous population of virus having
closely related yet heterogeneous genomes termed  quasispecies. The extensive
genetic heterogeneity of HCV has important diagnostic and clinical implications.
Detection of HCV RNA in serum by nucleic acid amplification methods has
become important in confirming the diagnosis of hepatitis C and in assessing
response to antiviral therapy (7) . Many different qualitative and quantitative
assays for HCV RNA have been described. These assays have employed
reverse transcriptase polymerase chain reaction (RT-PCR), nucleic acid
sequence-based amplification, and branched DNA as amplification methods
(7) . Standardization of in-house-developed RT-PCR assays for HCV RNA has
been difficult, as demonstrated by the results of a recent international collabo-rative proficiency testing study (8) . In this performance survey of 86 laborato-ries, only 16% of the laboratories performed faultlessly, with 29% missing
only a weak positive specimen, and 55% reporting false-positive or false-negative results. An RT-PCR test for HCV RNA for only research use is avail-able from Roche Molecular Systems (AMPLICOR HCV Test) (9) . The test has
undergone several preclinical evaluations and is currently used in FDA clinical
trials in the United States (9–11) . Because this test will likely become the first
standardized HCV RNA test marketed to clinical laboratories in the United
States, it is the method described in detail in this chapter.
The AMPLICOR test is based on five major processes: specimen prepara-tion, reverse transcription of target RNA to generate complementary DNA
(cDNA), PCR amplification of target cDNA using HCV-specific complemen-tary primers, hybridization of the amplified products to oligonucleotide probes
specific to the target(s), and detection of the probe-bound amplified products
by color formation.
HCV RNA is isolated from serum or plasma by lysis of virus particles with
a chaotropic agent followed by precipitation of the RNA with alcohol. An
internal control (IC) RNA is introduced into the specimen with the specimen
diluent and serves as an amplification control for each processed specimen.
The HCV IC is an in vitro RNA transcript with primer-binding sequences iden-tical to those of the HCV target sequences, a randomized internal sequence of
similar length and base composition as the HCV target sequences, and a unique
probe-binding region that differentiates the IC from the target amplicon. The
IC is introduced into each amplification reaction and is coamplified with target
RNA from the clinical specimen.
Detection and Typing of HCV 365
The appropriate selection of primers and probes is critical to the ability
of the test to detect all the HCV genotypes. The 5vuntranslated region (UTR)
of the HCV genome is the most conserved RNA sequence among the known
HCV genotypes (12) . The AMPLICOR test uses primers KY78 and KY80
to amplify a sequence of 244 nucleotides within the highly conserved
5v UTR.
The thermostable recombinant enzyme Thermus thermophilus DNA poly-merase (rTth) has both RT and DNA polymerase activity in the presence of
manganese (13) . The processed specimens are added to the amplification mix-ture in reactions tubes in which both reverse transcription and PCR amplifica-tion occur. The antisense primer (KY78) is biotylinated at the 5vend and the
sense primer (KY80) is not. The reaction mixture is heated to allow the
antisense primer to anneal to the HCV and IC target RNA. In the presence of
excess deoxynucleoside triphosphates (dNTPs), rTth polymerase extends the
annealed primer forming cDNA. Following reverse transcription of the target
RNA, the reaction mixture is heated to denature the RNA cDNA hybrids and
expose the primer-binding sequences. As the mixture cools, the sense primer
binds to the cDNA strand, and the rTth polymerase extends the sense primer to
synthesize a second DNA strand. This completes the first cycle of PCR, yield-ing a double-stranded DNA (dsDNA) copy of the target RNA. The reaction
mixture is heated again to separate the dsDNA and expose the primer-binding
regions. As the mixture cools, the primers, KY78 and KY80, anneal to the
target DNA. rTth extends the annealed primers along the target templates to
produce a 244-bp dsDNA product. This process is repeated for 37 cycles, each
cycle effectively doubling the amount of amplicon DNA.
Selective amplification of target nucleic acid from the clinical specimen is
achieved by the use of uracil-N-glycosylase (UNG) (AmpErase). UNG recog-nizes and catalyzes the destruction of DNA strands containing deoxyuridine,
but not DNA containing thymidine (14) . Deoxyuridine is not present in natu-rally occurring DNA, but it is always present in amplicon owing to the use
deoxyuridine triphosphate as one of the dNTPs in the master mix. Therefore,
only amplicon contains deoxyuridine. UNG, which is included in the master
mix, catalyzes the cleavage of deoxyuridine containing DNA at deoxyuridine
residues by opening the deoxyribose chain at the C1 position. When heated in
the first thermal cycling step at the alkaline pH of the master mix, the amplicon
DNA strand breaks at the position of the deoxyuridine residue, rendering the
DNA nonamplifiable. UNG is not active above 55°C (i.e., throughout the ther-mal cycling steps) and therefore does not destroy newly synthesized amplicon.
Following amplification, any residual enzyme is denatured by the addition of
the denaturation solution. The use of UNG in the AMPLICOR test helps pre-vents false-positive results owing to crosscontamination with amplicon.
366 Nolte
Following amplification, the HCV and IC amplicons are chemically dena-tured to form single-stranded DNA by the addition of denaturation solution,
and aliquots of the denatured amplicon are added to microwell plates coated
with HCV-specific (KY150) and IC-specific (SK535) oligonucleotide probes,
respectively. This hybridization increases the overall specificity of the test.
Following hybridization, the microwell plates are washed to remove any
unbound material, and an avidin-horseradish peroxidase (Av-HRP) conjugate
is added to each well of the microwell plates. The Av-HRP binds to the biotin-labeled amplicon captured by the plate-bound oligonucleotide probe. The
microwell plates are washed again to remove unbound Av-HRP, and a sub-strate solution containing H2O2 and 3,3v,5v,5v-tetramethylbenzidine (TMB) is
added to the microwell plates. In the presence of H2O2, the bound HRP cata-lyzes the oxidation of TMB to form a color complex. The reaction is stopped
by the addition of a weak acid, and the optical density (OD) at 450 nm is mea-sured using an automated microwell plate reader.
The six major genotypes and subtypes of HCV vary in their worldwide dis-tribution. In the United States, type 1 is predominant (75%) and types 2, 3, and
4 comprise the remaining 25% (15 ,16) . Several studies suggest that genotypes
other than 1 are more likely to be associated with a sustained response to inter-feron alone or in combination with ribavirin (17 ,18) . Although treatment should
not be withheld from patients based on genotype of the virus, this information
can help in advising patients on the likelihood of response to treatment and in
determining the duration of therapy (18) .
Various methods have been used for HCV genotyping, including genomic
amplification and sequencing  (19–22) , PCR with genotype-specific primers
(23 ,24) , restriction fragment-length polymorphism of PCR amplicons (25) , dif-ferential hybridization  (26) , allele-specific oligonucleotide hybridization of
PCR amplicon  (27) , cleavase fragment length polymorphism  (28) , PCR
amplicon heteroduplex tracking assay, and detection of genotype-specific anti-bodies by immunoenzymatic methods  (29 ,30) . Genomic amplification and
sequencing, followed by sequence comparison and phylogenetic tree construc-tion for confirmation, is currently the “gold standard.” Genomic regions com-monly used for this approach include nonstructural region 5, envelope 1 region,
and core region. This method is costly, labor-intensive, not standardized from
laboratory to laboratory, but does lend itself to large-scale testing.
A line-probe assay (LiPA) employing reverse hybridization of PCR
amplicon to type-specific probes on nitrocellulose strips is available commer-cially (INNO-LiPA HCV II; Innogenetics, Norcross, GA) for HCV genotyping
(27) . The LiPA is based on variations found in the 5v UTR of the different
HCV genotypes. It is a simple and quick procedure that has compared favor-ably with more cumbersome procedures in several recent evaluations (31–33) .
Detection and Typing of HCV 367
Because the LiPA works well with the amplicon from the AMPLICOR
HCV test, laboratories could detect HCV and determine its genotype from a
single PCR.
2. Materials
2.1. Detection of HCV RNA Using
the AMPLICOR HCV Test, Version 2.0
2.1.1. Collection of Specimens
The AMPLICOR HCV test is for use only with serum or plasma specimens.
For serum samples, collect blood in VACUTAINER serum separator tubes
(Becton-Dickinson no. 367784 or 367789), and for plasma, collect blood in
tubes containing EDTA (lavender top, Becton-Dickinson no. 6454 or equiva-lent) or acid citrate dextrose (yellow top, Becton Dickinson no. 4606 or equiva-lent) as the anticoagulant. Specimens collected in tubes containing heparin as
the anticoagulant are unsuitable for this test.
2.1.2. Preparation of Specimens
The manufacturer provides items 1–6. The composition of several reagents
is proprietary.
1. Lysis reagent: 68% guanidinium thiocyanate, 3% dithiothreitol, glycogen, Tris-HCl buffer. Warm to 25–37°C and mix thoroughly prior to use.
2. Specimen diluent: synthetic poly rA RNA, EDTA, sodium azide, Tris-HCl buffer.
3. Negative human plasma: nonreactive by antibody tests for HCV, human immu-nodeficiency virus-1 (HIV-1) and HIV-2, and HBsAg; ProClin 300 (Rohm and
Haas).
4. Positive control: noninfectious in vitro transcribed RNA containing HCV
sequences, synthetic poly rA RNA, EDTA, sodium azide.
5. Negative control: synthetic poly rA RNA, EDTA, sodium azide.
6. Internal control: noninfectious in vitro transcribed RNA containing HCV primer-binding sequences and a unique probe-binding region, synthetic poly rA RNA,
EDTA, sodium azide (see Note 1).
7. 70% Ethanol: Add 1 vol of deionized water to 2.75 vol of 95% ethanol. Prepare
fresh daily.
8. Isopropyl alcohol.
2.1.3. Reverse Transcription and cDNA Amplification
All reagents are provided by the manufacturer.
1. Master mix: primers KY78 and KY80; dATP, dCTP, dGTP, and dUTP; rTth DNA
polymerase; UNG; glycerol; potassium acetate; dimethyl sulfoxide; bicine buffer;
sodium azide (see Note 2).
2. Manganese solution: manganese, acetic acid, amaranth dye, sodium azide.
368 Nolte
2.1.4. Detection of Amplicon
All reagents are provided by the manufacturer.
1. HCV DNA probe-coated microwell plate: twelve 8-well strips in a resealable
pouch with desiccant.
2. Internal control probe-coated microwell plate: twelve 8-well strips in a resealable
pouch with desiccant.
3. Denaturation solution: 1.6% NaOH, EDTA, thymol blue.
4. Hybridization buffer: sodium phosphate, sodium thiocynate, solubilizer.
5. Av-HRP conjugate: Av-HRP conjugate, bovine a-globulin, Emulsit 25 (Dai-ichi
Kogyo Seiyaku); 0.1% phenol, 1% ProClin 150 (Rohm and Haas).
6. Substrate A: citrate solution, 0.01% H2O2; 0.1% ProClin 150.
7. Substrate B: 0.1% TMB, 40% N,N-dimethylformamide (DMF).
8. 10X Wash concentrate: phosphate buffer, NaCl, EDTA, detergent, 0.5%
ProClin 300.
9. Stop reagent: 4.9% sulfuric acid.
2.2. Genotyping HCV Using LiPA (INN0-LiPA HCV II)
The most convenient source of amplicons for the LiPA assay is the PCR
products from the AMPLICOR HCV 2.0 assay. Alternatively, RT-PCR of HCV
can be performed using primers supplied by Innogenetics. The following pro-tocol uses AMPLICOR HCV 2.0 PCR products.
2.2.1. Line-Probe Assay
The following reagents are supplied by the manufacturer. The detailed com-position of these reagents is proprietary.
1. Nitrocellulose strips with 19 type-specific probes bound to the surface thorough
poly-T tails (Fig. 1).
2. Alkaline denaturation solution.
3. Hybridization solution containing saline sodium citrate (SSC) buffer with 0.1%
sodium dodecyl sulfate (SDS).
4. Stringent wash solution containing SSC buffer with 0.1% SDS.
5. Concentrated conjugate: streptavidin labeled with alkaline phosphatase. Prior to
use, dilute 1 100 conjugate diluent.
6. Concentrated 5-bromo-5-chloro-3-indolyl phosphate `-toluidine salt (BCIP) and
nitroblue tetrazolium chloride (NBT) substrate solution in DMF. Dilute 1 100 in
substrate buffer before use.
7. Conjugate diluent containing phosphate buffer containing NaCl, Triton, protein
stabilizers, and sodium azide.
8. Substrate buffer containing Tris buffer with NaCl and MgCl2.
9. Concentrated rinse solution: phosphate buffer with NaCl, Triton, and sodium
azide. Dilute 1 5 in distilled water before use.
10. Incubation trays containing eight troughs.
11. Transparent plastic reading chart for identification of positive lines.
Detection and Typing of HCV 369
The following items are not provided by the manufacturer and must be sup-plied by the user.
1. Shaking water bath, capable of 80 rpm, with an inclined lid and temperature
adjustable to a minimum of 50°C.
2. Liquid aspirator.
3. Tweezers.
4. Vortex mixer.
5. Pipets adjustable to deliver 1–20, 20–200, and 200–1000 µL.
6. Orbital shaker or rocker shaker capable of 160 and 50 rpm, respectively.
Fig. 1. Type- and subtype-specific probe location on the INNO-LiPA HCV II strip.
370 Nolte
3. Methods
3.1. Amplicor HCV Assay
3.1.1. Collection, Transport, and Storage of Specimens
1. Whole blood should be kept at 2–25°C and serum or plasma should be separated
within 6 h of collection to avoid degradation of HCV RNA. A minimum of
200 µL of serum or plasma is required for the test.
2. Serum or plasma may be transported at 2–8°C or frozen at –20 to –80°C.
3. Aliquot serum or plasma into appropriately labeled 1.5-mL polypropylene screw-cap tubes (Sarstedt cat. no. 72.692.105).
3.1.2. Preparation of Specimens
1. Label one 1.5-mL polypropylene screw-cap tube for each patient specimen and
kit control.
2. Prepare the working lysis reagent by adding 100 µL of IC to one bottle of lysis
reagent.
3. Add 400 µL of working lysis reagent to each labeled tube and cap.
4. Prepare plasma controls as follows: To each of three 1.5-mL tubes (two negative and
one positive control tubes) add 200 µL of negative plasma. Vortex for 3–5 s. Then
add 25 µL of the negative control to each of the two negative control tubes and 25 µL
of the positive control to the positive control tube. Vortex the control tube tubes for 3–5 s.
5. Vortex the patient specimens for 3–5 s. Add 200 µL of specimen to the appropri-ate tube containing the working lysis reagent and vortex again.
6. Incubate all the specimens and control tubes for 10 min at 60°C and vortex.
7. Add 600 µL of isopropyl alcohol (at room temperature) to each tube and vortex.
8. Incubate all the tubes for 2 min at room temperature.
9. Put a mark on each tube and place the tubes into the microcentrifuge with the
orientation mark facing outward, so that the resulting pellet will align with the
orientation mark. Centrifuge for 15 min at 13,000–16,000g at room temperature.
10. Using a new, fine-tip, disposable transfer pipet for each tube, carefully remove
and discard the supernatant from each tube, being careful not to disturb the pellet.
The pellet may not be visible but should be aligned with the orientation mark.
11. Add 1.0 mL of 70% ethanol to each tube and vortex.
12. Centrifuge the tubes for 5 min as in step 9. Use the orientation mark on the tubes
to align the pellet.
13. Remove and discard the supernatant as in step 10.
14. Pulse centrifuge the tubes for 5 s.
15. Using a P200 pipet fitted with a plugged tip, remove all the residual supernatant
from each tube.
16. Pipet 200 µL of specimen diluent into each tube. Resuspend the pellet by scrap-ing the pellet from the side of the tube using a plugged pipet tip and then vortex.
Let the particulate matter settle for 5–10 s. The processed specimens and controls
can be held at room temperature for up to 3 h or for up to 1 mo at –20 to –80°C
prior to amplification (see Note 3).
Detection and Typing of HCV 371
3.1.3. Reverse Transcription and cDNA Amplification
1. Place the appropriate number of MicroAmp tubes for patient and control testing
in a MicroAmp tray and lock in place with the retainer.
2. Prepare the working master mix by adding 100 µL of manganese solution to one
tube of master mix.
3. Recap the master mix tube and mix well by inverting the tube 10–15 times. Dis-card the remaining manganese solution.
4. Pipet 50 µL of the working master mix solution into each MicroAmp reaction
tube using an Eppendorf repeater pipet and an individually wrapped sterile
1.25-mL Combitip reservoir or a micropipet with a plugged tip. One tube of mas-ter mix is enough for 12 reactions.
5. Place the MicroAmp tray containing the master mix in a resealable plastic bag
without capping the tubes. Seal the bag securely and store the sample tray at
2–8°C until specimen preparation is completed. Amplification must begin within
4 h of preparation of the master mix.
6. Pipette 50  µL of each processed specimen and control into the appropriate
MicroAmp reaction tube containing the working master mix.
7. Cap the MicroAmp reaction tubes and move to the amplification and detection
area.
8. Place the tray/retainer assembly into the thermal cycler sample block.
9. Program the thermal cycler as follows: hold program, 5 min at 50°C; hold pro-gram, 30 min at 62°C; cycle program (37 cycles), 10 s at 90°C, 25 s at 58°C; hold
program, 91°C for a maximum of 3 h.
10. Start the thermal cycle program. The program runs approx 1 h and 45 min. Speci-mens must be removed within 3 h of the start of the final hold program.
3.1.4. Detection of Amplicon
1. Remove the MicroAmp caps carefully to avoid creating aerosols of amplicon.
Immediately pipet 100  µL of the denaturation solution into the reaction tubes
using a multichannel pipettor. Mix by pipetting up and down. Denatured amplicon
can be held at room temperature for no longer than 2 h or at 2–8°C for up to 1 wk.
2. Add 100 µL of hybridization buffer to each well to be tested on the HCV and IC
microwell plates.
3. Pipet 25 µL of each denatured amplicon to the appropriate wells of the HCV and
IC microwell plates.
4. Cover the microwell plates and incubate for 1 h at 37°C.
5. Wash the microwell plates five times with 1X wash solution using a microwell
plate washer.
6. Add 100 µL of av-HRP conjugate to each well. Cover the microwell plates and
incubate for 15 min at 37°C.
7. Wash the microwell plates five times with 1X wash solution using a microwell
plate washer.
8. Prepare the working substrate reagent by mixing 2.0 mL of substrate A and
0.5 mL of substrate B for each of the 16 tests, no earlier than 3 h prior to use.
372 Nolte
9. Pipet 100 µL of the working substrate reagent into each well being tested.
10. Allow color development for 10 min at room temperature in the dark.
11. Add 100 µL of stop reagent to each well.
12. Measure the OD at 450 nm within 30 min of adding the stop reagent. Record the
absorbance value for each patient specimen and control tested.
3.1.5. Interpretation of Results
The assay results for the HCV positive and HCV negative controls should
be *1.5 A450 and <0.25 A450, respectively. If the absorbance values are <1.5 for
the positive control and *0.25 for the negative controls, the run is invalid and
the entire test procedure should be repeated. The presence of HCV RNA in a
specimen is determined by comparing the absorbance at 450 nm of the
unknown specimen to the established cutoff values for the test. The test result
interpretation is based on the combination of the HCV and IC results as shown
in Table 1. If invalid negative test results are obtained, either another aliquot
of the original specimen should be processed or a new specimen should be
collected. If equivocal test results are obtained, repeat the entire test procedure
in duplicate using a new aliquot of the specimen. If both repeat test results are
*0.3, the specimen should be considered positive. Specimens with one or both
repeat test results that are <0.3 should be considered negative. The analytical
sensitivity of this test is difficult to determine because of the lack of a recog-nized “gold standard” HCV RNA preparation. In our hands, it is at least as sen-sitive as our in-house-developed assay at )100 copies/m  (34) . Recent progress
toward establishing international units for HCV RNA preparations should help
to determine the relative sensitivities of the various nucleic acid amplification
assays currently in use. The PCR products from the AMPLICOR assay may be
used in the LiPA assay (35) .
3.2. LiPA HCV II
3.2.1. Denaturation and Hybridization (see Note 4)
1. Heat a water bath to 50°C and prewarm the hybridization solution to at least
37°C but not more than 50°C. Mix the hybridization solution before use to dis-solve any crystals.
2. Remove the required number of strips and write an identification number above
the black line on the strips with a pencil.
3. Place one trough per test in the plastic tray.
4. Pipet 10 µL of denaturation solution into the upper corner of each trough.
5. Add 10 µL of the PCR product to the denaturation solution and carefully mix by
pipetting up and down. Allow denaturation to proceed for 5 min at room temperature.
6. Add 2 mL of hybridization solution to the denatured PCR product in each trough.
Take care not to contaminate adjacent troughs during pipetting.
Detection and Typing of HCV 373
7. Immediately place the strip with the black marker line side up into the trough.
The strips must be completely submerged in the solution.
8. Place the tray into the 50°C shaking water, set the speed at 80 rpm, and incubate
for 60 min.
3.2.2. Stringent Wash
1. Remove the tray from the water bath, hold the tray at a low angle, and aspirate the
liquid from the trough.
2. Add 2 mL of prewarmed stringent wash solution into each trough and rinse by
rocking the tray for 10–20 s at room temperature.
3. Repeat step 2 once more.
4. Incubate each strip in 2 mL of prewarmed stringent wash solution in the shaking
water bath at 50°C for 30 min.
3.2.3. Color Development
1. Wash each strip twice for 1 min using 2 mL of the diluted rinse solution.
2. Add 2 mL of diluted conjugate to each trough and incubate for 30 min while
agitating the tray on the shaker.
3. Wash each strip twice for 1 min with 2 mL of diluted rinse solution and wash
once more using 2 mL of substrate buffer.
4. Add 2 mL of substrate solution to each trough and incubate for 30 min while
agitating the tray on the shaker.
5. Stop the color development by washing the strips in 2 mL of deionized water
with agitation on the shaker for at least 5 min.
6. Using tweezers, remove the strips from the troughs and place them on absorbent
paper. Let the strips dry completely before reading the results. Store the devel-oped strips in the dark.
3.2.4. Interpretation of Results
After hybridization, streptavidin labeled with alkaline phosphatase is added
and binds to any biotinylated hybrid previously formed. Incubation with BCIP/
NBT chromogen results in a purple/brown precipitate. Consequently, a col-Table 1
Interpretation of AMPLICOR HCV Test, Version 2.0 Results
HCV result (A450) IC result (A450)   Interpretation
<0.3 *0.3 HCV RNA not detected
<0.3 <0.3 Invalid negative test result
*1.0 Any HCV RNA detected
*0.3, <1.0 Any Equivocal
374 Nolte
374
Table 2
INNO-LiPA HCV II Interpretation Chart
*This pattern is found with some isolates from Indonesia and is provisionally classified as 10a.
Detection and Typing of HCV 375
ored precipitate forms only when there is a perfect match between the probe
and the PCR products. A line is considered positive when a distinct purple/
brown band appears at the end of the test procedure.
The first line on the strip is the conjugate control line that controls for the
color development steps in the procedure. It should be lined up with the conju-gate control line on the transparent plastic reading strip. The second positive
line controls the addition of amplified product for hybridization. This line
should always be positive if the amplified product form of HCV is present.
Inhibition of PCR might be the reason for complete failure of the genotyping
test. Record all the line numbers that are positive on the strip and deduce the
HCV genotype using the interpretation chart (see Table 2). Coinfection with
different types of HCV may be recognized when lines from two different
major types are present.
The INNO-LiPA HCV II test is based on nucleotide variability of the
5vUTR, and the test yields high concordance with sequences obtained for this
region using nucleic acid sequencing techniques. However, subtyping may
occasionally not be possible owing to lack of subtype-specific sequence varia-tion in the 5v UTR.
The INNO-LiPA is effective in identifying HCV genotypes that occur most
frequently in North America, Europe, and Japan. However, DNA sequencing
is currently the best means of discriminating types endemic to other geographic
regions.
4. Notes
1. The IC control not only tests for inhibition of the amplification reaction but also
tests for RNA recovery. Therefore, a failed IC reaction may indicate the presence
of inhibitors in the specimen or poor recovery of RNA from the specimen. In our
experience, most failed IC reactions are resolved by simply processing another
aliquot of the original specimens and repeating the test.
2. The presence of UNG and deoxyuridine triphosphate in the PCR master mix
reduces the risk of amplicon contamination. UNG has been demonstrated to inac-tivate a least 1000 copies of deoxyuridine-containing HCV amplicon per PCR
(see Roche AMPLICOR HCV Test, version 2.0 package insert). However, con-tamination from HCV positive controls and clinical specimens can be avoided
only by good laboratory practices and careful adherence to procedure.
3. Recovery of the often invisible RNA pellet is the most challenging part of this
procedure. Careful attention to this step is critical to achieving reproducible
results (34) .
4. The hybridization, stringent washing, and color development steps have been
automated in the Auto-LiPA instrument.
376 Nolte
References
1. Choo, Q. L., Kuo, G., Weiner, A. J., Overby, L. R., Bradley, D. W., and Houghton,
M. (1989) Isolation of a cDNA clone derived from a blood-borne non-A non-B
viral hepatitis genome. Science 224, 359–362.
2. Alter, H. J. (1991) Descartes before the horse: I clone, therefore I am: the hepatitis
C virus in current perspective. Ann. Intern. Med. 115, 644–649.
3. Kuo, G., Choo, Q. L., Alter, H. J., et al. (1989) An assay for circulating antibodies
to a major etiologic virus of human non-A, non-B hepatitis.  Science 244,
362–364.
4. Simmonds, P., Alberti, A., Alter, H. J., et al. (1994) A proposed system
for the nomenclature of hepatitis C viral genotypes. Hepatology 19, 1321–1324
(letter).
5. Miller, R. H. and Purcell, R. H. (1990) Hepatitis C virus shares amino acid
sequence similarities with pestiviruses and flaviviruses as well as members of two
plant virus supergroups. Proc. Natl. Acad. Sci. USA 87, 2057–2061.
6. Ogata, N., Alter, H. J., Miller R. H., and Purcell, R. H. (1991) Nucleotide
sequence and mutation rate of the H strain of hepatitis C virus. Proc. Natl. Acad.
Sci. USA 88, 3391–3396.
7. Gretch, D. R. (1997) Diagnostic tests for hepatitis C. Hepatology 26, 43S–47S.
8. Daman, M., Cuypers, H. T. M., Zaaijer, H. L., Reesink, H. W., Schaasberg, W. P.,
Gerlich, W. H., Miesters, H. G. M., and Lelie, P. N. (1996) International collabo-rative study on the second EUROHEP HCV-RNA reference panel. J. Virol. Meth-ods 58, 175–185.
9. Young, K. K. Y., Resnick, R. M., and Myers, T. W. (1995) Detection of hepatitis
C virus RNA by a combined reverse transcription-polymerase chain reaction
assay. J. Clin. Microbiol. 31, 882–886.
10. Zeuzem, S., Ruster, B., and Roth, W. K. (1994) Clinical evaluation of a new poly-merase chain reaction assay (AmplicorTM HCV) for detection of hepatitis C virus.
Z. Gastroenterol. 32, 342–347.
11. Nolte, F. S., Thurmond, C., and Fried, M. W. (1995) Preclinical evaluation of
AMPLICOR hepatitis C virus test for detection of hepatitis C virus RNA. J. Clin.
Microbiol. 33, 1775–1778.
12. Bukh, J., Purcell, R. H., and Miller, R. H. (1992) Importance of primer selection
for the detection of hepatitis C virus RNA with the polymerase chain reaction.
Proc. Natl. Acad. Sci. USA 89, 187–191.
13. Myers, T. W. and Gelfand, D. H. (1991) Reverse transcription and DNA amplifi-cation by a thermus thermophilus DNA polymerase. Biochemistry 30, 7661–7666.
14. Longo, M. C., Berninger, M. S. and Hartley, J. L. (1990) Use of uracil DNA
glycosylase to control carry-over contamination in polymerase chain reactions.
Gene 93, 125–128.
15. Mahaney, K., Tedeschi, V., Maertens, G., Di Bisceglia, A., Vergalia, J.,
Hoofnagle, J., and Sallie, R. (1994) Genotypic analysis of hepatitis C virus in
American patients. Hepatology 20, 1405–1411.
Detection and Typing of HCV 377
16. Lau, J. Y. N., Mizokami, M., Kolberg, J. A., et al. (1995) Application of six hepa-titis C virus genotyping systems to sera from chronic hepatitis C patients in the
United States. J. Infect. Dis. 171, 218–289.
17. McHutchison, J. G., Gordon, S., Schiff, E. R., et al. (1998) Interferon alpha-2b
alone or in combination of ribavirin as initial treatment for chronic hepatitis C.
N. Engl. J. Med. 339, 1485–1499.
18. Poynard, T., Marcellin, P., Lee, S. S., et al. (1998) Randomised trial of interferon
_2b plus ribavirin for 48 weeks or for 24 weeks versus interferon _2b plus pla-cebo for 48 weeks for treatment of chronic infection with hepatitis C virus. Lancet
352, 1426–1432.
19. Cha, T. A., Beall, E., Irvine, B., et al. (1992) At least five related, but distinct,
hepatitis C viral genotypes exist. Proc. Natl. Acad. Sci. USA 89, 7144–7148.
20. Chan, S. W., McOmish, F., Holmes, E. C., et al. (1992) Analysis of a new hepati-tis C virus type and its phyylogenetic relationship to existing variants.  J. Gen.
Virol. 73, 1131–1141.
21. Bukh, J., Purcell, R. H., and Miller, R. H. (1993) At least 12 genotypes of hepati-tis C virus predicted by sequence analysis of the putative E1 gene of isolates col-lected worldwide. Proc. Natl. Acad. Sci. USA 90, 8234–8238.
22. Simmonds, P., Holmes, E. C., Cha, T.-A., et al. (1993) Classification of hepatitis
C virus into six major genotypes and a series of subtypes by phylogenetic analysis
of the NS-5 region. J. Gen. Virol. 74, 2391–2399.
23. Okamoto, H., Sugiyama, Y., Okada, S., et al. (1992) Typing hepatitis C virus by
polymerase chain reaction with type-specific primers: application to clinical sur-veys and tracing infectious sources. J. Gen. Virol. 73, 673–679.
24. Chayama, K., Tsubota, A., Arase, Y., et al. (1993) Genotypic subtyping of hepa-titis C virus. J. Gastroenterol. Hepatol. 8, 150–156.
25. Nakao, T., Enomoto, N., Tadada, N., Tadada, A., and Date, T., (1991) Typing of
hepatitis C virus genomes by restriction fragment length polymorphism. J. Gen.
Virol. 72, 2105–2112.
26. Takada, N., Takase, S., Enomoto, N., Takada, A., and Date, T. (1992) Clinical
backgrounds of the patients having different types of hepatitis C virus genomes.
J. Hepatol. 14, 35–40.
27. Stuyver, L., Rossau, R., Wyseur, A., Duhamel, M., Vanderborght, B., Van
Heuverswyn, H., and Maertens, G. (1993) Typing of hepatitis C virus isolates
and characterization of new subtypes using a line probe assay. J. Gen. Virol. 74,
1093–1102.
28. Marshall, D. J., Heisler, L. M., Lyamichev, V., Murvine, C., Olive, D. M., Ehrlich,
G. D., Neri, B. P., and deArruda, M. (1997) Determination of hepatitis C virus
genotypes in the United States by cleavase fragment length polymorphism analy-sis. J. Clin. Microbiol. 35, 3156–3162.
29. Simmonds, P., Rose, K. A., Graham, S., et al. (1993) Mapping of serotype-specific immunodominant epitopes in the HS-4 region of hepatitis C virus (HCV).
Use of type-specific peptides to serologically differentiate infections with HCV
type 1, type 2 and type 3. J. Clin. Microbiol. 31, 1493–1503.
378 Nolte
30. Bhattacherjee, V., Prescott, L. E., Pike, I., et al. (1995) Use of NS-4 peptides to
identify type-specific antibody to hepatitis C virus genotypes 1, 2, 3, 4, 5 and 6.
J. Gen. Virol. 76, 1737–1748.
31. Andonov, A. and Chaudhary, R. K. (1995) Subtyping of hepatitis C virus isolates
by a line probe assay using hybridization. J. Clin. Microbiol. 33, 254–256.
32. Lau, J. Y. N., Simmonds P., and Urdea, M. S. (1995) Implications of variations of
“conserved” regions of hepatitis C virus genome. Lancet 346, 425, 426.
33. Hadziyannis, E., Larsen, N., Thurmond, C., Mauer, D., and Nolte, F. (1997)
Genotyping of hepatitis C virus with direct sequencing: comparison with a line
probe assay, in Program and Abstracts of the Annual Meeting of the Association
of Molecular Pathology, San Diego, CA.
34. Nolte, F. S., Thurmond, C., and Mitchell, P. S. (1994) Isolation of hepatitis
C virus RNA from serum for reverse transcription-PCR. J. Clin. Microbiol. 32,
519,520.
35. Hadziyannis, E., Fried, M., and Nolte, F. S. (1995) Evaluation of two methods for
quantitation of hepatitis C virus RNA, in  Program and Abstracts of the 46th
Annual Meeting of the American Association for the Study of Liver Diseases,
Chicago, p. 356A.
Detection and Speciation of Mycobacteria 379
29
Detection and Speciation of Mycobacteria
in Formalin-Fixed, Paraffin-Embedded
Tissue Sections
Diana Mohl and Thomas J. Giordano
1. Introduction
The ability to detect mycobacterial DNA by polymerase chain reaction
(PCR)–based methodology in formalin-fixed, paraffin-embedded tissue sec-tions is useful in several clinical scenarios. The major use of this type of assay
is in those instances in which infectious disease is not clinically suspected and
microbiological cultures are not performed. In these cases, the only tissue avail-able for examination is that present in routinely prepared paraffin blocks after
histologic examination. The presence of necrotizing granulomatous inflamma-tion should result in special stains for acid-fast organisms. However, in many
such cases the special stains are unsatisfactory, because the number of organ-isms present is very low (as in Mycobacterium tuberculosis). Thus, the tedious
examination of multiple serial sections is often necessary to identify the patho-genic organism, and often no organism is found. Therefore, more sensitive
detection methods are needed. PCR-based detection of mycobacterial DNA is
more sensitive and can be used either to verify the presence of organisms seen
on special stains or to identify an occult organism. By combining the PCR
assays with restriction analyses of the products, it is often possible to speciate
the pathogenic organism.
In addition, given the rapidity of the PCR assays compared to mycobacterial
culture, this technology can be used whenever mycobacterial disease is suspected,
but the clinical situation precludes waiting for culture results. In these cases,
often the determination of typical vs atypical organisms is all the information one
needs to provide until culture confirmation and speciation becomes available.
379
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
380 Mohl and Giordano
A variety of assays has been employed to detect mycobacterial DNA in
formalin-fixed, paraffin-embedded tissues (1,2) . Most have used a nested PCR
approach to amplify the 65-kDa surface antigen gene (3) , the IS6110 insertion
sequence (4–6) , the mtp40 sequence  (7) , or the 16S rRNA gene  (8 ,9) . The
assays have been shown to be rapid, specific, and sensitive. The assay described
herein is based on that of Cook et al.  (10) and employs nested PCR of the
65-kDa surface antigen gene followed by an algorithmic restriction analysis
for speciation.
2. Materials
2.1. Slides
1. One positive control slide (see Note 1).
2. Four to six slides of the case (see Note 2).
2.2. DNA Extraction
1. 1.5-mL Microfuge tubes.
2. Xylene.
3. Razor blades, single edge.
4. 100% Ethanol.
5. 70% Ethanol.
6. Proteinase K (20 mg/mL) (Gibco-BRL, Gaithersburg, MD).
7. 50 mM Tris-HCl, pH 8.3.
8. Dry ice.
2.3. Polymerase Chain Reaction
1. 0.5-mL Microfuge tubes (or 0.2 mL, depending on the type of thermal cycler).
2. Taq DNA polymerase and buffer with MgCl2 (Gibco-BRL).
3. dNTPs (10 mM) (Gibco-BRL).
4. Sterile H2O.
5. Oligonucleotides (1 µg/mL stocks). Sequences are shown in Table 1.
2.4. Analysis of PCR Products
1. NuSieve agarose (FMC Bioproducts, Rockland, ME).
2. Agarose, multipurpose (Gibco-BRL).
3. DNA size markers, e.g., phi-X174/HaeIII digest (Gibco-BRL).
4. 10X Gel-loading buffer (Gibco-BRL).
5. 10X Tris-borate EDTA (TBE) buffer (Gibco-BRL).
6. Ethidium bromide (10 mg/mL stock).
2.5. Restriction Analysis of PCR Products
1. Restriction enzymes BstUI, CfoI, MboI (Gibco-BRL).
2. Reaction buffer provided by manufacturer.
3. Mineral oil (Sigma, St. Louis, MO).
Detection and Speciation of Mycobacteria 381
2.6. Equipment
1. Thermal cycler.
2. Pipets with filtered pipet tips.
3. Agarose gel apparatus.
4. Tabletop centrifuge.
5. Tube racks.
6. Shaking incubator.
7. Heat blocks (37 and 60°C).
8. Ice bath.
3. Methods
The following methods are those of Cook et al.  (10) with only minor
modifications.
3.1. DNA Extraction
1. Label the appropriate number of 1.5-mL microfuge tubes with the date and
sample number (see Note 3).
2. Pipet 100 µL of xylene into each tube.
3. Using a clean razor blade, scrape the tissue off the glass slide.
4. Place the tissue into the tube containing the xylene and tap to make sure the tissue
is submerged.
5. Stand at room temperature for 5–10 min to deparaffinize the tissue.
6. Spin the xylene and tissue at 12,800g in a microcentrifuge for 5 min (see Note 4).
7. Remove the supernatant with a pipet, making sure the tissue pellet remains in the
tube (see Note 4). Blot the excess xylene with a tissue.
Table 1
Sequences of PCR Primers
Round 1: mycobacteria primers
5v-AAGGAGATCGAGCTGGAGGA-3v (upstream 1)
5v-AGGCGTTGGTTCGCGAGGG-3v (upstream 2)
5v-TGATGACGCCCTCGTTGCC-3v (downstream)
Round 2: mycobacteria primersa
5v-GTCTCAAACGCGGCATCG-3v (upstream)
5v-GTCACCGATGGACTGGTC-3v (downstream)
Round 1: `-globin primers
5v-CCATAGGCAGAGAGAGTCAGTG-3v (upstream 1)
5v-TGCCAGAAGAGCCAAGGACAGG-3v (downstream 1)
Round 2: `-globin primersa
5v-CACATAGGCAGAGAGCGTCAGTG-3v (upstream)
5v-CCTATCAGAAACCCAAGAGTC-3v (downstream)
a Round 2 product sizes are 130 and 322 bp for mycobacteria and `-globin, respectively.
382 Mohl and Giordano
8. Add 100 µL of 100% ethanol to the tissue pellet. Gently tap to resuspend.
9. Spin at 14,000 rpm in a tabletop centrifuge for 5 min.
10. Pour off the supernatant and remove the excess liquid with a Kimwipe (see
Note 4).
11. Repeat steps 8–10 with 70% ethanol.
12. Place the open tubes in a rack and heat in a 60°C oven for 15 min or until the
pellets are dry.
13. Prepare fresh proteinase K digestion solution (0.2 mg/mL in 50 mM Tris-HCl,
pH 8.3) from a stock solution of 20 mg/mL of proteinase K.
14. Add 100 µL of the proteinase K digestion solution to each tube.
15. Tap the tubes to dissolve the pellet quick-spin.
16. Incubate for 6 h to overnight at 37°C in a shaking incubator.
17. Quick-spin the tubes to collect the liquid at the bottom of the tubes.
18. Place the tubes in a dry ice–ethanol bath for 1 min. Remove the tubes and place
them into a boiling water bath for 8 min to inactivate the proteinase K.
19. Place the tubes on ice for 5 min, and then spin at 12,800g in a tabletop centrifuge
for 5 min.
20. Transfer the supernatant, which contains the DNA, to a clean, labeled tube.
Alternatively, the DNA can be stored in the same tube; spin again before remov-ing the DNA.
3.2. Polymerase Chain Reaction
1. Prepare PCR master mix for round 1. The master mix contains dNTPS, 1X Taq
buffer with MgCl2, and oligonucleotide primers. It is prepared in large batches,
aliquoted, and stored at –20°C. The master mix for round 1 (makes mix for 200
reactions) contains the following (see Note 5): 25 µL of mycobacteria primers
(three each at 1 µg/µL), 2.5 µL of `-globin primers (two at 1 µg /mL), 20 µL of
10 mM dNTPs, 1000 µL of 10X Taq buffer with MgCl2, and 2540 µL of H2O
(total of 3700 µL/200 rxn = 18.5 µL/rxn).
2. Label 0.5-mL tubes with the date and sample number.
3. In a 1.5-mL tube, add 18.5 µL of master mix for each reaction plus 1. Thus, if
there are nine reactions, add 185 µL (9 + 1 = 10 × 18.5 µL = 185 µL).
4. To the tube with the master mix, add 0.5 µL of Taq DNA polymerase (5 U/µL)
for each reaction.
5. Mix well by tapping.
6. Pipet 19 µL of the master mix with Taq into prelabeled 0.5-mL tubes.
7. Add 31 µL of the DNA to each tube. Mix well and quick-spin.
8. Place the tubes in a thermal cycler and run the following program: initial denatur-ation for 4 min at 94°C; followed by 36 cycles of 94°C for 1 min, 57°C for 2 min,
and 72°C for 2 min; followed by a 10-min extension at 72°C. Bring to 25°C and
hold at that temperature. This will take about 4 h.
9. Remove the tubes and quick-spin.
Detection and Speciation of Mycobacteria 383
10. Label a new set of 0.5 mL tubes with the same information as in round 1, but
specify them as round 2.
11. Prepare the master mix for round 2 (makes mix for 200 reactions): 25  µL of
Mycobacteria round 2 primers (two, each at 1 µg/µL) 20 µL of `-globin primers
round 2 (two, each at 1 µg/mL) 20 µL of 10 mM dNTPs 1000 µL of 10X Taq
buffer with MgCl2, and 7835 µL of H2O (total of 8900 µL).
12. In a 1.5-mL tube, add 44.5 µL of master mix for round 2 for each reaction plus 1.
Thus, if there are nine reactions, add 445 µL (9 + 1 = 10 × 44.5 µL = 445 µL).
13. To the tube with the master mix, add 0.5 µL of Taq DNA polymerase (5 U/µL)
for each reaction.
14. Mix well by tapping.
15. Pipet 45 µL of the master mix with Taq into prelabeled 0.5 mL tubes.
16. Add 5 µL of the round 1 products to the round 2 tubes (see Note 6).
17. Place the tubes in the thermal cycler and run the following program: initial dena-turation for 4 min at 94°C, followed by 36 cycles of 94°C for 1 min, 52°C for
2 min, and 72°C for 2 min. Finish with a 10-min extension at 72°C.
18. Remove the tubes and quick-spin to collect the solution at the bottom of the tubes.
19. Analyze the products on a 3% agarose gel.
3.3. Agarose Gel Electrophoresis
1. Analyze the products on a 3% agarose/0.5X TBE gel buffer in a submersible
apparatus of choice (see Note 7).
2. Use size markers of choice to cover 100–400 bp.
3. Run 10 µL of PCR products along with 2 µL of 10X gel-loading buffer.
4. Photograph using a UV light box.
5. Proceed to restriction analysis if the duplicate extractions of the case contain the
appropriate mycobacterial band (130 bp). The `-globin product (322 bp) should
be present in all reactions except the mock extraction (see Fig. 1A).
3.4. Restriction Analysis of PCR Products
1. Digest 10  µL of each PCR product with three separate restriction enzymes—
CfoI, BstUI, and MboI—according to the supplier’s recommendations. Use 5 U
of enzyme/10 µL of PCR product (see Note 8).
2. After digestions are complete, load the entire digestion onto a 3% agarose gel as
before and electrophorese until the bromophenol tracking dye has migrated
approximately three fourths the length of the gel.
3. Analyze the restriction patterns using the algorithm of Cook et al. (10) for pat-terns that match those of known mycobacterial organisms. The patterns associ-ated with the most important organisms,  M. tuberculosis and Mycobacterium
avium intracellulare, are shown in Table 2. See Figs. 1B and 2 for examples (see
Note 9).
384 Mohl and Giordano
Fig. 1. (A) Results of two rounds of nested PCR for mycobacterial DNA. The case
is extracted and amplifed in duplicate (lanes 2 and 3). Lane 4, mock extraction; lane 5,
positive mycobacterial control; lane M, DNA markers; G, 322-bp  `-globin internal
control band; M, 130-bp mycobacterial band. (B) Restriction analysis of the products
from (A). Each PCR product is digested with a panel of restriction enzymes—CfoI,
BstUI, and MboI—and the digests are analyzed as a group. Group A–C represents the
restriction digests of the PCR products of lanes 2, 3, and 5 in (A), respectively. The
arrows represent 322, 130, 120, 76, and 65 bp. This restriction pattern matches that of
M. tuberculosis. Lane M: DNA size markers; lanes 2, 5, and 8: = CfoI digest; lanes 3,
6, and 0: MboI digest; lanes 4, 7, and 10: BstUI digest.
Table 2
Restriction Enzyme Patterns for M. tuberculosis and M. avium-intracellulare
Enzyme M. tuberculosis (bp) M. avium-intracellulare (bp)
CfoI 75 + 65 130 (uncut)
BstUI 120 85–90
MboI 130 (uncut) 80–100
Detection and Speciation of Mycobacteria 385
4. Notes
1. The best positive control tissues are obtained from culture-proven patients who
have evidence of disease at the time of autopsy. Several paraffin blocks from
such a case can provide control tissues for a significant period of time. The tissue
sections of the control can be cut into batches and stored in a slide box at room
temperature.
2. The tissue sections of the case need to be cut with a clean knife and with careful
attention not to pick up any tissue bath floaters. The sections should be about
5 µm thick, should placed on plain glass slides (not treated slides), and should not
be heated. Use the tissue from multiple sections of the tissue. Tissue fixed in B5
is suboptimal and should be avoided.
3. For each clinical sample and control pair, include a “mock” extraction that con-tains only the extraction reagents. This is used to monitor contamination in the
reagents. Clinical samples should be extracted in duplicate. If more than one
block is available, it is possible to combine a section from each block into a
single tube. Always wear gloves and handle the tubes from the sides; that is, do
not touch the rims of the tubes or the cap. Use filtered pipet tips to prevent con-tamination of the pipetmen.
4. It is very easy to dislodge and lose the tissue at this step. Remove the samples as
soon as the centrifuge stops spinning. If unable to do so, spin again and remove
the xylene.
Fig. 2. Restriction analysis of a different positive mycobacterial assay. In this case,
the restriction digests do not match those of M. tuberculosis. The pattern is that of an
atypical organism, most consistent with M. avium-intracellulare. A and B represent
the restriction digests of duplicate extractions of the case and C represents the digests
of an M. tuberculosis control. Arrows represent 322, 130, and 100 bp. Lane M: DNA
size markers; lanes 2, 5, and 8: CfoI digest; lanes 3, 6, and 0: MboI digest; lanes 4, 7,
and 10: BstUI digest.
386 Mohl and Giordano
5. Two upstream primers specific to the 65-kDa surface antigen are used to increase
the number of amplifyable cases (10) . Because the assay consists of two rounds
of nested PCR, the final products are the same regardless of which first-round
primer was used in the amplification. Add a drop of mineral oil to the PCR reac-tions if the thermal cycler does not have a heated lid.
6. This is best performed in a “clean” environment, such as a “template tamer” or a
UV biological hood to minimize contamination. Care should be taken not to con-taminant any of the round 2 reactions with any round 1 products.
7. We use a mixture of multipurpose and Nusieve agarose when preparing the 3%
gels. This agarose boils over easily, so be careful when preparing the gel. Prepare
the gel with ethidium bromide by adding 6.5 µL of 10 mg/mL stock per 100 mL
of gel.
8. The restriction digests should be set up in a designated “post-PCR” room to avoid
contamination by PCR product. We use a 4-h incubation at 37°C for CfoI and
MboI and 60°C for BstUI. Add a drop of mineral oil to the BstUI digest to prevent
evaporation.
9. If the restriction patterns of the duplicate extractions of the case are the same but
do not match the algorithm of Cook et al. (10) , you may consider sequencing the
PCR product to determine the species of origin. If the patterns of the duplicate
extractions do not match each other, then contamination is likely the source of
the mycobacterial DNA.
References
1. Osaki, M., Adachi, H., Gomyo, Y., Yoshida, H., and Ito, H. (1997) Detection of
mycobacterial DNA in formalin-fixed, paraffin-embedded tissue specimens by
duplex polymerase chain reaction: application to histopathologic diagnosis. Mod.
Pathol. 10, 78–83.
2. Totsch, M., Bocker, W., Brommelkamp, E., Fille, M., Kreczy, A., Ofner, D.,
Schmid, K. W., and Dockhorn-Dworniczak, B. (1996) Diagnostic value of differ-ent PCR assays for the detection of mycobacterial DNA in granulomatous lym-phadenopathy. J. Pathol. 178, 221–226.
3. Bascunana, C. R. and Belak, K. (1996) Detection and identification of mycobac-teria in formalin-fixed, paraffin-embedded tissues by nested PCR and restriction
enzyme analysis. J. Clin. Microbiol. 34, 2351–2355.
4. Ferrara, G., Cannone, M., Guadagnino, A., Nappi, O., and Barberis, M. C. (1999)
Nested polymerase chain reaction on vaginal smears of tuberculous cervicitis: a
case report. Acta Cytol. 43, 308–312.
5. Salian, N. V., Rish, J. A., Eisenach, K. D., Cave, M. D., and Bates, J. H. (1998)
Polymerase chain reaction to detect Mycobacterium tuberculosis in histologic
specimens. Am. J. Respir. Clin. Care Med. 158, 1150–1155.
6. Berk, R. H., Yazici, M., Atabey, N., Ozdamar, O. S., Pabuccuoglu, U., and Alici,
E. (1996) Detection of Mycobacterium tuberculosis in formaldehyde solution-fixed, paraffin-embedded tissue by polymerase chain reaction in Pott’s disease.
Spine 21, 1991–1995.
Detection and Speciation of Mycobacteria 387
7. Marchetti, G., Gori, A., Catozzi, L., Vago, L., Nebuloni, M., Rossi, M. C., Esposti,
A. D., Bandera, A., and Franzette, F. (1998) Evaluation of PCR in detection of
mycobacterial tuberculosis from formalin-fixed, paraffin-embedded tissues: com-parison of four amplification assays. J. Clin. Microbiol. 36, 1512–1517.
8. Hardman, W. J., Benian, G. M., Howard, T., McGowan, J. E. Jr., Metchock, B.,
Murtagh, J. J. (1996) Rapid detection of mycobacteria in inflammatory necrotiz-ing granulomas from formalin-fixed, paraffin-embedded tissue by PCR in clini-cally high-risk patients with acid-fast stain and culture negative tissue biopsies.
Am. J. Clin. Pathol. 106, 184–389.
9. Kox, L. F., van Leeuwen, J., Knijper, S., Jansen, H. M., and Kolk, A. H. (1995)
PCR assay based on DNA coding for 16S rRNA for detection and identification
of mycobacteria in clinical samples. J. Clin. Microbiol. 33, 3225–3233.
10. Cook, S. M., Bartos, R. E., Pierson, C. L., and Frank, T. S. (1994) Detection and
characterization of atypical mycobacteria by the polymerase chain reaction.
Diagn. Mol. Pathol. 3, 53–58.
Quantitation of HIV-1 RNA in Plasma 389
30
Ultrasensitive Quantitation of Human
Immunodeficiency Virus Type 1 RNA
in Plasma by the AMPLICOR and COBAS
AMPLICOR HIV-1 MONITOR™ Tests
Steven Herman, James Novotny, and Maurice Rosenstraus
1. Introduction
The measurement of plasma human immunodeficiency virus type 1 (HIV-1)
RNA levels has become an important tool for identifying individuals likely to
benefit from antiretroviral therapy  (1–7) as well as monitoring patients on
therapy (1 ,8–14) , and is now regarded as standard medical practice for manag-ing the treatment of HIV-1-infected individuals (15–21) . Three commercially
available assays for measuring HIV-1 RNA levels are available. The first-generation AMPLICOR HIV-1 MONITOR™ Test, which uses reverse
transcriptase-polymerase chain reaction (RT-PCR) technology, has a lower
limit of quantitation of 400 copies/mL of plasma (22,23). The NucliSens HIV-1
QT Test (Organon Teknika, Boxtel, Netherlands), a second-generation assay
based on the nucleic acid sequence–based amplification technique, has a lower
limit of quantitation of 100 HIV-1 RNA copies/mL of plasma  (24) . The
Quantiplex HIV-1 Version 2.0 Test (Chiron, Emeryville, CA), which uses the
branched-chain DNA signal amplification technique, has a lower limit of
quantitation of 500 HIV-1 RNA copies/mL of plasma (25 ,26) .
Recently, the use of combination therapy has resulted in rapid and potent
antiretroviral and immunologic effects that lead to a sharp decline in plasma
HIV-1 RNA concentration, frequently to an undetectable level (10 ,12 ,14) . A
more sensitive method with a lower detection limit for plasma HIV-1 RNA is
therefore required. By employing a modified specimen preparation procedure,
389
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
390 Herman, Novotny, and Rosenstraus
which incorporates an ultracentrifugation step to concentrate virus from plasma
(27 ,28) , we increased the sensitivity of the AMPLICOR HIV-1 MONITOR
Test eightfold, achieving a lower limit of quantitation of 50 HIV-1 RNA
copies/mL of plasma ( [29 ,30] ; N. Michael, personal communication).
Roche Molecular Systems has incorporated the UltraSensitive processing
procedure into the first-generation AMPLICOR HIV-1 MONITOR Test; the
second-generation AMPLICOR HIV-1 MONITOR Test, Version 1.5; and the
fully automated COBAS AMPLICOR HIV-1 MONITOR™ Test, Version 1.5.
By offering the option of using either the UltraSensitive or Standard process-ing method, these tests achieve the wide dynamic range required to measure
viral RNA titer throughout the course of HIV-1 infection. The first-generation
AMPLICOR HIV-1 MONITOR Test has been cleared by the Food and Drug
Administration. The second-generation Version 1.5 tests, which are commer-cially available outside the United States, are upgraded tests that contain a
primer pair and modified thermal cycling conditions designed to yield equiva-lent quantitation of all group M subtypes of HIV-1.
2. Materials
2.1. AMPLICOR Kits
2.1.1. AMPLICOR HIV-1 MONITOR Test, Version 1.5
1. AMPLICOR HIV-1 MONITOR, Version 1.5 Kit (contains all specimen prepara-tion, amplification, and detection reagents).
2.1.2. COBAS AMPLICOR HIV-1 MONITOR Test, Version 1.5
1. COBAS AMPLICOR HIV-1 MONITOR, Version 1.5 Kit (contains all specimen
preparation, amplification, and detection reagents).
2.2. Laboratory Supplies and Equipment
2.2.1. Preamplification: Reagent Preparation Area
2.2.1.1. COMMON EQUIPMENT
1. Plastic resealable bag.
2. Eppendorf® Repeater™ pipet with 1.25-mL Combitip® Reservoir (sterile, indi-vidually wrapped).
3. Pipettors (200-µL capacity) with aerosol barrier or positive displacement
RNase-free tips.
4. Latex gloves, powderless.
2.2.1.2. AMPLICOR TEST
1. Perkin-Elmer MicroAmp® reaction tubes.
2. Perkin-Elmer MicroAmp caps.
3. Perkin-Elmer MicroAmp tray/retainers.
4. Perkin-Elmer MicroAmp tray base.
Quantitation of HIV-1 RNA in Plasma 391
2.2.1.3. COBAS AMPLICOR TEST
1. COBAS AMPLICOR A-ring fitted with 12 A-tubes.
2. COBAS AMPLICOR A-ring holder.
2.2.2. Preamplification: Specimen Preparation Area
1. Microcentrifuge (max. relative centrifugal force [RCF] 16,000g, min. RCF
12,500g); Eppendorf 5415C, HERMLE Z230M, or equivalent.
2. Refrigerated ultracentrifuge with fixed-angle rotor (45°, capacity for 24 1.5-mL
tubes) with an RCF of 23,600g (Heraeus Centrifuge 17 RS or Biofuge 28 RS with
HFA 22.1 rotor, Heraeus Biofuge Stratos with the 3331 rotor or equivalent). The
rotor lid must form an aerosol-tight seal to the rotor to contain aerosols in case of
tube failure.
3. 2.0-mL Sterile, nonsiliconized, conical polypropylene screw-cap tubes (Sarstedt
72.694.006 or equivalent).
4. 1.5-mL Sterile, nonsiliconized, conical polypropylene screw-cap tubes (Sarstedt
72.692.105 or equivalent).
5. Tube racks (Sarstedt 93.1428).
6. 70% Ethanol (freshly diluted from 95% using deionized water).
7. Isopropyl alcohol, reagent grade.
8. Sterile fine-tip transfer pipets, RNase-free.
9. Vortex mixer.
10. Latex gloves, powderless.
11. Sterile, disposable, polystyrene serologic pipets (5, 10, and 25 mL).
12. Pipettors (capacity 12.5, 25, 50, 100, 200, 400, 500, 600, 800, and 1000 µL) with
aerosol barrier or positive displacement RNase-free tips.
2.2.3. Postamplification Area
2.2.3.1. AMPLICOR TEST
1. Perkin-Elmer GeneAmp PCR System 9600 or GeneAmp PCR System 2400 ther-mal cycler.
2. MicroAmp Base and cap-installing tool for Perkin-Elmer Applied Biosystems
GeneAmp PCR System 9600 or GeneAmp PCR System 2400.
3. Multichannel pipettor (25 and 100  µL) or electronic pipettor (Impact® or
AMPLICOR®).
4. Aerosol barrier or positive displacement RNase-free pipettor tips (25 and 100 µL)
and barrier-free tips (100 µL).
5. Microwell plate washer.
6. Microwell plate reader.
7. Disposable reagent reservoirs.
8. Microwell plate lid (cat. no. 07-200-376; Fisher).
9. Incubator (37 ± 2°C).
10. Graduated vessels.
11. Distilled or deionized water.
12. Latex gloves, powderless.
392 Herman, Novotny, and Rosenstraus
2.2.3.2. COBAS AMPLICOR TEST
1. COBAS AMPLICOR Analyzer and printer.
2. Operator’s manual for the COBAS AMPLICOR Analyzer.
3. HIV-1 MONITOR Test Method Manual for the COBAS AMPLICOR Analyzer.
4. COBAS AMPLICOR D-cups.
5. Distilled or deionized water.
6. 5-mL Serologic pipets.
7. Graduated cylinder (minimum 1 L).
8. Vortex mixer.
9. Latex gloves, powderless.
3. Methods
The AMPLICOR and COBAS AMPLICOR HIV-1 MONITOR tests are
based on five major processes: specimen preparation, reverse transcription of
target RNA to generate cDNA, PCR amplification of target cDNA using
HIV-1 specific complementary primers, hybridization of the amplified prod-ucts (amplicon) to oligonucleotide probes specific to the target(s), and detec-tion of the probe-bound amplified products by colorimetric determination.
Either of two specimen processing methods can be used with the tests. The
Standard method has a limit of quantitation of 400 copies of HIV-1 RNA/mL
of plasma and a dynamic range of 400–750,000 copies of HIV-1 RNA/mL of
plasma. The UltraSensitive method has a limit of quantitation of 50 copies of
HIV-1 RNA/mL of plasma and a dynamic range of 50–75,000 copies
of HIV-1 RNA/mL of plasma. The user has the option of using either method.
The Standard method would be used when a high viral RNA titer is expected,
e.g., to obtain titers at baseline. The UltraSensitive method would be used when
very low viral titer is expected, e.g., after initiation of therapy or after the viral
titer exhibits a substantial decrease.
For the AMPLICOR test, the user programs a thermal cycler to perform
reverse transcription and amplification automatically; hybridization, detection,
and calculations are performed manually. For the COBAS AMPLICOR HIV-1
test, the user simply enters the test name and specimen identification codes.
The COBAS AMPLICOR Analyzer (31) then uses preprogrammed parameters
to automatically perform reverse transcription, amplification, detection, and
calculation of results without user intervention.
To achieve selective amplification of target nucleic acid from the clinical
specimen, the amplification reaction mixtures for both HIV-1 MONITOR
tests contain AmpErase® and deoxyuridine triphosphate (dUTP)  (32) . A
Quantitation Standard (HIV-1 QS) is utilized as a reference for calculating the
HIV RNA concentration in the specimen.
Quantitation of HIV-1 RNA in Plasma 393
3.1. Selective Amplification
AmpErase (uracil-N-glycosylase) recognizes and catalyzes the destruction
of DNA strands containing deoxyuridine, but not DNA containing thymidine
(32) . Deoxyuridine is not present in naturally occurring DNA, but is always
present in amplicon owing to the use of dUTP as one of the dNTPs in the
master mix reagent; therefore, only amplicon contains deoxyuridine.
Deoxyuridine renders contaminating amplicon susceptible to destruction by
AmpErase prior to amplification of the target DNA. AmpErase, which is
included in the master mix reagent, catalyzes the cleavage of deoxyuridine-containing DNA at deoxyuridine residues by opening the deoxyribose chain at
the C1 position. When heated in the first thermal cycling step at the alkaline
pH of the master mix, the amplicon DNA chain breaks at the position of the
deoxyuridine, thereby rendering the DNA nonamplifiable. AmpErase is inac-tive at temperatures above 55°C (i.e., throughout the thermal cycling steps)
and therefore does not destroy target amplicon. Following amplification, any
residual enzyme is denatured by the addition of the denaturation solution,
thereby preventing the degradation of target amplicon. AmpErase in the
AMPLICOR HIV-1 MONITOR tests has been demonstrated to inactivate at
least 103 copies of deoxyuridine-containing HIV-1 amplicon per PCR.
3.2. Quantitation Standard
The HIV-1 quantitation standard (QS) is a noninfectious 233 nucleotide in
vitro transcribed RNA molecule that contains primer binding sites identical to
those of the HIV-1 target. The HIV-1 QS generates an amplification product of
the same length and base composition as the HIV-1 target sequence. The
HIV-1 QS contains a unique probe binding region that allows amplicon gener-ated from it to be distinguished from HIV-1 amplicon.
In the Standard procedure, a known number of HIV-1 QS copies is added to
each individual specimen at the first step of the specimen-processing proce-dure and is carried through the specimen preparation, reverse transcription,
PCR amplification, hybridization, and detection steps along with the HIV-1
target. Levels of HIV-1 RNA in the test specimens are determined by compar-ing the HIV-1 signal to the HIV-1 QS signal for each specimen. Therefore, the
HIV-1 QS compensates for any effects of inhibition and controls variation in
the specimen preparation, amplification, and detection processes to allow the
accurate quantitation of HIV-1 RNA in each specimen.
In the UltraSensitive procedure, a known number of HIV-1 QS copies is
added to each individual specimen immediately after it has been concentrated
by ultracentrifugation and is then carried through all the remaining assay steps.
394 Herman, Novotny, and Rosenstraus
The HIV-1 QS cannot be added to the specimen prior to ultracentrifugation
because it is in the form of pure RNA. Whereas HIV-1 RNA associated with
viral particles is efficiently recovered during ultracentrifugation, free RNA
molecules such as the HIV-1 QS are not. Thus, the HIV-1 QS does not control
or compensate for variation in virus recovery during the initial ultracentrifuga-tion step.
3.3. Collection, Transport, and Storage of Specimens
Both the COBAS AMPLICOR and AMPLICOR HIV-1 MONITOR tests
are for use with plasma specimens alone. Blood should be collected in sterile
tubes using EDTA (lavender top, Becton Dickinson no. 6454 or equivalent) or
acid citrate dextrose (ACD) (yellow top, Becton Dickinson no. 4606 or equiva-lent) as the anticoagulant. Specimens containing heparin as the anticoagulant
are unsuitable for this test. Store whole blood at 2–25°C for no longer than 6 h.
Specimens containing ACD as the anticoagulant will yield test results that are
approx 15% lower than those obtained from specimens containing EDTA as
the anticoagulant owing to the dilution effect of the 1.5 mL of ACD anticoagu-lant present in the blood collection tube. Whole blood may be stored, or trans-ported, at 2–25°C for up to 6 h from the time of collection.
Plasma must be separated from whole blood within 6 h of collection by
centrifugation at 800–1600g for 20 min at room temperature (see Note 1).
After separation from whole blood, plasma must be transferred to a sterile
polypropylene tube. Plasma specimens may be stored, or transported, at 2–8°C
for up to 5 d, or frozen at –20°C. It is recommended that specimens be stored in
600 to 700-µL aliquots in sterile, 2.0-mL polypropylene screw-cap tubes (such
as Sarstedt 72.694.006). Plasma specimens may be frozen and thawed up to
three times.
3.4. Work Flow
Both the COBAS AMPLICOR and AMPLICOR HIV-1 MONITOR tests
can be completed in 1 d or over the course of 2 d. If the testing is to be com-pleted in a single workday, perform the procedures in Subheading 3.5. first
and then proceed to  Subheading 3.6. If the testing is to be completed over
the course of 2 d, perform the procedures described in Subheadings 3.6.1. or
3.6.2. on d 1 and store the processed specimens as described in step 19 or 14 of
the UltraSensitive or Standard specimen-processing procedures, respectively.
On d 2, perform the procedures described in  Subheading 3.5. first and
then proceed to  step 20 or 15 of the UltraSensitive or Standard specimen-processing procedures, respectively.
Quantitation of HIV-1 RNA in Plasma 395
The laboratory should contain two distinct work areas, one for
preamplification activities and one for postamplification activities. Within the
preamplification area, there should be separate work spaces for preparation of
reagents and preparation of specimens (see Note 2).
3.5. Preparation of Reagents
Each kit contains reagents sufficient for two 12-test runs, which may be
performed separately or simultaneously. It is recommended that one replica-tion of the AMPLICOR HIV-1 MONITOR (–) control, AMPLICOR HIV-1
MONITOR low (+) control, and AMPLICOR HIV-1 high (+) control be
included in each test run. For the most efficient use of reagents, specimens and
controls should be processed in batches that are multiples of 12.
1. Determine the appropriate number of MicroAmp reaction tubes (AMPLICOR)
or A-rings (COBAS AMPLICOR) needed for patient specimen and control test-ing. Place the reaction tubes in the MicroAmp tray and lock into place with the
retainer or place the A-ring(s) in the A-ring holder(s).
2. Prepare working master mix by adding 100  µL of HIV-1 Mn2+ to one vial of
HIV-1 master mix (see Notes 3–6). Mix by inverting 10–15 times (see Note 7).
The pink dye in the manganese solution is used for visual confirmation that the
manganese solution has been added to the master mix. Discard the remaining
manganese solution. The working master mix must be stored at 2–8°C and used
within 4 h of preparation.
3. Add 50 µL of working master mix to each reaction tube or A-tube using a repeat
pipettor or a pipettor with an aerosol barrier or positive displacement tip. Do not
cap the reaction tubes or close the covers of the A-tubes at this time. Discard
unused working master mix.
4. Place the MicroAmp tray and the appropriate number of reaction tube caps or the
A-ring(s) in a resealable plastic bag and seal the bag securely. Move the tray or
A-ring(s) to the specimen preparation work space in the preamplification area.
Store the reaction tubes or A-ring(s) containing working master mix at 2–8°C
until preparation of specimen and control is completed. Working master mix is
stable for 4 h at 2–8°C in reaction tubes or A-tubes sealed in the plastic bag.
3.6. Preparation of Specimen
In both the UltraSensitive and Standard specimen-processing methods,
HIV-1 RNA is extracted from viral particles with HIV-1 lysis reagent and
the extracted RNA is then recovered by alcohol precipitation (see Note 8). The
HIV-1 lysis reagent contains the chaotropic reagent guanidine thiocyanate. The
UltraSensitive method differs in two ways from the Standard method. First,
viral particles are concentrated from 500 µL of plasma by ultracentrifugation
prior to extraction, whereas 200 µL of whole plasma is extracted directly in the
396 Herman, Novotny, and Rosenstraus
Standard method. Second, the recovered RNA is resuspended in a final volume
of 100 µL in the UltraSensitive method compared with 400 µL in the Standard
method. Thus, compared to the Standard method, the UltraSensitive method
produces a 10-fold (2.5 times more plasma in one fourth the volume) more
concentrated processed specimen. This increase in concentration is possible
because the ultracentrifugation step eliminates soluble plasma components that
potentially interfere with RT-PCR.
3.6.1. UltraSensitive Method
1. Precool the ultracentrifuge and rotor to 2–8°C as described in the operating
instructions for the ultracentrifuge.
2. Prepare 70% ethanol (see Notes 9 and 10). For 12 tests, mix 11.0 mL of 95%
ethanol and 4.0 mL of deionized water.
3. Label one 1.5-mL screw-cap tube for each patient specimen and label three addi-tional tubes as HIV-1 (–) C, HIV L(+)C, and HIV H(+)C.
4. Prepare the UltraSensitive working lysis reagent. Vortex HIV-1 QS for at least
10 s before use (see Notes 11 and 12). For each batch of up to 12 specimens and
controls, add 25  µL of HIV-1 QS to one bottle of HIV-1 lysis buffer and mix
well. It is not necessary to measure the volume of HIV-1 lysis buffer. The pink
dye in the HIV-1 QS is used for visual confirmation that the HIV-1 QS has been
added to the lysis reagent. Discard the remaining HIV-1 QS. UltraSensitive work-ing lysis reagent is stable for 4 h at room temperature. If using frozen specimens,
thaw the specimens at room temperature and vortex for 3–5 s. Spin the specimen
tube briefly to collect the specimen in the base of the tube. Take care to avoid
contaminating gloves when manipulating specimens (see Note 13).
5. Add 500  µL of each patient specimen to the appropriately labeled tubes (see
Note 14).
6. Add 500 µL of normal human plasma to each of the appropriately labeled control
tubes (see Note 15).
7. Put an orientation mark on each tube and place the tubes in the ultracentrifuge
with the orientation marks facing outward, so that the pellets will align with the
orientation marks. Centrifuge the specimens and controls at 23,600g for 60 min
at 2–8°C.
8. Using a new, fine-tip disposable transfer pipet for each tube, carefully remove
and discard the supernatant from each tube, being careful not to disrupt the pellet
(which may not be visible) (see Note 16). Remove as much liquid as possible
without disturbing the pellet. Withdraw the supernatant slowly, allowing the liq-uid to drain completely off the sides of the tube. Do not use vacuum aspiration.
9. Add 600  µL of working lysis reagent to each of the labeled tubes and cap the
tubes. Check that the working lysis reagent is pink to confirm that HIV-1 QS was
added to the lysis reagent.
10. Prepare the UltraSensitive controls as follows:
a. Vortex HIV-1 (–), HIV-1 low (+), and HIV-1 high (+) controls for 3–5 s.
Quantitation of HIV-1 RNA in Plasma 397
b. Add 12.5 µL of HIV-1 (–) control to the tube labeled HIV-1 (–) C containing
UltraSensitive working lysis reagent. Cap the tube and vortex for 3–5 s.
c. Add 12.5 µL of HIV-1 low (+) control to the tube labeled HIV-1 L(+)C contain-ing UltraSensitive working lysis reagent. Cap the tube and vortex for 3–5 s.
d. Add 12.5 µL of HIV-1 high (+) control to the tube labeled HIV-1 H(+)C contain-ing UltraSensitive working lysis reagent. Cap the tube and vortex for 3–5 s.
11. Incubate the specimen and control tubes for 10 min at room temperature.
12. For each specimen and control tube, remove the cap, add 600 µL of 100% isopro-pyl alcohol (at room temperature), recap the tube, and vortex vigorously for
3–5 s. Do not have more than one tube open at a time.
13. Put an orientation mark on each tube and place the tubes in the microcentrifuge
with the orientation marks facing outward, so that the pellets will align with the
orientation marks. Centrifuge the specimens and controls at maximum speed
(12,500–16,000g) for 15 min at room temperature.
14. Using a new, fine-tip disposable transfer pipet for each tube, carefully remove
and discard the supernatant from each tube, being careful not to disrupt the pellet
(which may not be visible). Remove as much liquid as possible without disturb-ing the pellet. Withdraw the supernatant slowly, allowing the liquid to drain com-pletely off the sides of the tube. Do not use vacuum aspiration.
15. Add 1.0 mL of 70% ethanol (at room temperature) to each tube, recap, and vortex
for 3–5 s.
16. Place the tubes into a microcentrifuge with the orientation marks facing outward
and centrifuge the tubes for 5 min at maximum speed (12,500–16,000g) at room
temperature.
17. Using a new, fine-tip disposable transfer pipet for each tube, carefully remove
the supernatant without disturbing the pellet. The pellet should be clearly visible
at this step. Remove as much of the supernatant as possible. (Note: residual etha-nol can inhibit amplification.)
18. Repeat step 17 to remove as much of the remaining supernatant as possible.
19. Add 100 µL of HIV-1 diluent to each tube. Recap the tubes. Vortex vigorously
for 10 s to resuspend the extracted RNA. Some insoluble material may remain.
Start the reverse transcription/amplification procedure within 2 h of specimen
and control preparation. If amplification cannot be undertaken within 2 h of
preparation, the samples can be stored frozen at –20°C or colder for up to 1 wk,
with no more than one freeze/thaw. More than one freeze/thaw cycle may result
in loss of copy number. If processed specimens and controls were stored frozen
prior to amplification, thaw at room temperature and vortex for 5 s before pro-ceeding to step 20.
20. Add 50 µL of each processed specimen and control to appropriate reaction tubes
or A-tubes containing working master mix using a pipettor with an aerosol bar-rier or a positive displacement tip. Use a new tip for each specimen and control.
Be careful to avoid transferring any precipitated material that may not have gone
back into the solution. Cap the reaction tubes and seal the caps using the
MicroAmp cap-installing tool or cap the A-tubes (see Notes 17–20).
398 Herman, Novotny, and Rosenstraus
21. Record the positions of the controls and specimens. Reverse transcription/ampli-fication must be started within 45 min of the time that the processed specimens
and controls were added to the reaction tubes or A-tubes containing working
master mix. Move the prepared specimens and controls in the MicroAmp trays or
A-rings to the amplification/detection area.
3.6.2. Standard Method
1. Prepare 70% ethanol. For 12 tests, mix 11.0 mL of 95% ethanol and 4.0 mL of
deionized water.
2. Label one 2.0-mL screw-cap tube for each patient specimen and label three addi-tional tubes as HIV-1 (–) C, HIV L(+)C, and HIV H(+)C (see Note 13).
3. Prepare Standard working lysis reagent. Vortex HIV-1 QS for at least 10 s before
use (see Notes 11 and 12). For each batch of up to 12 specimens and controls, add
100 µL of HIV-1 QS to one bottle of HIV-1 lysis reagent and mix well. It is not
necessary to measure the volume of HIV-1 lysis reagent. The pink dye in the
HIV-1 QS is used for visual confirmation that HIV-1 QS has been added to the
lysis reagent. Discard the remaining HIV-1 QS. Standard working lysis reagent
is stable for 4 h at room temperature. If using frozen specimens, thaw the speci-mens at room temperature and vortex for 3–5 s. Spin the specimen tube briefly to
collect the specimen in the base of the tube. Take care to avoid contaminating
gloves when manipulating specimens.
4. Add 600 µL of Standard working lysis reagent to each of the labeled tubes and
cap the tubes. Check that the working lysis reagent is pink to confirm that HIV-1
QS was added to the Standard lysis reagent.
5. Prepare Standard controls as follows:
a. Vortex negative human plasma, HIV-1 (–) control, HIV-1 low (+) control,
and HIV-1 high (+) control for 3–5 s.
b. Add 200 µL of negative human plasma to each of the three control tubes. Cap
the tubes and vortex for 3–5 s.
c. Add 50 µL of HIV-1 (–) control to the tube labeled HIV-1 (–) C containing
Standard working lysis reagent and negative human plasma. Cap the tube and
vortex for 3–5 s.
d. Add 50 µL of HIV-1 low (+) control to the tube labeled HIV-1 L(+)C contain-ing Standard working lysis reagent and negative human plasma. Cap the tube
and vortex for 3–5 s.
e. Add 50 µL of HIV-1 high (+) control to the tube labeled HIV-1 H(+)C con-taining Standard working lysis reagent and negative human plasma. Cap the
tube and vortex for 3–5 s.
6. Add 200 µL of each patient specimen to the appropriately labeled tubes contain-ing Standard working lysis reagent. Cap the tubes and vortex for 3–5 s.
7. Incubate the specimen and control tubes for 10 min at room temperature.
8. For each specimen and control tube, remove the cap, add 800 µL of 100% isopro-pyl alcohol (at room temperature), recap the tube, and vortex vigorously for
3–5 s. Do not have more than one tube open at a time.
Quantitation of HIV-1 RNA in Plasma 399
9. Put an orientation mark on each tube and place the tubes in the microcentrifuge
with the orientation marks facing outward, so that the pellets will align with the
orientation marks. Centrifuge the specimens and controls at maximum speed
(12,500–16,000g) for 15 min at room temperature.
10. Using a new, fine-tip disposable transfer pipet for each tube, carefully remove
and discard the supernatant from each tube, being careful not to disrupt the pellet
(which may not be visible). Remove as much liquid as possible without disturb-ing the pellet. Withdraw the supernatant slowly, allowing the liquid to drain com-pletely off the sides of the tube. Do not use vacuum aspiration.
11. Add 1.0 mL of 70% ethanol (at room temperature) to each tube, recap, and vortex
for 3–5 s.
12. Place the tubes in a microcentrifuge with the orientation marks facing outward,
and centrifuge the tubes for 5 minutes at maximum speed (12,500–16,000g) at
room temperature.
13. Using a new, fine-tip disposable transfer pipet for each tube, carefully remove
the supernatant without disturbing the pellet. The pellet should be clearly visible
at this step. Remove as much of the supernatant as possible. (Note: Residual
ethanol can inhibit amplification.)
14. Add 400 µL of HIV-1 diluent to each tube. Recap the tubes. Vortex vigorously
for 10 s to resuspend the extracted RNA. Some insoluble material may remain.
Start the reverse transcription/amplification procedure within 2 h of specimen
and control preparation. If amplification cannot be undertaken within 2 h of
preparation, the samples can be stored frozen at –20°C or colder for up to 1 wk,
with no more than one freeze/thaw. More than one freeze/thaw cycle may result
in loss of copy number. If processed specimens and controls were stored frozen
prior to amplification, thaw at room temperature and vortex for 5 s before pro-ceeding to step 15.
15. Add 50 µL of each processed specimen and control to appropriate reaction tubes
or A-tubes containing Working master mix using a pipettor with an aerosol bar-rier or a positive displacement tip. Use a new tip for each specimen and control.
Be careful to avoid transferring any precipitated material that may not have gone
back into the solution. Cap the reaction tubes and seal the caps using the
MicroAmp cap-installing tool or cap the A-tubes.
16. Record the positions of the controls and specimens. Reverse transcription/ampli-fication must be started within 45 minutes of the time that the processed speci-mens and controls were added to the reaction tubes or A-tubes containing
Working master mix. Move the prepared specimens and controls in the
MicroAmp trays or A-rings to the amplification/detection area.
3.7. Reverse Transcription and PCR Amplification
The AMPLICOR HIV-1 Version 1.5 tests amplify and detect a 155-base
target sequence located in a highly conserved region of the HIV-1 gag gene
(33) , defined by the primers SK145 and SKCC1B. Each primer is biotinylated
at its 5v end.
400 Herman, Novotny, and Rosenstraus
The reverse transcription and amplification reaction are performed with the
thermostable recombinant enzyme  Thermus thermophilus DNA Polymerase
(rTth pol). In the presence of manganese and under the appropriate buffer con-ditions, rTth pol has both RT and DNA polymerase activity (34) . This allows
both reverse transcription and PCR amplification to occur in the same reaction
mixture.
3.7.1. Reverse Transcription
The reaction tubes are placed in a thermal cycler that automatically per-forms the heating and cooling steps required for reverse transcription and
amplification. First, the reactions are heated to allow the downstream primer
(SKCC1B) to anneal specifically to the HIV-1 and HIV-1 QS target RNA. In
the presence of excess deoxynucleoside triphosphates, including deoxy-adenosine, deoxyguanosine, deoxycytidine, thymidine, and dUTPs, rTth pol
extends the annealed primer, forming a complementary (cDNA) strand.
3.7.2. PCR Amplification
Following reverse transcription of the HIV-1 and HIV-1 QS target RNA, the
reaction mixture is heated to denature the RNA:cDNA hybrid and expose the
HIV-1 and HIV-1 QS target sequences. As the mixture cools, the upstream
primer (SK145) anneals to the cDNA strand and the rTth pol catalyzes the
extension reaction, yielding a double-stranded DNA (dsDNA) copy of the tar-get region of each HIV-1 and HIV-1 QS RNA. This completes the first cycle of
PCR. The reaction mixture is heated again to separate the dsDNA and expose
the primer target sequences. As the mixture cools, the primers anneal to the
target DNA. In the presence of excess dNTPs, rTth pol extends the annealed
primers along the target templates to produce a 155-bp dsDNA molecule
termed an amplicon. This process is automatically repeated for the appropriate
number of cycles, each cycle effectively doubling the amount of amplicon
DNA. Amplification occurs only in the region of the HIV-1 genome between
the primers.
3.7.3. Performing Reverse Transcription/PCR Amplification
(see Notes 21–23)
3.7.3.1. AMPLICOR HIV-1 MONITOR, VERSION 1.5 TEST
1. Place the tray/retainer assembly into the thermal cycler block.
2. Program the Perkin-Elmer GeneAmp PCR System 9600 or GeneAmp PCR Sys-tem 2400 thermal cycler as follows:
a. HOLD program: 2 min at 50°C.
b. HOLD program: 30 min at 60°C.
Quantitation of HIV-1 RNA in Plasma 401
c. CYCLE program (8 cycles): 10 s at 95°C, 10 s at 52°C, 10 s at 72°C.
d. CYCLE program (23 cycles): 10 s at 90°C, 10 s at 55°C, 10 s at 72°C.
e. HOLD program: 15 min at 72°C.
In the CYCLE programs, the ramp times should be left at the default setting
(0 00), which is the maximum rate, and the allowed setpoint error at the default
setting (2°C). Link the four programs together into a METHOD program.
3. Start the METHOD program. The program runs approx 1 h and 15 min.
4. After the thermal cycling is complete (i.e., during the final 15-min HOLD pro-gram), remove the caps from the tubes and immediately pipet 100 µL of denatur-ation solution into each reaction tube using a multichannel pipettor and mix by
pipetting up and down five times. Do not allow the reaction tubes to remain in the
thermal cycler beyond the end of the final HOLD program, and do not extend the
final HOLD program beyond 15 min.
5. The denatured amplicon can be held at room temperature no more than 2 h before
proceeding to the detection reaction. If the detection reaction cannot be performed
within 2 h, recap the tubes and store the denatured amplicon at 2–8°C for up
to 1 wk.
3.7.3.2. COBAS AMPLICOR HIV-1 MONITOR TEST
The COBAS AMPLICOR Analyzer will automatically perform the correct
thermal cycling program, as well as all subsequent assay procedures, after the
user starts the instrument by performing the following operations:
1. Examine the quantities of reagents on board the COBAS AMPLICOR Analyzer.
Prepare enough reagent cassettes to complete the workload.
2. Mix HIV probe suspension 1 (IM PS1, v1.5) well by vortexing. Add 2.5 mL of
IM PS1, v1.5 to one cassette of probe suspension 2 (IM4, v1.5). Place the cas-sette on the test-specific reagent rack. Discard the used IM PS1, v1.5 vial. Record
the date of reagent preparation on the IM4, v1.5 cassette.
3. Mix QS probe suspension 1 (IQ PS1, v1.5) well by vortexing. Add 2.5 mL of IQ
PS1, v1.5 to one cassette of QS probe suspension 2 (IQ4, v1.5). Place the cassette
on the test-specific rack. Discard the used IQ PS1, v1.5 vial. Record the date of
reagent preparation on the IQ4, v1.5 cassette.
4. Prepare working substrate by pipetting 5 mL of substrate B (SB) into one cassette
of substrate A (SB3). Pipet up and down to mix. Discard the open SB vial. Record
the date of preparation on the SB3 cassette.
5. Place the working substrate in the generic reagent rack.
6. Place the cassette of Amplicon dilution reagent (AD3) in the test-specific reagent
rack. Record the date on the AD3 cassette.
7. Place the cassettes of denaturation solution (DN4) and conjugate (CN4) in the
generic reagent rack. Record on each cassette the date it was opened.
8. Identify the reagent racks as generic or test specific using the keypad, bar-code
scanner, or AMPLILINK™ software as described in the operator’s manual for
the analyzer.
402 Herman, Novotny, and Rosenstraus
9. Configure the reagent racks by inputting reagent positions and lot numbers into
the instrument using the keypad, bar-code scanner, or AMPLILINK software as
described in the operator’s manual.
10. Load the reagent rack onto the instrument using the keypad, bar-code scanner, or
AMPLILINK software as described in the operator’s manual. Make sure that
each reagent cassette is in its assigned position and that each cassette fits tightly
into its rack.
11. Place the D-cup rack on the D-cup platform. Six D-cups are required for each
specimen or control and two D-cups are required for each cassette of working
substrate to allow for blanking by the analyzer.
12. Place the A-ring(s) into the thermal cycler segment(s) of the analyzer.
13. Load the A-ring(s) into the analyzer using the keypad, bar-code scanner, or
AMPLILINK software as described in the operator’s manual.
14. Create an A-ring work list as described in the operator’s manual. (Note: At this
time the user will be required to enter the lot-specific Quantitation Standard copy
number and the low [+] and high [+] control ranges provided on the COBAS
AMPLICOR HIV-1 Test, v1.5 Data Cards. Enter appropriate control ranges based
on specimen preparation utilized.)
15. Tightly close the cover of the thermal cycler segment(s).
16. Start the analyzer as described in the operator’s manual.
17. Wait for the analyzer to indicate that the load check has passed.
3.8. Hybridization Reaction
Following PCR amplification, the HIV-1 and HIV-1 QS amplicons are
chemically denatured to form single-stranded DNA by adding denaturation
solution to the reaction tubes (see Notes 22 and 23). The denatured amplicon is
serially diluted and aliquots of undiluted and diluted amplicon are hybridized
to HIV-1-specific (SK102) and HIV-1 QS-specific (CP35) oligonucleotide
probes that are bound to a solid phase. Microwell plates serve as the solid
phase for the AMPLICOR test, and magnetic microparticles serve as the solid
phase for the COBAS AMPLICOR test. In the manual AMPLICOR test, the
microwell plates are incubated at 37°C for 60 min. The COBAS AMPLICOR
Analyzer performs all these steps automatically without user intervention. The
following specific instructions apply to the manual AMPLICOR test:
1. Allow the microwell plates to warm to room temperature before removing from
the foil pouch. Add 100  µL of hybridization buffer to each well required for
testing. Add 25 µL of the denatured amplicon to the HIV-1 wells in row A of the
microwell plates, and mix up and down 10 times with a 12-channel pipettor with
plugged tips (see Note 24). Make serial fivefold dilutions in the HIV-1 wells in
rows B–F as follows. Transfer 25 µL from row A to B and mix as before. Con-tinue through row F. Mix row F as before, and then remove and discard 25 µL.
Discard the pipet tips. These operations may also be performed using the
AMPLICOR Electronic Pipettor.
Quantitation of HIV-1 RNA in Plasma 403
2. Add 25 µL of the denatured amplicon to the QS wells in row G of the microwell
plates and mix up and down 10 times with a 12-channel pipettor with plugged
tips. Transfer 25 µL from row G to H. Mix as before, and then remove 25 µL
from row H and discard. These operations may also be performed using the
AMPLICOR Electronic Pipettor (see Note 25).
3. Cover the microwell plates and incubate for 1 h at 37 ± 2°C.
4. Wash the microwell plates five times with the working wash solution using an
automated microwell plate washer. Program the washer as follows:
a. Aspirate the contents of the well.
b. Fill each well to the top (400–450 µL). Soak for 30 s and aspirate dry.
c. Repeat step 4b four more times.
d. After automated washing is completed, tap the plate dry.
3.9. Detection of Reaction
Following the hybridization reaction, the microwell plates (or tubes of mag-netic microparticles) are washed to remove any unbound material (see Notes
26 and 27). An avidin-horseradish peroxidase (Av-HRP) conjugate is added to
each well of the microwell plates (or to tubes of magnetic microparticles) and
incubated at 37°C for the appropriate amount of time. Av-HRP binds to the
biotin-labeled amplicon captured by the plate-bound oligonucleotide probes.
The microwell plates (or tubes of magnetic microparticles) are washed again to
remove unbound Av-HRP, and a substrate solution containing hydrogen per-oxide and 3,3v,5,5v-tetramethylbenzidine (TMB) is added to the microwell
plates (or tubes of magnetic microparticles). In the presence of hydrogen per-oxide, the bound Av-HRP catalyzes the oxidation of TMB to form a colored
complex. In the AMPLICOR test, the reaction is stopped by addition of a weak
acid, and the optical density (OD) at 450 nm is measured using an automated
microwell plate reader. In the COBAS AMPLICOR test, the reaction is not
stopped; after a precisely timed incubation, the OD of the reaction at 660 nm is
measured by a spectrophotometer integrated in the COBAS AMPLICOR Ana-lyzer. The COBAS AMPLICOR Analyzer performs all these steps automati-cally without user intervention. The following specific instructions apply to
the manual AMPLICOR assays:
1. Add 100  µL Av-HRP conjugate to each well. Cover the microwell plates and
incubate for 15 min at 37 ± 2°C.
2. Wash the microwell plates as described in Subheading 3.8., step 4.
3. Prepare working substrate solution. For each microwell plate, measure 12 mL of
substrate A and 3 mL of substrate B. Mix together to prepare working substrate.
Protect the working substrate from direct light. Working substrate must be at
room temperature and used within 3 h of preparation (see Notes 28 and 29).
4. Pipet 100 µL of working substrate solution into each well.
404 Herman, Novotny, and Rosenstraus
5. Allow color to develop for 10 min at room temperature in the dark.
6. Add 100 µL of stop reagent to each well.
7. Measure the OD at 450 nm (single wavelength) within 10 min of adding the stop
reagent (see Notes 30 and 31).
3.10. Calculation of HIV-1 RNA Concentration
Viral RNA load is quantitated by utilizing the HIV-1 QS, which is added to
the test sample at a known concentration. For the AMPLICOR test, the follow-ing calculations are performed manually. The COBAS AMPLICOR Analyzer
automatically performs these calculations and reports the viral titer, or the
appropriate message when the viral titer cannot be calculated from the test
results.
1. For each specimen or control, the wells in rows A, B, C, D, E, and F represent
neat and 1 5, 1 25, 1 125, 1 625, and 1 3125 dilutions, respectively, of the
HIV-1 amplicon. For each sample, select the well giving the lowest HIV-1 signal
having an OD *0.20 and )2.0 absorbance units. The absorbance values should
decrease with the serial dilutions, with the highest value for each specimen and
control in well A and the lowest in well F.
a. If all HIV-1 absorbance values are <0.2, do not proceed with  steps 2–7.
Report results as “HIV-1 RNA not detected (<400 copies/mL)” or “HIV-1 RNA
not detected (<50 copies/mL),” depending on whether the Standard or Ultra-Sensitive method was used.
b. If all HIV-1 absorbance values are >2.0, the HIV-1 copy number is above the
linear range of the assay. Do not proceed with steps 2–7. Report the result as
“Not determined.” If the Standard processing method was used, prepare a
1 50 dilution of the original specimen with HIV-1 negative human plasma
and repeat the test. Calculate the HIV-1 concentration as described in steps
2–7 and multiply the result by 50. If the UltraSensitive processing method
was used, retest the original specimen using the Standard specimen-process-ing procedure.
c. If HIV-1 absorbances do not follow the pattern of decreasing from well A to
F, examine the data to determine whether an error in dilution occurred. If the
out-of-sequence values and all more concentrated wells are >2.3, the signals
are saturated: no error occurred. If the out-of-sequence values and all more
diluted wells are <0.1, the signals are at background: no error occurred. In
both cases, calculate the viral titer as described in steps 2–7. If the out-of-sequence values fall between 0.2 and 2.0, an error occurred. The result for
that specimen is invalid, and the entire test procedure for that specimen
(including specimen processing) must be repeated.
2. Subtract the background absorbance (0.07 for the AMPLICOR test) from the
absorbance value of the selected well.
3. Calculate the HIV-1 total OD by multiplying the background-corrected HIV-1
absorbance by the dilution factor associated with the selected well.
Quantitation of HIV-1 RNA in Plasma 405
4. For each specimen or control, the wells in rows G and H represent neat and 1 5
dilutions, respectively, of the HIV-1 amplicon. For each sample, select the well
giving the lowest HIV-1 QS signal having an OD  *0.30 and <2.0 absorbance
units. For each specimen and control, the absorbance value for row G should be
higher than the value for row H.
a. If all HIV-1 QS absorbance values are <0.3, the result for that specimen is
invalid. Either the processed specimen was inhibitory to amplification or the
RNA was not recovered during specimen processing. Repeat the entire proce-dure (including specimen processing) for that specimen.
b. If all HIV-1 QS absorbance values are >2.0, an error occurred and the result
for that specimen is invalid. Repeat the entire procedure (including specimen
processing) for that specimen.
c. If HIV-1 QS absorbance of well H is greater than the absorbance for well G,
an error occurred and the result for that specimen is invalid. Repeat the entire
procedure (including specimen processing) for that specimen.
5. Subtract the background absorbance (0.07 for the AMPLICOR test) from the
absorbance value for the selected well.
6. Calculate the HIV-1 QS total OD by multiplying the background-corrected QS
absorbance by the dilution factor associated with the selected well.
7. Calculate the HIV-1 RNA copies/mL of plasma for each specimen using the
following equation:
in which the sample volume factor is 4 for the UltraSensitive method or 40 for
the Standard method (the equivalent of 250 or 25 µL of plasma is added to each
amplification reaction, respectively).
3.11. Daily Maintenance of COBAS AMPLICOR Analyzer
1. Wipe the initialization post with a lint-free moist cloth and dry.
2. Wipe the D-cup handler tip with a lint-free moist cloth and dry.
3. Check the wash buffer reservoir and fill if necessary.
4. Prepare working wash buffer (1X) by adding 1 vol of 10X wash concentrate to
9 vol of distilled or deionized water. Mix well. Keep a minimum of 3–4 L of
working wash buffer (1X) in the wash buffer reservoir of the system at all times.
5. Empty the waste container.
6. Prime the analyzer.
7. During the priming, check the syringes and tubing as well as the transfer tip.
8. Prior to each run, do the following:
a. Check the waste container and empty if necessary.
b. Check the wash buffer reservoir and add buffer if necessary.
c. Replace used D-cup racks.
d. Prime the analyzer.
HIV-1 RNA copies/mL =
× Input HIV-1 QS copies/PCR × sample volume factor
(Total HIV-1 OD/Total HIV-1 QS OD)
406 Herman, Novotny, and Rosenstraus
3.12. Test Performance Characteristics
The AMPLICOR HIV-1 MONITOR, Version 1.5 and COBAS AMPLICOR
HIV-1 MONITOR, Version 1.5 tests have a dynamic range of 50–75,000 cop-ies of HIV RNA/mL of plasma for specimens processed by the UltraSensitive
method (26) . The dynamic range for specimens processed by the Standard
method is 400–750,000 copies of HIV-1 RNA/mL of plasma. For specimens
that fell within the overlapping portion of the linear ranges, the UltraSensitive
method yielded RNA concentrations that were only slightly lower (median =
22% lower) than those obtained by the Standard method (26) . This agreement
means that laboratories can switch between the two specimen-processing meth-ods, as dictated by the HIV-1 RNA titer, to obtain an overall dynamic range of
50–750,000 copies of HIV-1/mL of plasma.
The AMPLICOR HIV-1, Version 1.5 and COBAS AMPLICOR HIV-1
MONITOR, Version 1.5 tests amplified all group M subtypes with equal effi-ciency (26) . By contrast, the first-generation AMPLICOR HIV-1-MONITOR
Test amplified subtypes A, E, F, and G less efficiently than the group B. The
improved performance of the new tests was achieved by replacing the primer
pair in the first-generation test with a single pair of consensus primers that
recognize equally all group M subtypes.
4. Notes
1. Before work begins, all work surfaces used for PCR should be cleaned with 10%
bleach followed by 70% alcohol (isopropanol or ethanol). This process will
destroy bacteria and viruses, and will denature DNA or RNA. Rinsing surfaces
with alcohol destroys any residual bleach, which could react with target nucleic
acid and interfere with test performance. UV lighting should be used to irradiate
DNA that may be on the work surface. It is good practice to turn on the UV light
20–30 min prior to beginning work.
2. Work flow in the laboratory must proceed in a unidirectional manner, beginning
in the preamplification area and moving to the postamplification area.
Preamplification activities must begin with reagent preparation and proceed to
specimen preparation. Supplies and equipment must be dedicated to each
preamplification activity and not used for other activities or moved between
areas. Gloves must be worn in each area and must be changed before leaving that
area. Equipment and supplies used for preparing reagents must not be used for
preparing specimens or for processing amplified DNA or other sources of target
DNA. Postamplification supplies and equipment must remain in the post-amplification area at all times.
3. Separate sets of pipets and pipet tips must be dedicated for use in the reagent,
specimen preparation, and detection procedures. Doing so will protect reagents
from specimen or amplicon contamination.
Quantitation of HIV-1 RNA in Plasma 407
4. Glove powder can nonspecifically inhibit each of the major steps in the PCR
detection process. For this reason, all AMPLICOR PCRs should be performed
with powder-free gloves.
5. For the AMPLICOR test, soak all Perkin-Elmer amplification trays, retainers,
and bases in 10% bleach and rinse thoroughly with deionized water before reuse.
6. Use aerosol barrier tips for all liquid transfers to prevent reagent contamination.
The use of aerosol barrier tips will also prevent RNases from being introduced
into reaction tubes or A-tubes, which can destroy target RNA and lead to false-negative results.
7. Master mix should be inverted 10–15 times, not vortexed. Vortexing can inacti-vate the rTth pol in the master mix.
8. Use aerosol barrier tips for all liquid transfers to prevent reagent contamination
and cross contamination of nucleic acid from specimen to specimen. Aerosol
barrier tips also prevent RNases from being introduced, which can destroy target
RNA and lead to false-negative results.
9. Use only 70% ethanol; the 70% ethanol wash was found to be more reproducible
during the development of this assay. Ethanol should be made fresh daily with
deionized H2O. Do not use diethylpyrocarbonate (DEPC)-treated water; various
preparations of DEPC water have been reported to cause reduced signals.
10. Isopropanol and ethanol must be reagent-grade quality or better.
11. Dissolve the lysis reagent completely before adding the well-vortexed HIV-1 QS.
Lysis buffer may be warmed at 37°C, but no warmer, for up to 30 min. Always bring
warmed lysis reagent to room temperature before adding the HIV-1 QS.
12. The HIV-1 QS must never be heated to 37°C. The HIV-1 QS must be well
vortexed before adding the lysis reagent. Original vials of all AMPLICOR con-trols and standards should be vortexed as follows: upright for 5 s, upside down
for 5 s, upright for 10 s; finish by tapping the vial on the counter to remove any
liquid inside the cap.
13. Use only sterile 2-mL screw-cap tubes for storing and preparing specimens. Flip-cap tubes can cause specimen aerosols when tubes are opened. They can also
cause splashes that can result in cross contamination or a biosafety hazard.
14. To avoid cross contamination when transferring aliquots of specimen to labeled
preparation tubes, uncap one specimen tube and one labeled preparation tube at a
time. After transferring the specimen aliquot, recap both the specimen and the
labeled preparation tubes before proceeding to the next specimen. Use a new
aerosol barrier pipet tip for each specimen. Replace gloves immediately if they
become contaminated with specimen.
15. When preparing controls, combine negative human plasma with the lysis buffer
in the control tubes and vortex well before adding the control. This will destroy
possible RNase activity in the negative human plasma that could degrade control
RNA. It is best to add the control last and then start the 10-minute RT incubation.
16. Use fine-tip transfer pipets to aspirate all supernatants.
17. Arrange tubes of processed specimens, amplification tubes, and waste containers
so that the pipet does not have to pass over open reaction tubes.
408 Herman, Novotny, and Rosenstraus
18. Uncap one tube of processed specimen at a time. After transferring an aliquot of
specimen to the amplification tube, recap the tube before proceeding to the next
specimen.
19. Use aerosol barrier tips when transferring aliquots of processed specimens to
prevent pipettors from becoming contaminated with nucleic acid.
20. To avoid cross contamination between samples, cap each A-tube (for COBAS
tests) or each column of MicroAmp tubes (for AMPLICOR tests) immediately
after adding the processed specimens.
21. Use only Perkin-Elmer 9600 or 2400 thermal cyclers for performing manual
AMPLICOR tests. The amplification parameters have been optimized for these
systems and may yield suboptimal results when performed on other thermal
cyclers.
22. Remove caps from amplification tubes slowly and with extreme care just prior to
adding the denaturation solution. If the caps are opened too rapidly, droplets of
reaction mixture may splash into neighboring tubes. Because reaction tubes may
contain extremely high levels of amplicon, the introduction of a small amount of
contaminating reaction mixture into a negative tube may be sufficient to produce
a false-positive result.
23. Use plugged tips and a 12-channel multichannel pipettor (range 50–300 µL) or
the AMPLICOR Electronic Pipettor to add denaturation solution to the wells.
Change the tips for each row. Pipet up and down to mix well.
24. Use plugged tips and a 12-channel multichannel pipettor (range 5–50 µL) or the
AMPLICOR Electronic Pipettor to transfer aliquots of reaction mixture to the
microwell plates and perform the serial dilutions. Work carefully to avoid splash-ing, which could result in cross contamination of neighboring wells.
25. Recap reaction tubes with new caps immediately after transferring the reaction
mixture to the microwell plates. This will help prevent accidental spillage, which
could contaminate equipment and work surfaces with large amounts of amplicon.
26. Clean disposable reagent reservoirs should be used for detection reagents. If
reagent reservoirs are reused they must be labeled for specific reagents and
cleaned thoroughly between each use with deionized water. Do not reuse reagent
reservoirs more than five times.
27. Working wash buffer must be prepared by measuring the 10X wash concentrate
and then adding nine equal parts of deionized water. Owing to overfill, there may
be up to 120 mL of wash buffer concentrate per bottle. Prepare wash buffer as
follows: Mix well 100 mL of 10X wash concentrate + 900 mL of deionized
water.
28. Substrates A and B must be measured when preparing the working substrate. All
vials are overfilled so that the exact amount of reagents may be pipeted from the
bottle. The overfill between substrates A and B might not be proportional.
29. Mix substrates A and B no more than 3 h before use. Store protected from light.
Excessive storage or exposure to light may cause high background signals and
false-positive results.
Quantitation of HIV-1 RNA in Plasma 409
30. Microwell plates are measured at 450 nm without a reference filter. All
AMPLICOR tests have been optimized for this measurement. Using a reference
filter will falsely depress the sensitivity and results.
31. Measure the absorbance within 10 min of adding stop reagent to the microwell
plates.
References
1. Lathey, J. L., Hughes, M. D., Fiscus, S. A., Pi, T., Jackson, J. B., Rasheed, S.,
Elbeik, T., Reichman, R., Japour, A., D’Aquila, R. T., Scott, W., Griffith, B. P.,
Hammer, S. M., and Katzenstein, D. A. for the AIDS Clinical Trials Group Proto-col 175 Team. (1998) Variability and prognostic values of virologic and CD4 cell
measures in human immunodeficiency virus type 1-infected patients with
200–500 CD4 cells/mm3 (ACTG 175). J. Infect. Dis. 177, 617–624.
2. Mellors, J. W., Kingsley, L. A., Rinaldo, C. R. Jr., Todd, J. A., Hoo, B. S., Kokka,
R. P., and Gupta, P. (1995) Quantitation of HIV-1 RNA in plasma predicts out-come after seroconversion. Ann. Intern. Med. 122, 573–579.
3. Mellors, J. W., Rinaldo, C. R. Jr., Gupta, P., White, R. M., Todd, J. A., and
Kingsley, L. A. (1996) Prognosis in HIV-1 infection predicted by the quantity of
virus in plasma. Science 272, 1167–1170.
4. O’Brien, W. A., Hartigan, P. M., Martin, D., Esinhart, J., Hill, A., Benoit, S.,
Rubin, M., Simberkoff, M. S., and Hamilton, J. D. for the Veterans Affairs Coop-erative Study Group on AIDS. (1996) Changes in plasma HIV-1 RNA and CD4+
lymphocyte counts and the risk of progression to AIDS.  N. Engl. J. Med. 334,
426–431.
5. Saksela, K., Stevens, C. E., Rubinstein, P., Taylor, P. E., and Baltimore, D. (1995)
HIV-1 messenger RNA in peripheral blood mononuclear cells as an early marker
of risk for progression to AIDS. Ann. Intern. Med. 123, 641–648.
6. Wong, M. T., Dolan, M. J., Kozlow, E., Doe, R., Melcher, G. P., Burke, D. S.,
Boswell, R. N., and Vahey, M. (1996) Patterns of virus burden and T cell pheno-type are established early and are correlated with the rate of disease progression in
human immunodeficiency virus type 1-infected persons.  J. Infect. Dis. 173,
877–887.
7. Mellors, J. W., Munoz, A., Giorgi, J. V., Margolick, J. B., Tassoni, C. J., Gupta,
P., Kingsley, L. A., Todd, J. A., Saah, A. J., Detels, R., Phair, J. P., and Rinaldo,
C. R. Jr. (1997) Plasma viral load and CD4+ lymphocytes as prognostic markers
of HIV-1 infection. Ann. Intern. Med. 126, 946–954.
8. Eron, J. J., Benoit, S. L., Jemsek, J., MacArthur, R. D., Santana, J., Quinn, J. B.,
Kuritzkes, D. R., Fallon, M. A., and Rubin, M. for the North American HIV Work-ing Party. (1995) Treatment with lamivudine, zidovudine, or both in HIV-positive
patients with 200 to 500 CD4+ cells per cubic millimeter. N. Engl. J. Med. 333,
1662–1669.
9. Fiscus, S. A., Hughes, M. D., Lathey, J. L., Pi, T., Jackson, B., Rasheed, S., Elbeik,
T., Reichman, R., Japour, A., Byington, R., Scott, W., Griffith, B. P., Katzenstein,
410 Herman, Novotny, and Rosenstraus
D. A., and Hammer, S. M. for the AIDS Clinical Trials Group Protocol 175 Team.
(1998) Changes in virologic markers as predictors of CD4 cell decline and pro-gression of disease in human immunodeficiency virus type 1-infected adults
treated with nucleosides. J. Infect. Dis. 177, 625–633.
10. Gulick, R. M., Mellors, J. W., Havlir, D., Eron, J. J., Gonzalez, C., McMahon, D.,
Richman, D. D., Valentine, F. T., Jonas, L., Meibohm, A., Emini, E. A., and
Chodakewitz, J. A. (1997) Treatment with indinavir, zidovudine, and lamivudine
in adults with human immunodeficiency virus infection and prior antiretroviral
therapy. N. Engl. J. Med. 337, 734–739.
11. Hughes, M. D., Johnson, V. A., Hirsch, M. S., Bremer, J. W., Elbeik, T., Erice,
A., Kuritzkes, D. R., Scott, W. A., Spector, S. A., Basgoz, N., Fischl, M. A., and
D’Aquila, R. T. for the ACTG 241 Protocol Virology Substudy Team. (1997)
Monitoring plasma HIV-1 RNA levels in addition to CD4+ lymphocyte count
improves assessment of antiretroviral therapeutic response. Ann. Intern. Med. 126,
929–938.
12. Myers, M. W., Montaner, J. G., and Group, T. I. S. (1996) A randomized, double-blinded comparative trial of the effects of zidovudine, didanosine and nevirapine
combinations in antiviral naive, AIDS-free, HIV-infected patients with CD4
counts 200–600/mm3, in Program and Abstracts: XI International Conference on
AIDS (Vancouver, BC, Canada), XIth International Conference on AIDS Society,
Vancouver, BC, Canada, p. 291.
13. O’Brien, W. A., Hartigan, P. M., Daar, E. S., Simberkoff, M. S., and Hamilton, J. D.,
for the VA Cooperative Study Group on AIDS. (1997) Changes in plasma HIV
RNA levels and CD4+ lymphocyte counts predict both response to antiretroviral
therapy and therapeutic failure. Ann. Intern. Med. 126, 939–945.
14. Perelson, A. S., Essunger, P., Cao, Y., Vesanen, M., Hurley, A., Saksela, K.,
Markowitz, M., and Ho, D. D. (1997) Decay characteristics of HIV-1-infected
compartments during combination therapy. Nature 387, 188–191.
15. BHIVA Guidelines Co-ordinating Committee. (1997) British HIV Association
guidelines for antiretroviral treatment of HIV seropositive individuals. Lancet 349,
1086–1092.
16. Carpenter, C. C., Fischl, M. A., Hammer, S. M., et al. (1996) Antiretroviral therapy
for HIV infection in 1996: recommendations of an international panel. Interna-tional AIDS Society-USA. JAMA 276, 146–154.
17. Centers for Disease Control and Prevention. (1998) Guidelines for the use of
antiretroviral agents in HIV-infected adults and adolescents.  Morbid. Mortal.
Weekly Rep. 47(RR-5), 43–82.
18. Centers for Disease Control and Prevention. (1998) Guidelines for the use of
antiretroviral agents in pediatric HIV infection.  Morbid. Mortal. Weekly Rep.
47(RR-4), 1–43.
19. Centers for Disease Control and Prevention. (1998) Public Health Service Task
Force recommendations for the use of antiretroviral drugs in pregnant women
infected with HIV-1 for maternal health and for reducing perinatal HIV-1 trans-mission in the United States. Morbid. Mortal. Weekly Rep. 47(RR-2), 1–30.
Quantitation of HIV-1 RNA in Plasma 411
20. Volberding, P. A. (1996) HIV quantification: clinical applications. Lancet 347,
71–73.
21. Saag, M. S., Holodniy, M., Kuritzkes, D. R., O’Brien, W. A., Coombs, R.,
Poscher, M. E., Jacobsen, D. M., Shaw, G. M., Richman, D. D., and Volberding,
P. A. (1996) HIV viral load markers in clinical practice. Nat. Med. 2, 625–629.
22. Mulder, J., McKinney, N., Christopherson, C., Sninsky, J., Greenfield, L., and
Kwok, S. (1994) Rapid and simple PCR assay for quantitation of human immuno-deficiency virus type 1 RNA in plasma: application to acute retroviral infection.
J. Clin. Microbiol. 32, 292–300.
23. Roche Diagnostic Systems. (1996) AMPLICOR HIV-1 MONITOR Test package
insert. Roche Diagnostic Systems, Branchburg, NJ, pp. 8,9.
24. Segondy, M., Ly, T. D., Lapeyre, M., and Montes, B. (1998) Evaluation of the
Nuclisens HIV-1 QT assay for quantitation of human immunodeficiency virus
type 1 RNA levels in plasma. J. Clin. Microbiol. 36, 3372–3374.
25. Lin, H. J., Pedneault, L., and Hollinger, F. B. (1998) Intra-assay performance
characteristics of five assays for quantification of human immunodeficiency virus
type 1 RNA in plasma. J. Clin. Microbiol. 36, 835–839.
26. Nolte, F. S., Boysza, J., Thurmond, C., Clark, W. S., and Lennox, J. L. (1998)
Clinical comparison of an enhanced-sensitivity branched-DNA assay and reverse
transcription-PCR for quantitation of human immunodeficiency virus type 1 RNA
in plasma. J. Clin. Microbiol. 36, 716–720.
27. Mulder, J., Resnick, R., Saget, B., Scheibel, S., Herman, S., Payne, H., Harrigan,
R., and Kwok, S. (1997) A rapid and simple method for extracting human immu-nodeficiency virus type 1 RNA from plasma: enhanced sensitivity.  J. Clin.
Microbiol. 35, 1278–1280.
28. Schockmel, G. A., Yerly, S., and Perrin, L. (1997) Detection of low HIV-1
RNA levels in plasma.  J. Acquir. Immune Defic. Syndr. Hum. Retrovirol. 14,
179–183.
29. Sun, R., Ku, J., Jayakar, H., Kuo, J. C., Brambilla, D., Herman, S., Rosenstraus,
M., and Spadoro, J. (1998) Ultrasensitive reverse transcription-PCR assay for
quantitation of human immunodeficiency virus type 1 RNA in plasma. J. Clin.
Microbiol. 36, 2964–2969.
30. Triques, K., Coste, J., Petter, J. L., Segarra, C., Mpoudi, E., Reynes, J., Delaporte,
E., Butcher, A., Dreyer, K., Herman, S., Spadoro, J., and Peeters, M. (1999) Effi-ciencies of four versions of the Amplicor HIV-1 Monitor test for quantification of
different subtypes of human immunodeficiency virus type 1. J. Clin. Microbiol.
37, 110–116.
31. DiDomenico, N., Link, H., Knobel, R., Caratsch, T., Weschler, W., Loewy, Z. G.,
and Rosenstraus, M. (1996) COBAS AMPLICOR: fully automated RNA and
DNA amplification and detection system for routine diagnostic PCR. Clin. Chem.
42, 1915–1923.
32. Longo, M. C., Berninger, M. S., and Hartley, J. L. (1990) Use of uracil DNA
glycosylase to control carry-over contamination in polymerase chain reactions.
Gene 93, 125–128.
412 Herman, Novotny, and Rosenstraus
33. Kwok, S. and Sninsky, J. J. (1993) PCR detection of human immunodeficiency
virus type 1 proviral DNA sequences, in  Diagnostic Molecular Microbiology
Principles and Applications (Persing, D. H., Smith, T. F., Tenover, F. C., and
White, T. J., eds.), ASM, Washington, DC, pp. 309–315.
34. Myers, T. W. and Gelfand, D. H. (1991) Reverse transcription and DNA ampli-fication by a  Thermus thermophilus DNA polymerase.  Biochemistry 30,
7661–7666.
Diagnosis of Hereditary Thrombotic Disorders 413
31
Molecular Diagnosis of Hereditary
Thrombotic Disorders
James G. Donnelly
1. Introduction
Deep vein thrombosis (DVT) can be the result of coagulation pathway
defects at the molecular level or damage to the vascular endothelium. Some of
the acquired causes of DVT include malignancy, trauma, prolonged immobili-zation, and pregnancy  (1) . Thrombophilia can be owing in part to both
acquired and inherited defects. The relative risk for thrombosis is increased by
estrogen replacement therapy  (2) and homocysteinemia  (3) . Homocysteine
metabolism is influenced by the use of alcohol, anticonvulsant drugs,
cyclosporine, methotrexate, inadequate dietary vitamin B12, folate and pyri-doxine intake, organ transplantation, and reduced creatinine clearance (4–8) .
Similar to venous thrombosis, homocysteinemia has genetic factors that influ-ence susceptibility (9–11) .
Numerous defects within the coagulation pathway have been observed.
Many mutations in the genes for protein C, protein S, and antithrombin III
have been described, making it impractical to use molecular techniques to
screen routinely for defects. Quantitative and functional assays are used
instead.
Three genes with common missense mutations have been found to add to
the relative risk of thrombosis (9–11) . Two of these gene mutations, factor V
1691AAG and prothrombin 20210GAA, have direct effects on the coagula-tion/anticoagulation pathways, and the enzyme expressed from the third gene,
methylenetetrahydrofolate reductase 677CAT (MTHFR C677T), influences
the conversion of homocysteine to methionine.
413
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
414 Donnelly
The detection of a genetic defect in factor V has shed light on the mecha-nism of the major cause of activated protein C resistance (APCR) (9) . A mis-sense mutation in the exon 10 coding region for the A2 domain of  factor V
1691AAG results in the substitution of arginine at residue 506 for glutamine.
This mutation has a dominant phenotype that results in the loss of an antigen-presenting cell (APC)-specific proteolytic site on factor Va required for
downregulation. This mutation is known as  factor VLeiden (9) and can be
detected using restriction digestion with MnlI. The frequency of factor VLeiden
is 5–7% in Western countries (9 ,12 ,13) . Moreover, APCR has been identified
in up to 40% of patients with DVT (9 ,13 ,14) .
The missense mutation, G20210A, within the 3vuntranslated region (UTR)
of the prothrombin gene also increases the risk of thrombosis (11) . This muta-tion is not specific for known restriction enzymes, and, therefore, a mutagenic
primer is used to create a HindIII site in the presence of the 20210A allele. The
frequency of this mutation is approx 1% in the general population (11) , 18% in
DVT patients with an established family history of venous thrombosis, and 6%
in consecutive patients with DVT  (11) . Individuals inheriting a copy of the
A20210 allele of the prothrombin gene have significantly elevated levels of
circulating prothrombin (11) . The G20210A mutation occurs downstream from
exon 14, the final coding region of the prothrombin gene. This mutation in the
prothrombin UTR may affect transcriptional regulation.
The missense mutation of MTHFR 677CAT results in a thermolabile variant of
this enzyme and reduced catalytic activity. Conversion of homocysteine to methi-onine is affected in individuals homozygous for the 677T allele. The frequency for
the MTHFR 677T allele is approx 0.38 in the Western Hemisphere (10). In one
study, 10% of French-speaking Canadians were shown to be homozygous (10).
The influence of this gene on homocysteine is related to folate concentration in
plasma. Fasting homocysteine concentrations in plasma are increased significantly
in 677T homozygotes when plasma folate concentrations are <15.4 nmol/L (15).
With this particular mutation, plasma homocysteine levels respond to increased
dietary folate. Hyperhomocysteinemia has been shown to contribute a two- to
threefold increase in risk for recurrent thrombosis  (16). This mutation can be
detected using the restriction enzyme Hinf I and amplified DNA.
Heritable factors for hypercoagulability such as underproduction of endog-enous anticoagulant proteins or increased synthesis of coagulant proteins will
result in increased risk of thrombosis. It is important to determine the heritable
risk of thrombosis to properly treat, anticipate future risk, and identify other
family members who may also be at risk. Identification of asymptomatic fam-ily members will permit these individuals to make informed decisions con-cerning choice of birth control method, estrogen therapy, and prophylactic
anticoagulant therapy in high-risk situations such as pregnancy and surgery.
Diagnosis of Hereditary Thrombotic Disorders 415
I present three polymerase chain reaction (PCR)-based genotyping assays
that can be performed simultaneously using the same master mix, DNA prepa-ration, and reaction conditions. Each assay can then be subjected to restriction
digestion and separation using agarose gel electrophoresis with visualization
using ethidium bromide (EtBr) staining. I also describe a second confirmatory
assay for factor VLeiden, using allele-specific oligonucleotide primers (17) .
A rare mutation of factor V at 1692AAC has been reported (18) . Although
this missense mutation does not confer resistance to APC, it has the same
digestion pattern as factor VLeiden. This 1692C allele has been reported in only
one individual and is likely a rare, isolated mutation. However, some laborato-ries may wish to confirm positive  factor VLeiden genotypes using the second
method. I also present algorithms that can be used to assist in appropriate assay
selection and, if followed, will minimize wasteful testing.
2. Materials
2.1. Isolation of Genomic DNA
1. Wizard™ Genomic DNA Purification Kit (Promega, Madison, WI).
2. Additional erythrocyte lysis fluid: 10 mM KCl, 10 mM NH4Cl, 10 mM
Tris-HCL, pH 7.5. Dissolve 0.75 g of KCl, 5.35 g of NH4Cl, and 1.21 g of Tris base
in 800 mL of deionized water. Adjust the pH to 7.50 with 1 M HCl. Adjust the
volume to 1.0 L with deionized water.
2.2. PCR Amplification
1. Primers (Applied Biosystems) as follows:
a. Factor VLeiden A1691G:
FV1 5v-ACCCACAGAAAATGATGCCCAG-3v.
FV2 5v-TGCCCCATTATTTAGCCAGGAG-3v(9) .
b. MTHFR C677T:
MTHFR1 5vTGAAGGAGAAGGTGTCTGCGGGA3v.
MTHFR2 5vAGGACGGTGCGGTGAGAGTG3v(10) .
c. Prothrombin G20210A:
UTPT1 5vTCTAGAAACAGTTGCCTGGC3v.
UTPT2 5vATAGCACTGGGAGCATTGAAGC3v(11) .
d. Factor VLeiden 1691G allele-specific confirmatory assay (optional):
FVASO1 5vCTTTCAGGCAGGAACAACACC3v.
FVASO2 5vTGGACAAAATACCTGTATACCTT3v(17) .
2. AmpliTaq™ DNA Polymerase Stoffel fragment 10X PCR buffer (Applied
Biosystems).
3. 10X Stock MgCl2 (25 mM) (Applied Biosystems).
4. AmpliTaq DNA Polymerase Stoffel fragment (10 U/µL) (Applied Biosystems).
5. GeneAmp™ dNTPs (10 mM stock solutions) (Applied Biosystems).
6. Perkin-Elmer 9600 Thermocycler (Applied Biosystems).
416 Donnelly
2.3. Restriction Enzyme Analysis
1. MnlI restriction enzyme (10 U/µL) (MBI Fermantis) for factor VLeiden.
2. Hinf I restriction enzyme (10 U/µL) (Promega) for MTHFR C677T.
3. HindIII restriction enzyme (10 U/µL) (Promega) for prothrombin G20210A.
4. EcoRI digested lambda phage DNA base pair marker for HindIII digest internal
control (Roche).
2.4. High-Resolution Agarose Gel Electrophoresis
1. Metaphor™ high-resolution agarose (FMC Bioproducts).
2. Agarose (Gibco-BRL).
3. 1X Tris-borate EDTA (TBE) (Sigma, St. Louis, MO): 100 mM Tris, 90 mM boric
acid, 1.0 mM EDTA, pH 8.33. Gel-Mix Running Mate™ TBE Buffer (Gibco-BRL) is used routinely in our laboratory rather than making this buffer from
individual components.
4. Gel loading solution: 50% (v/v) glycerol and deionized water with a few grains
of bromophenol blue stain.
5. EtBr solution (10 mg/mL).
3. Methods
Leukocyte (white blood cell [WBC]) nuclei serve as the source of DNA for
thrombophilic genotyping assays. The PCR assays described herein require the
sequential isolation of WBCs from red blood cells (RBCs). WBCs can be stored
at –70°C for extended periods. When ready for analysis, DNA is then isolated
from the WBC nuclei.
DNA is separated from the bulk of nuclear protein by differential precipita-tion of the proteins in a sodium dodecyl sulfate—ammonium acetate solution.
The DNA is then precipitated from the supernatant solution using 2 vol of
isopropanol. The pellet of DNA is washed with several volumes of 70% ethyl
alcohol to remove salts and detergent. The DNA pellet is then rehydrated with
DNA rehydration solution (TBE buffer). The method presented is modified
from the Wizard Genomic DNA Purification Kit and is intended to maximize
the quantity of DNA isolated. Using this procedure, enough DNA can be ob-tained from 2.5 mL of blood to perform between 20 and 50 separate PCRs.
This is useful for banking DNA for future studies as new gene mutations are
implicated in hereditary thrombosis.
3.1. Specimen Requirements
EDTA or citrate anticoagulated blood stored at room temperature for up to
4 d is suitable. Prepare the WBCs as soon as possible and store frozen. Hep-arinized whole blood has not been analyzed using these PCR genotyping meth-ods, and heparin is known to inhibit PCRs.
Diagnosis of Hereditary Thrombotic Disorders 417
3.2. Preparation of WBCs (see Note 1)
1. Centrifuge EDTA or citrate anticoagulated whole blood for 5 min at 1200g. The
blood should be stored at room temperature. Do not store the blood at 4°C for
prolonged periods. Glycolysis is inhibited in blood stored at 4°C and, therefore,
WBC integrity is compromised and hemolysis is increased.
2. While the tube is being centrifuged, pipet 900  µL of RBC lysis solution to a
labeled 1.5-mL microcentrifuge tube.
3. Using a disposable plastic pipet, remove the white cell layer (buffy coat) at the
plasma-cell interface and add it to the tube. Some plasma and RBCs will also be
transferred. Cap and mix.
4. Incubate the mixture at room temperature for 10 min. Invert two to three times
while incubating. Centrifuge for 1 min in a microcentrifuge to pellet the WBC
nuclei.
5. Remove the supernatant with a fine-tip disposable pipet. Remove as much of the
liquid as possible without disturbing the pellet. Occasionally RBCs do not lyse
completely. If this occurs, take the supernatant off and add RBC fresh lysis solu-tion to a total volume of 1.2–1.4 mL. Mix and reincubate for 10 min, centrifuge,
and remove the supernatant. Try to remove as much of the RBC stroma layer as
possible. At this point, the WBC pellet should be almost completely free of
hemoglobin. If it is not, step 5 can be repeated.
6. If the DNA is not to be prepared immediately, freeze and store the WBC pellets
at –70°C.
3.3. Preparation of DNA
1. Add 300 µL of the nuclei lysis solution to the tube containing the freshly thawed
and resuspended WBCs. Vortex vigorously or rub across the top of a tube rack to
ensure complete disruption of the pellet. The solution is very viscous at this point.
There must be no clumps of WBCs in the tube; otherwise the yield will be
reduced (see Notes 2–4).
2. Add 100  µL of protein precipitation solution to the nuclear lysate and vortex
vigorously for 10–20 s. If the solution does not show signs of a precipitate, and
instead has a viscous gel of nuclei, add another 100 µL of the protein precipita-tion solution and vortex again.
3. Centrifuge at 12,800g for 3 min at room temperature. A brown pellet should be
visible.
4. Using a disposable pipet, transfer the supernatant to a clean 1.5-mL micro-centrifuge tube containing 600 µL of isopropanol.
5. Gently mix the solution by inversion until white threads of DNA form. Should no
precipitate be obvious, shake the tube three or four times.
6. Centrifuge for 1 min at 12,800g at room temperature. The DNA will be a visible
white pellet. Using a permanent marker, place a line on the bottom of the tube
prior to centrifugation. Orient the tube in the centrifuge with the line facing outward.
After centrifugation, the DNA pellet will be visible against the mark on the tube.
418 Donnelly
7. Aspirate the supernatant and add 600 µL of 70% ethanol to the DNA. Invert the
tube gently to wash the pellet and the walls of the tube. Do not attempt to resus-pend the pellet. Centrifuge for 1 min at 12,800g at room temperature.
8. Aspirate the ethanol using a pipet. Repeat the ethanol wash once more. The DNA
pellet is not firmly attached to the tube and care must be taken to avoid aspirating
the pellet when removing the ethanol solution.
9. Air-dry the pellet to ensure that the ethanol is evaporated completely. Avoid
overdrying the pellet.
10. Reconstitute DNA pellet by adding 50 µL of TBE, DNA hydration solution to the
tube. Incubate at 65°C for 1 h on a heating block or at 4°C overnight. Genomic
DNA will not come back into the solution readily. It is better to prepare the DNA
the day before it is to be used. Mix well with a pipet before use, but do so gently
to avoid shearing the DNA.
11. Store the rehydrated DNA at 2–8°C and use within 3 to 4 d; DNA stability is
variable after 4 d. DNA is stable for several years when stored at –20°C.
3.4. Preparation of Working Primer Solutions
Deblocked primers are obtained from 40-nmol scale synthesis (see Notes
5 and 6).
1. Dilute primers in 200 µL of deionized water and further dilute a 5-µL aliquot of
this stock into 500 µL (1/100 dilution).
2. Test the diluted primer at three or four volumes, typically 2, 4, 6, and 8 µL in PCR
reactions. Then run out the reactions on a 2% agarose, 0.5X TBE (Gibco-BRL) gel.
3. Use the concentration that provides the optimal yield of PCR product to calculate
the desired dilution of the stock primer. For method consistency, the volume of
the primer is fixed at 2.5 µL in the PCR assays; therefore, the final dilution of the
stock will depend on the optimal volume of primer determined in the test.
3.5. PCR Assays for Factor VLeiden, MTHFR C677T,
and Prothrombin G20210A
A master reaction mixture is prepared separately for each gene using
reagents from Perkin-Elmer. Table 1 lists the volumes of each reagent required
for one tube. A Perkin-Elmer 9600 thermocycler is used for the reactions. The
amplification conditions are as follows:
1. Initial 5-min denaturation step at 95°C: Place the rack in the thermocycler when
the temperature is >85°C (see Note 7).
2. Thirty cycles of 94°C for 30 s, 50°C for 30 s, and 72°C for 1 min.
3. Final 5-min incubation at 72°C: This is particularly important for sharpening the
prothrombin PCR product band.
4. The optional confirmatory PCR assay for specimens positive for  factor VLeiden
uses the same master mix reagents (see Subheading 3.4.) with allele-specific
primers (see Subheading 2.2., step 1, part d), under the same reaction condi-tions as in steps 1–3 with 55°C used as the annealing temperature.
Diagnosis of Hereditary Thrombotic Disorders 419
3.6. Verification of PCR Product (optional)
1. Use a 2% Gibco-BRL agarose gel with 0.5X TBE and 0.5 µg/mL of EtBr. Add
10 µL of PCR product and 5 µL of gel-loading solution per well in the gel.
2. To the left lane of each gel run, add a 50-bp marker (2.5 µL) (MBI Fermentas).
3. Run at 100 V, in the same TBE buffer strength used to make the gel, until the dye
front has migrated approx 3 cm.
3.7. Restriction Digestion
1. Digest 10 µL of each of the specific PCR products with 1 U of MnlI, factor V, and
Hinf I for MTHFR, and 1 µ of HindIII for prothrombin (see Note 8).
2. As a restriction enzyme control, add EcoRI digested lambda phage (0.5 µg) to the
prothrombin reaction prior to digestion. Allow digestion to proceed for a mini-mum of 2 h at room temperature (see Note 9).
3.8. 2.5% High-Resolution Gel for Digest Products
For two 50-mL gels, mix the following components: 100 mL of 1X cold
TBE, 2.5 g of Metaphor agarose, 1.0 g of agarose. Before pouring the gel add
5 µL of EtBr (final concentration of 0.5 µg/mL from a 10 mg/mL stock).
1. Measure the buffer and transfer to a 500-mL Erlenmeyer flask. Weigh out the
agarose and add to the buffer while swirling the flask. Mix well and loosely cap
with a cotton gauze plug.
2. Microwave to melt the agarose. Periodically and carefully mix the solution to
avoid boil-over of the agarose. Continue heating until the agarose is completely
melted. Let the gel cool briefly and add 5 µL of EtBr stain. EtBr is considered
mutagenic, and gloves must be worn when handling the solution, gels, and buff-ers. Swirl the molten agarose until the EtBr is completely dispersed. Then pour
the gel into the casting stand.
Table 1
Preparation of PCR Master Mix
Master mix Volume (µL/tube) Final concentration
Distilled/deionized water 23 —
10X Stoeffel buffer   5 1X
10X MgCl2   5 2.5 mM
dNTPs   2 each 200 µM
Stoeffel Taq   0.25 2.5 U
Primer 1   2.5 20 µM
Primer 2   2.5 20 µM
.
420 Donnelly
3. One hundred milliliters of molten agarose is sufficient for two 5 × 8 cm mini gels.
Place one 12-well comb in each gel and leave undisturbed until completely
solidified. Refrigerate the gel for at least 30 min before using.
4. Cold 1X TBE should be used for the electrophoresis. Load a 50-bp DNA ladder
into the left-most lane and then 20 µL of digested product in the adjacent lanes.
5. Electrophorese the DNA fragments at 100 V until the dye front has migrated two
thirds to three fourths through the gel.
3.9. Separation of Restriction-Digested PCR Products
1. Separate the digested products using a gel consisting of 2.5% Metaphor agarose
and 1% agarose prepared with 1X TBE  (see Note 10).
2. Confirm positive  factor VLeiden in the optional allele-specific PCR assay by
amplifying a DNA product of 234 bp in size observed on a 2% agarose gel elec-trophoresed with 0.5X TBE and stained with EtBr. Prepare the gels in advance.
The gels may be stored in sealed plastic containers with a few milliliters of water
to keep them from drying out.
3.10. Documentation of Results
After running the gel for a suitable length of time as judged by the bro-mophenol blue stain, visualize the bands under UV light and document. We use a
Gel Doc™1000 (Bio-Rad, Hercules, CA) video capture system with Molecular
Analyst™ (Bio-Rad) software for documentation of gels (see Note 11).
3.11. Interpretation of Results
Restriction isotyping yields specific band patterns for each allele. The base
pair size of each fragment obtained after the restriction enzyme digestion is
determined from its position relative to the base pair markers. Figure 1 is rep-resentative of the typical digest separations using high-resolution agarose.
Table 2 lists the expected fragment size for each allele. Table 3 shows sug-gested work flow for these assays.
The allele patterns are superimposed for heterozygotes. The  prothrombin
20210G allele is not digested by HindIII. Therefore, EcoRI digested lambda
phage virus DNA is added prior to digestion. A faint band at 564 bp indi-cates successful digestion in the absence of the 20210A allele. Figure 2 illus-trates the optional allele-specific confirmation assay for  factor VLeiden (see
Notes 12–15).
MTHFR genotyping is useful for identifying hyperhomocysteinemic
patients who would benefit from folate supplementation. Approximately 28%
of our venous thrombotic patients are hyperhomocysteinemic (19) . Therefore,
homocysteine should be included in all initial patient assessments. Individuals
homozygous for T677 MTHFR may have normal or elevated tissue folate
Diagnosis of Hereditary Thrombotic Disorders 421
pools. However plasma folate, part of which is derived from hepatic portal
recirculation, is decreased. Jacques et al. (15) observed that homocysteine was
reduced when plasma folate concentrations were >15.4 nmol/L.
Hyperhomocysteinemic patients who are not homozygous for the  677T
allele of MTHFR, and who have folate and vitamin B12 values within the refer-ence interval and have no other acquired cause of impaired homocysteine
metabolism, should be screened for cystathionine beta synthase (CBS) defi-ciency. An abbreviated 2-h methionine-loading assay is suitable for screening
CBS deficiency (20) . Although a small percentage of individuals can have a
normal fasting homocysteine and be CBS deficient, we do not routinely per-form the methionine-loading test in our institution.
4. Notes
1. Other preparations for DNA such as buccal scrapings may be suitable for DNA.
However, most patients with DVT will be monitored for anticoagulant therapy or
require homocysteine and B vitamin monitoring. Therefore, WBC DNA is avail-able at some point in the normal course of treating these patients.
Fig. 1. Thrombophilia genotyping panel. Lanes A and I, 50-bp ladder; lane B, fac-tor V 1691AG (heterozygous Leiden); lane C, factor V 1691AA (wild type); lane D,
MTHFR 677CT; lane E, MTHFR 677TT (homozygous thermolabile variant); lane F,
MTHFR 677CC (wild type); lane G, prothrombin 20210GA; lane H, prothrombin
20210GG (wild type). The band at 564 bp is the digestion control using  EcoRI
digested lambda DNA. Inverted image (positive–negative) of EtBr-stained gel.
422 Donnelly
2. Smaller volumes can be used for DNA isolation. This preparation is optimized
for large yields of DNA. Banked DNA is useful for future studies concerning
thrombophilia.
3. The RBC lysis solution is similar to that supplied by Promega. Following the
procedure outlined, this reagent is limiting using the commercial kit, and there-fore it is economical to prepare this component.
4. Should the occasional PCR not work, repeat using one half or two times the quan-tity of DNA. If this does not solve the problem, reprecipitate the DNA in isopro-panol and wash the pellet again with 70% ethanol. We have not had this problem;
however, contamination with detergents could occur if the pellet were not washed
adequately.
5. Good quality primers from a reliable source are essential for successful PCR.
The optimization step will ensure that the primers work before attempting to pro-cess patient specimens.
6. It is not necessary to purchase purified primers. Deblocked primers are adequate.
7. The rack is not loaded on the thermocycler until the temperature reaches 85°C.
This is not a true hot-start technique. However, this does help prevent false prim-ing and ensure a more specific start to the amplifications.
8. The digestion can take place in the parent tube of DNA to save an aliquoting step.
9. The tubes can be left overnight if required.
10. The restriction digest products are easily separated by the outlined procedure
without the use of polyacrylamide gel electrophoresis. The use of agarose gels
greatly reduces the analysis processing time.
11. Detection of small DNA fragments is difficult without optimal staining and a
good photodocumentation system. Digital video capture is not required for
detection of small DNA fragments. However, running the gels longer than neces-sary to visualize the bands is not recommended, because the longer the gels are
electrophoresed the more the bands can diffuse through the gel.
Table 2
Interpretation of Restriction Digest
Gene Allele DNA restriction fragments (bp)
Factor V 1691 A 104, 82, 37
1691 G 141, 82
MTHFR 677 C 198
677 T 175
Prothrombin 20210 G 345a
20210 A 322
a Undigested PCR product. It requires an internal restriction enzyme control as outlined in
Subheading 3.
Diagnosis of Hereditary Thrombotic Disorders 423
12. The assays for heritable thrombophilia are expensive and utilization should be
monitored closely. The prevalence in the patient population is one way to test for
appropriate utilization. The prevalence of  factor VLeiden, prothrombin A20210,
and MTHFR TT677 in our patients is 21.5, 5.4, and 12.4%, respectively. The
prevalence of APCR is 50.3% in our tested population. Approximately 6% of our
patients cannot be tested for APCR because of anticoagulation or estrogen
therapy.
13. Wasserman et al. (21) and Adcock et al. (22) have described approaches to the
incorporation of  factor VLeiden genotyping into coagulation investigations. The
normalized APCR ratio was more sensitive than the APCR ratio when used as a
preliminary screen to eliminate unnecessary factor VLeiden genotyping (21) . The
sensitivity and specificity of the normalized ratio using a cutoff value of <0.85
were 100 and 51%, respectively, in Wasserman et al.’s study (21) . The APCR
ratio cutoff of <1.57 was more specific (79%); however, sensitivity decreased to
94%. In the absence of preliminary screening for APCR, we found that approx
21.5% of our DVT patients were heterozygous for factor VLeiden. APCR results
Fig. 2. Optional confirmation assay for factor VLeiden. Lane A, 50-bp marker; lanes
B and E, allele-specific primers for  factor V 1691A; lanes C and D, allele-specific
primers for  factor V 1691G. Lanes B and C indicate 1691AG, and lanes D and E
indicate 1691AA. Allele-specific primer for 1691A is 5vTGGACAAAATACCTGT-ATACCTT3v. This primer does not need to be used when the allele-specific oligo-nucleotide assays are used for confirmation.
424 Donnelly
are an economical screening tool for factor VLeiden genotyping. Care must be taken
to ensure that the patient history is accurate and communicated to the laboratory
because the APCR assay will be abnormal irrespective of factor V genotype when
a patient is anticoagulated, pregnant, or taking estrogens.
14. A mutation of factor V A1692C that destroys the MnlI restriction site but does not
confer APC resistance has been reported in one patient  (18) . It appears to be
unlikely that this mutation is common. However, some laboratories may wish to
confirm the results of restriction assay for factor VLeiden until they are reasonably
certain that the mutation is not present in their patient population.
15. We chose not to use the allele-specific oligonucleotide assay as our primary
assay because of the uncertainty of amplification and the cost of running two
assays per patient. As a primary assay, two sets of primers are required for factor
V: the wild-type forward primer and the allele-specific reverse primer. Figure 2
shows the results of both reactions. Additionally an internal control must be used
in this assay to confirm the integrity of the PCR should this be selected as a
stand-alone assay for  factor V genotyping. However, even with the use of an
internal control product, there is no guarantee that the allele-specific primers for
factor V have amplified the target DNA. Moreover, this assay also is not cost-effective because two PCR tubes must be set up for each assay and two sets of
primers; the factor V primers and the internal control primer must be used when
this assay is run alone.
Table 3
Suggested Work Flow for Thrombophilia Genotyping
Day 1 Days 2 and 3
Prepare DNA from fresh and frozen Remove PCR tubes from thermocycler
leukocytes in the morning.       and aliquot for digest.
Prepare master mix and run Prepare digest working enzyme solution.
amplification in the afternoon. Add EcoRI digested lambda to the
prothrombin aliquots.
Add working enzyme solution to the
aliquots.
Prepare high-resolution electrophoresis
gels and refrigerate for a minimum of
30 min after the agarose has set.
Load high-resolution agarose gels with
50-bp DNA ladder and digested PCR
products.
Separate at 100 V or less in cold 1X TBE.
Record digest results.
Diagnosis of Hereditary Thrombotic Disorders 425
Acknowledgments
I am indebted to Ferne Shirley for her superior organizational skills and
many contributions to the thrombophilia genotyping procedures.
References
1. Appleby, R. D. and Olds, R. J. (1997) The inherited basis of venous thrombosis.
Pathology 29, 341–347.
2. Helmerhorst, F. M., Bloemenkamp, K. W., Rosendaal, F. R., and Vandenbroucke,
J. P. (1997) Oral contraceptives and thrombotic disease: risk of venous throm-boembolism. Thromb. Haemost. 78, 327–333.
3. D’Angelo, A., Mazzola, G., Crippa, L., Fermo, I., and Vigano, D. S. (1997)
Hyperhomocysteinemia and venous thromboembolic disease. Haematologica 82,
211–219.
4. Cravo, M. L., Gloria, L. M., Selhub, J., et al. (1996) Hyperhomocysteinemia in
chronic alcoholism: correlation with folate, vitamin B-12, and vitamin B-6 status.
Am. J. Clin. Nutr. 63, 220–224.
5. Lucock, M. D., Wild, J., Schorah, C. J., Levene, M. I., and Hartley, R. (1994) The
methylfolate axis in neural tube defects: in vitro characterisation and clinical
investigation. Biochem. Med. Metab. Biol. 52, 101–114.
6. Arnadottir, M., Hultberg, B., Vladov, V., Nilsson, E. P., and Thysell, H. (1996)
Hyperhomocysteinemia in cyclosporine-treated renal transplant recipients. Trans-plantation 61, 509–512.
7. Keuzenkamp, J. C., De, A. R., Blom, H. J., Bokkerink, J. P., Trijbels, J. M. (1996)
Effects on transmethylation by high-dose 6-mercaptopurine and methotrexate
infusions during consolidation treatment of acute lymphoblastic leukemia.
Biochem. Pharmacol. 51, 1165–1171.
8. Brattstrom, L., Lindgren, A., Israelsson, B., Andersson, A., and Hultberg, B.
(1994) Homocysteine and cysteine: determinants of plasma levels in middle-aged
and elderly subjects. J. Intern. Med. 236, 633–641.
9. Bertina, M. R., Koeleman, B. P. C., Koster, T., et al. (1994) Mutation in blood
coagulation factor V associated with resistance to activated protein C. Nature 369,
64–67.
10. Frosst, P., Blom, H. J., Milos, R., et al. (1995) A candidate genetic risk factor for
vascular disease: a common mutation in methylenetetrahydrofolate reductase. Nat.
Genet. 10, 111–113.
11. Poort, S. R., Rosendaal, F. R., Reitsma, P. H., and Bertina, R. M. (1996) A com-mon genetic variation in the 3v-untranslated region of the prothrombin gene is
associated with elevated plasma prothrombin levels and an increase in venous
thrombosis. Blood 88, 3698–3703.
12. Koster, T., Rosendaal, F. R., de Ronde, H., Briet, E., Vandenbroucke, J. P., and
Bertina, R. M. (1993) Venous thrombosis due to poor anticoagulant response to
activated protein C: Leiden Thrombophilia Study. Lancet 342, 1503–1506.
13. Svensson, P. J. and Dalback, B. (1994) Resistance to activated protein C as a basis
for venous thrombosis. N. Engl. J. Med. 330, 517–522.
426 Donnelly
14. Griffin, J. H., Evatt, B., Wideman, C., and Fernandez, J. A. (1993) Anticoagulant
protein C pathway defective in majority of thrombophilic patients.  Blood 82,
1908–1993.
15. Jacques, P. F., Bostom, A. G., Williams, R. R., et al. (1996) Relation between
folate status, a common mutation in methylenetetrahydrofolate reductase, and
plasma homocysteine concentrations. Circulation 93, 7–9.
16. den Heiger, M., Blom, H. J., Gerrits, W. B., et al. (1995) Is hyperhomo-cysteinaemia a risk factor for recurrent venous thrombosis?  Lancet 345,
882–885.
17. Ridker, P. M., Hennekens, C. H., Lindpaintner, K., Stampfer, M. J., Eisenberg,
P. R., and Miletich, J. P. (1995) Mutation in the gene coding for coagulation fac-tor V and the risk of myocardial infarction, stroke, and venous thrombosis in
apparently healthy men. N. Engl. J. Med. 332, 912–917.
18. Liebman, H. A., Sutherland, D., Bacon, R., and McGehee, W. (1996) Evaluation
of a tissue factor dependent factor V assay to detect factor V Leiden: demonstra-tion of high sensitivity and specificity to a generally applicable assay for activated
protein C resistance. Br. J. Haematol. 95, 550–553.
19. Donnelly, J. G. (1998) Homocysteine and plasma folate relationship in patients
with venous thrombosis, end-stage renal disease or atherosclerosis. Clin. Chem.
44(Suppl. 6), A167,A168.
20. Bostom, A. G., Roubenoff, R., Dellaripa, P., et al. (1995)Validation of abbrevi-ated oral methionine-loading test. Clin. Chem. 41, 948,949.
21. Wasserman, L. M., Edson, J. R., Key, N. S., Chibbar, R., and McGlennen, R. C.
(1997) Detection of the factor V Leiden mutation: development of a testing algo-rithm combining a coagulation assay and molecular diagnosis. Am. J. Clin. Pathol.
108, 427–433.
22. Adcock, D. M., Fink, L., and Marlar, R. A. (1997) A laboratory approach to the
evaluation of hereditary hypercoagulability. Am. J. Clin. Pathol. 108, 434–449.
RhD Genotyping by Allele-Specific PCR 427
32
Prenatal Genotyping of the RhD Locus
to Identify Fetuses at Risk for Hemolytic
Disease of the Newborn
Martin J. Hessner and Daniel B. Bellissimo
1. Introduction
Hemolytic disease of the newborn (HDN) can occur when there are
fetomaternal incompatibilities within any number of different erythrocyte anti-gen systems, including the RhD, Cc, Ee, Kidd and Duffy, and Kell antigen
systems. In these disorders, maternal antibodies are developed by
alloimmunization of the mother to fetal red blood cells during pregnancy when
the fetal cells carry an alloantigen inherited from the father. The maternal anti-bodies result in the destruction of fetal erythrocytes leading to severe hemolytic
anemia and hyperbilirubinemia. Permanent neurologic damage can result from
HDN, and in extreme cases loss of the fetus or death of the neonate may occur.
In subsequent pregnancies, it is important to determine the status of the incom-patible allele in the fetus. If the father is heterozygous or homozygous for the
allele, the chance of the fetus inheriting the paternal alloallele to which the
mother is immunologically sensitized is 50 or 100%, respectively. Fetuses that
do not inherit the alloallele will not be at risk for HDN.
Investigative and therapeutic measures used for alloimmunized pregnant
women involve some risk to the fetus. Currently, women who present with
alloantibody titers to red cell antigens are monitored by amniotic fluid spectro-photometric analysis to detect deviation from linearity at 450 nm, the wave-length at which bilirubin absorbs (1) . Accurate determination of fetal risk is
achieved through serial analysis, generally weekly for several weeks. Although
427
From: Methods in Molecular Medicine, vol. 49: Molecular Pathology Protocols
Edited by: A. A. Killeen © Humana Press Inc., Totowa, NJ
428 Hessner and Bellissimo
the risk of placental trauma during amniocentesis has been greatly reduced
since the introduction of ultrasound imaging techniques, there is still a 2% risk
of placental trauma (2) . Alternatively, percutaneous umbilical blood sampling
allows direct measurements of all fetal blood parameters including blood
groups. However, the procedure is technically more difficult and because of
the risk of fetomaternal hemorrhage and further sensitization of the mother, its
use is limited. The molecular characterization of many blood antigen systems
has enabled the development of molecular diagnostic assays that are useful in
identifying fetuses at risk for HDN. Prenatal identification of the relevant geno-types for fetuses potentially at risk for HDN requires fetal DNA isolated from
a single amniocentesis. When fetuses are shown to be compatible with sensi-tized mothers, and therefore not at risk for HDN, the need for expensive and
invasive monitoring throughout the pregnancy can be obviated.
The characterization of the  RhD and RhCcEe genes has provided the
molecular basis for prenatal testing for RhD. The Rh locus on chromosome 1
contains two distinct but highly homologous genes: RhD and RhCcEe (3,4) .
Generally, RhD-positive individuals possess one or two copies of the RhD gene
and two copies of the RhCcEe gene, whereas RhD-negative individuals retain
two copies of the RhCcEe gene but typically lack the RhD gene (4) . The RhD
and RhCcEe genes are arranged tandemly and are believed to have arisen
through duplication of a single ancestral gene. The Rh genes, which are >95%
homologous at the nucleotide sequence level, both consist of 10 exons span-ning more than 75 kb, and both encode peptides of 417 amino acid residues
with a predicted molecular mass of 30–35 kDa (5–8) .
A number of sequence differences between the RhCcEe and RhD genes can
be utilized in genotyping assays to identify the presence or absence of the RhD
gene. Polymerase chain reaction (PCR)-based RhD typing assays, utilizing
sequence differences in intron 4, exon 3, exon 7, and the untranslated region
within exon 10, have been previously described (7 ,9–12) . However, discrepan-cies between serotyping and genotyping have been observed. These discrepan-cies are largely owing to the existence of allelic variants, the molecular basis of
which is often the result of recombination between the RhD and RhCeEe genes.
In these hybrid genes, some RhD sequences are replaced with the correspond-ing sequences from the  RhCcEe gene (9 ,13) . For other variants, all that is
known is that the sequences targeted by PCR primers or restriction enzymes
are altered or deleted. These variants can potentially cause the misdiagnosis of
the fetal RhD status during prenatal genotyping. To develop a reliable tech-nique for prenatal RhD typing of fetuses at risk for HDN, we have evaluated
the suitability of four different regions of the  RhD gene for genotyping and
compared these results to those obtained by serology.
RhD Genotyping by Allele-Specific PCR 429
The first RhD genotyping method is based on multiplexing the oligonucle-otide primers of Arce et al. (7) with oligonucleotide primers derived from those
described by Bennett et al. (10) . The RhD genotyping strategy described by
Arce et al. (7) detects a 600-bp deletion within intron 4 of the RhD gene that is
not present within intron 4 of the  RhCcEe gene (10) . Amplification with a
primer pair that targets exons 4 and 5 of the RhD and the RhCcEe genes results
in a 1200-bp RhCcEe product and a 600-bp product in RhD-positive individu-als. The RhD-specific oligonucleotide primers derived from those described
by Bennett et al. (10) specifically amplify a 193-bp product of the 3vuntrans-lated region of exon 10. The second RhD genotyping method evaluated
involves specific amplification of a 96-bp product from exon 7 of the  RhD
gene, previously described by Simsek et al.  (12) . Finally, the third method
involves specific amplification of a 111-bp product from exon 3 of the RhD
gene as previously described by Beckers et al. (9) .
To evaluate these genotyping methods and their possible discrepancies with
serologic typing, 50 Rh-phenotyped individuals (38 RhD positive and 12 Rh
negative) were selected. The RhD-positive samples were chosen for their
unusual serotyping results suggesting the samples were RhD variants. DNA
was isolated from the peripheral blood, and these samples were genotyped
using the aforement