Understanding Aging

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 TM
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 TM
Humana Press
Edited by
Yvonne A. Barnett
Christopher R. Barnett
Edited by
Yvonne A. Barnett
Christopher R. Barnett
and Protocols
and Protocols
Understanding Aging 1
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Understanding Aging
Bernard L. Strehler
1. Background
Enormous advances in our understanding of human aging have occurred
during the last 50 yr. From the late 19th to the mid-20th centuries only four
comprehensive and important sources of information were available:
1. August Weismann’s book entitled  Essays on Heredity and Kindred Biological
Problems (the first of these essays dealt with The Duration of Life; 1). Weissmann
states (p. 10) “In the first place in regulating the length of life, the advantage to
the species, and not to the individual, is alone of any importance. This must be
obvious to any one who has once thoroughly thought out the process of natural
2. A highly systematized second early source of information on aging was the col-lection of essays edited by Cowdry and published in 1938. This 900+ page vol-ume contains 34 chapters and was appropriately called Problems of Aging.
3. At about the same time Raymond Pearl published his book on aging  (2). Pearl
believed that aging was the indirect result of cell specialization and that only the
germ line was resistant to aging. Unfortunately Pearl died in the late 1930s and is
largely remembered now for having been the founding editor of Quarterly Review
of Biology while he was at the Johns Hopkins University, this author’s alma mater.
4. Alexis Carrel wrote a monumental scientific and philosophical book, Man, the
Unknown (3). Carrel believed that he had demonstrated that vertebrate cells could
be kept in culture and live indefinitely, a conclusion challenged by others (more
on this later).
Probably the most useful of all the more recent books published on aging
was Alex Comfort’s The Biology of Senescence (4), which supplied much of
the source information that this author used in writing Time, Cells and Aging
(5–7; I am most grateful to Dr. Christine Gilbert, of Cyprus, for her efforts in
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the revision of the third edition of  Time, Cells and Aging, and for the most
stimulating discussions we have had over the years). The extremely useful and
thoroughly documented book called Developmental Physiology and Aging by
Paul Timeras (8) is a fine source of critical appraisals of the science in both
areas. Many of the more recent books on aging are cited later. The success of
my own journal (Mechanisms of Ageing and Development) is largely due to the
work of our excellent editorial board and to the careful work and prodding of
my dear wife, Theodora Penn Strehler, who passed away on 12 February, 1998.
This chapter is dedicated to her living memory and the love she gave to me for
50 years of marriage and joy and sadness — and the kindness she showed to all
who knew her. Requiescat in pacem.
2. Overview of a Systematic Approach
My own synthesis and analysis of the nature and causes of aging were pre-sented in a book called  Time, Cells and Aging. To use terms consistently in
discussing aging, a set of four properties that all aging processes must meet are
defined in that book:
1. Aging is a process; i.e., it does not occur suddenly, but rather is the result of very
many individual events.
2. The results of aging are deleterious in the sense that they decrease the ability of
an individual to survive as he or she ages.
3. Aging is universal within a species. However, aging may not occur in every spe-cies. Thus, certain “accidents” such as those that result from a specific infection
are not part of the aging process.
4. Aging is intrinsic to the living system in which it occurs (i.e., it reflects the quali-ties of DNA, RNA, and other structures or organelles that were inherited from the
parental generation).
The central thesis presented in  Time, Cells and Aging is that the possible
causes of aging can be divided into:
1. Those that are built into the system as specific DNA or RNA coding (or catalytic)
sequences, and
2. Those that are the result of controllable or uncontrollable environmental factors
including radiation, nutrition, and lifestyle.
Two key phenomena are shown by aging animals:
1. The probability of a human dying doubles about every 8 yr, a fact that was first
discovered by an English Insurance Actuary by the name of Benjamin Gompertz
about 165 yr ago  (9). Thus, the following equation, derived from Gompertz’s
work, accurately describes the probability of dying as a function of age in a par-ticular environment: R = k + R0eat where, R(ate) of death at any age equals the
probability of dying at age 0 multiplied by an age-dependent factor that is equal
Understanding Aging 3
to e raised to the a times t power, where a is a function of the doubling time and
t is the age attained. A better fit to observed mortality rates is given by adding a
constant (k) (which largely reflects environmental factors).
If one plots log R against t(age) one obtains a remarkably precise straight line,
usually between ages 30 and 90. A Gompertz curve is obtained for the mortality
rate vs age for a variety of animals—humans, horses, rats, mice, and even Droso-phila melanogaster, a much studied insect.
2. A second general fact or law is provided by my own summary and analysis of the
pioneering quantitative work of Nathan Shock on maximum functional ability of
various body systems’ ability to do work as humans age. Shock’s studies (on
humans) implied to me that after maturity is reached the following equation
describes a multitude of maximum work capacity of various body parts: Wmax =
Wmax(30) (1 – Bt) where B varies from about 0.003 per yr to almost 0.01 per yr—
depending on the system whose maximal function is being measured. For exam-ple, maximum nerve conduction velocity declines by about 0.003 per yr (10) and
vital capacity as well as maximum breathing capacity declines by about 1% per
yr (11).
The Gompertz and Shock equations pose the following puzzling and key
question: “How can a linearly declining ability in various functions cause a
logarithmic increase in our chances of dying as we age ?” A probable answer to
this question was provided by this author in collaboration with Prof. Albert
Mildvan (12–14). Our theory made two assumptions. The first of these is that
the equation derived from Shock’s work (that the maximum work capacity of a
variety of body systems declines linearly after maturity is reached) is valid.
This, as shown earlier, is the very simple equation: Wmax = Wmax(30) (1 – Bt),
where Wmax is the maximum ability to do work at age t, Wmax(30) is the maxi-mum ability to do work at age 30, where B is the fraction of function lost per yr,
and t is the age in years. Of course B varies from species to species and the t
term is some small fraction of the maximum longevity of a species.
The second assumption is that the energy distribution of challenges to sur-vival is very similar to the kinetic energy distribution of atoms and molecules
as defined in the Maxwell–Boltzmann equation. This equation or law defines
how kinetic energy is distributed in a collection of atoms or molecules at a
specific temperature (where temperature is defined as the average kinetic
energy and is equal to KE = 0.5 mv2). This distribution has a maximum value
near the average kinetic energy of the particles in the system. But higher and
higher energies are generated through random successive multiple collisions
between particles. The reason that this is possible is easily understood through
an analogy in which the particles are seen as billiard balls. Consider the case
when one of two spherical billiard balls can absorb momentum from another
such sphere. This happens in billiards when one ball strikes the second ball
squarely. In that case, the moving billiard ball stops and the formerly stationary
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one moves off at about 45° from the direction in which the first one was moving.
The law of conservation of momentum is mv = K for any two colliding structures.
Because the balls are not perfectly elastic some heat will be generated during
the collision, but this is a very small fraction of the total momentum and kinetic
energy of the two particles. This is evident from the fact that one cannot feel a
warming of either of the billiard balls after such a collision and the fact that the
ball that is struck moves at about the same velocity that the first ball had before
the two balls collided. Now consider the special case where two such billiard
balls are traveling at right angles to each other when they collide and that the
collision between them is “on center” so that one of the balls stops dead in its
tracks and the other ball moves off at a 45° angle at a speed that conserves total
momentum. (That is, the moving ball is now moving along the line that defined
the center of gravity of the two balls as they were moving before they collided.)
If momentum the two balls is conserved (the momenta are added) then the
speed of the struck moving ball should be twice that which both of the balls
had before they collided. There is no obvious reason why momentum is not
conserved in this manner. But the kinetic energy (1/2)mv2 of the moving ball
will be much greater than the sum of the kinetic energies they had before colli-sion. (In fact the total kinetic energy of the two balls moving at the same veloc-ity before they collided is  two times as great after they collide than it was
before this special kind of collision happened!) This is a most surprising seem-ing “violation” of the Law of Conservation of Energy. It would seem to follow
from this that certain kinds of very improbable collisions result in an increase
in the kinetic energy of the pair of balls. This seems almost obvious from the
fact that the kinetic energies of atoms or molecules is not equal among atoms
or molecules in a closed system. Instead, it follows the Maxwell–Boltzmann
distribution. Where does this energy come from? Perhaps from the Einsteinian
conversion of mass to energy. Thus it appears that if one constructs a device
in which collisions of the non-random kind described previously took place
one should be able to get more energy out of the system than one puts in—
essentially because the structure of such a machine minimizes the entropy of
collisions by causing only certain very rare collisions to take place. I have spent
many months testing this revolutionary theory, but the results produced from
my “Perpetual Motion Machine” have failed to demonstrate any such gain in
kinetic energy. There appears to be no other explanation for the distribution of
kinetic energy among atoms and molecules than the kind of collisions discussed
here! It’s unfortunate that it doesn’t work at the macro level. In any event, if a
small probability exists that improbable collisions, such as discussed previ-ously, are rare and cause an increase in momentum of one of the balls or atoms
then the probability that a series of similar collisions that increase momentum
of particular atom or molecule will give that atom or molecule greater and
Understanding Aging 5
greater energy will decrease very rapidly as the number of such improbable
events increases. In fact, the number of such combined events will decrease
logarithmically as the energy possessed by such an atom or molecule increases
linearly. Such a decreasing exponential is part of the classical form of the Max-well–Boltzmann equation—and defines the number of atoms with momenta
greater than some particular high value. In fact, the distribution of momentum
is described by a symmetrical bell-shaped curve (a Maxwellian curve) whereas
the distribution of energy follows the Maxwell–Boltzmann curve.
To return to the Gompertz equation as it applies to the probability of dying
vs age, Mildvan and I postulated that the energy distribution of challenges to
living systems is very similar to the Maxwell–Boltzmann distribution. For
example. obviously one knows that small challenges such as cutting a finger or
tripping or stumbling are very frequent compared to the chance of falling down
the stairs, being hit by a speeding automobile, or experiencing an airplane crash.
Similarly, the frequency of coming down with a very serious diseases (infec-tions by a new influenza virus, blood clots in the coronary arteries or key arter-ies in the brain, aortic aneurysms, cancer) is much rarer than is coming down
with a minor infection (e.g., a cold or acne) or bumping one’s shin against a
coffee table. It may have been that the “Sidney” flu somehow was exported
from Hong Kong to Australia by a “carrier” passenger in an airplane and thence
to the Uunited States via another carrier who gave it to someone who infected
my great grandson, who in turn infected our entire family at Christmas time,
1997 and led to my sadness at losing the person, Theodora (my wife), I had
deeply loved and enjoyed for 50 years. The separate events leading to this per-sonal tragedy were each improbable, but they resulted in a very large challenge
that one of us was unable to overcome! This illustrates the principle that it
takes many unlikely events to lead to a major challenge to humans—or to
The theory of absolute reaction rates states that R = C(kt/h)e–(F*/RT), where
F* is the free energy of activation of a reaction. The free energy of activation is
in turn defined as the amount of energy needed to break a bond that must be
broken in order for a chemical reaction to occur. Of course the free energy
needed is derived from multiple collisions and the number of particles that
possess a given excess energy equal to that required for a given reaction to
occur increases as a function of the absolute temperature. Note that the RT (gas
constant times absolute temperature) leads to an exponentially decreasing rate
of reaction as T (absolute temperature) is lowered linearly because the T term
is in the dividend of the negative exponential term e–(F*/RT). If one plots the log
of the rate against 1/T one obtains a straight line whose slope is a measure of
the minimum amount of energy (T*) required to cause a reaction to happen.
Such a plot is called an Arrhenius plot. Therefore, if one defines the events that
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lead to possible death similarly and takes into account the linear decline in the
body’s ability to resist challenges (through the expenditure of the right kind of
energy in a particular system or systems) decreases linearly as we age, one
obtains the Gompertz equation. Thus, the Gompertz equation results from the
logarithmic distribution of size of challenges we encounter interacting with
linear loss of functions of various kinds during aging observed by Shock.
3. Ten Key Experimental Questions—Plus Some Answers
Although several hundred specific questions or theories regarding the
source(s) of aging in humans and other nucleated species (eukaryotes) are pos-sible, only 10 of the most carefully examined “theories” are highlighted here.
Space does not permit a complete discussion of each of these questions.
1. How does the temperature of the body affect the rate of aging?
The activation energy of a particular chemical reaction is the amount of
energy that is derived from accidental collisions among atoms or molecules to
break the bonds needed for the reaction to occur. If the reaction is a catalyzed
one then the activation energy is about 10–20 kcal/mol. By contrast, if the reac-tion is not catalyzed the energy required is that which will break a bond in a
reacting substance. Covalent bonds require between 75 and 130 kcal to be bro-ken, whereas in the presence of an appropriate catalyst the bond is weakened
by its combination with the catalyst so that it only takes 6–20 kcal to break it. If
one plots the log of the rate of a reaction against the reciprocal of the absolute
temperature one often obtains a remarkably straight line. Such a plot is called
an Arrhenius plot (after the man who discovered it). The slope of the straight
line obtained in such a plot will generally be high (50–200 kcal for uncatalysed
reactions and 6–19 kcal for catalyzed ones. In order to calculate the activation
energy of aging I plotted my own results on the effects of temperature in Droso-phila life-spans (15,16) together with those of Loeb and Northrup (17,18) and
others and found the activation energy to be between 15 and 19 kcal. Thus, in
the cold-blooded animal, Drosophila (a fruit fly), the rate of aging appears to
be determined by a catalyzed reaction or possibly by the effects of temperature
on the rates of production and destruction of harmful substances such as .OH
radicals that attack DNA and other cell parts. It is known that trout live much
longer in cold lakes than in warmer ones but no quantitative studies of their
longevities at a variety of temperatures have, to my knowledge, been made.
Because mammals operate at essentially constant body temperatures, it is not
an easy matter to study the effect of body temperatures on humans or similar
mammals. One might find a correlation between the body temperatures of the
descendants of centenarians and the descendants of shorter lived persons, but
such a study is unlikely to be funded (as I know from personal experience!).
Understanding Aging 7
2. Are changes in connective tissue a key cause of aging?
There is no doubt the age-related alterations to the structure and therefore
biological properties of connective tissues can lead to cosmetic through to
pathological changes in vivo. The onset of such pathologies may in some
instances increase the chances of death.
It is widely recognized that changes in the elasticity of skin (less elasticity)
as we grow older occurs in humans. If one pinches the skin on the back of the
hand and pulls up on it, it returns to its original shape (flat) in a short time,
about 1 s for young persons and about 3 s or more for older skin. This change is
primarily due to the attrition of the elastic fibers that are present in the dermis.
If the skin is exposed during early life to large amounts of ultraviolet radiation
such as that in sunlight, some of the collagen is converted into a fiber that
resembles elastin. This transformation leads to the uneven contraction of the
skin, that is, wrinkles are formed. The collagen in the skin and elsewhere in the
body becomes less plastic as it matures (for a discussion of the chemical pro-cesses underlying these maturity changes please see 19–23). Alteration in the
physical properties of the elastic tissue found in blood vessels can lead to
changes in blood pressure in vivo.
There are many examples of pathologies that result from age-related alter-ations to connective tissues. Particularly in fair-skinned persons, exposure to
ultraviolet light can lead to damage of skin cells and may lead to basal cell and
squamous cell cancers (both of which are relatively easily treated) and even
melanomas (difficult to treat successfully if not diagnosed at very early stages).
Alterations to the structure of bone can lead to osteoporosis. Physical changes
to the cartilage in joints can lead to the onset of osteoarthritis.
3. Does a significant fraction of the mitochondria of old mammals suffer
from defects, either in DNA or in other key components?
The mitochondria we possess are all derived from our mother’s egg, as are
various other materials such as particular RNA molecules. Mitochondria are
the cell factories in which the energy provided when food is oxidized is con-verted into the unstable molecule called ATP. ATP is used to contract muscles,
to pump ions across neural membranes, and is used to manufacture proteins
and RNAs.
The production of ATP can be assayed (24–26; John Totter and I (at the
Oak Ridge National Laboratory in 1951) developed an assay for ATP using
McElroy’s reaction (24) that is able to measure a billionth of a gram of ATP
(1 millionth of a milligram). This method has been widely used in various bio-logical and biomedical studies but the description of the method was published
so many years ago (1951–52) that it is no longer associated with our names. In
my laboratory we used this assay to study the production of ATP by mitochon-
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dria obtained from animals of different ages. We found no differences between
mitochondria from 8-mo-old rat hearts and 24-mo-old rat hearts, using α-keto-glutaric acid as substrate. Later it was reported that some mitochondria from
old animals oxidize different substrates such as succinate less efficiently than
do mitochondria derived from young animals. Later in this book Miquel et al.
summarize the literature, including much of their own work, on various mor-phological and functional changes that accumulate with age in mitochondria.
These changes are thought to result from an accumulation of various types of
mutations in the mitochondrial genome (much of which codes for polypeptides
involved in Complex I and II of the respiratory redox chain) that result from
primarily reactive oxygen species damage to the mitochondrial genome that is
poorly, if at all, repaired. Turnbull et al. present two chapters later in this book
on the analysis of mitochondrial DNA mutations. Such an age-related decrease
in mitochondrial function has been proposed to lead to the bioenergetic decline
of cells and tissues and so contribute to the aging process (27).
4. Is a limitation in the number of divisions a body cell can undergo (in cell
culture) a significant cause of aging?
Alexis Carrel reported (3) that he was able to keep an embryonic chicken
heart alive for more than 22 yr. This is, of course, much longer than chickens
usually live and Carrel concluded that regular supplements of the growth me-dium with embryo extracts would keep these cultures alive for very long times,
perhaps indefinitely. To quote from p. 173 of the Carrel book, “If by an appro-priate technique, their volume is prevented from increasing, they never grow
old.” Colonies obtained from a heart fragment removed in January 1912, from
a chick embryo, are growing as actively today as 23 yr ago. In fact, are they
immortal? Maybe so. For many individuals, including myself at about 13 yr of
age, these findings were very exciting. Perhaps man would eventually be able
to conquer his oldest enemy, aging. It was at about that time that I decided on a
career in aging research.
In 1965 my good friend Leonard Hayflick reported some research he and a
colleague (Moorhouse) had carried out that appeared to be contrary to what the
renaissance man, Carrel, had concluded (28). Hayflick found that human fibro-blasts in a culture medium could go through only about 50 doublings, after
which the cells died or stopped dividing (now known as replicative senescence)
or both. Hayflick’s data have been confirmed by many persons, including this
author, who with Robert Hay (29) carried out similar experiments on chicken
fibroblasts that were only capable of about 20 doublings. However, because a
new layer of skin cells is produced about every 4 d (about 90 doublings per yr
and 9000 doublings in a 100-yr lifetime), and because red blood cells are pro-duced by the millions every 120 d and because the crypt cells in the lining of
Understanding Aging 9
the intestine give rise to the entire lining of the cleft in which the crypt cells lie,
it seemed to me unreasonable that the Hayflick limit applies to normal cells in
the body. In the case of skin cells Hayflick countered with the idea that if each
of the progenitor cells in the skin could divide only 50 times, then the reason
might be that cells moved out of the dividing cell structure (the one cell thick,
basal cell layer) that gives rise to the epidermis after they had gone through 40
or 50 doublings. This seemed a reasonable and possibly correct theory, so (with
the help of my late wife), we showed that the cells did not leave the basal layer
two or four or eight cells at a time, but rather the daughter cells of cells labeled
with tritiated thymine moved out of the basal layer randomly (the reader is
encouraged to read pp. 37–55 of the third edition of Time, Cells and Aging for
further discussion in this regard). Such a finding may cast strong doubt on the
relevance of in vitro clonal “aging” to the debilities of old age.
I offer one possibility that may account for the apparent contradiction
between the findings of Carrel on one hand and of Hayflick on the other. The
antibiotics routinely used during the “fibroblast cloning” experiments (and
other experiments performed since on the phenomenon of replicative senes-cence) might in themselves cause a decrease in the number of divisions pos-sible. Carrel was unable to use antibiotics in his studies because they were not
yet discovered or manufactured when he carried out his 22-yr experiment on
chick heart viability. Hayflick states in his recent book that he has evidence
that Carrel’s embryo extract supplements contained living cells and that this is
why the tissues Carrel studied remained alive for times greater than the life-time of a chicken. Carrel had to use very careful means to replace his media
every so often over a period of 20 yr. Besides, Carrel did not allow his organ
cultures to grow, so cell division was either absent or cells possibly present in
the embryo extracts he added were able to differentiate into replacement cells
for heart tissues. Because the heart is a syncytium of cells, it is difficult to
imagine how a steady state of replacement of old cells by cells possibly present
in the embryo extract could take place, particularly within the center of the
organ culture! This logic argues for the validity of Carrel’s reports. Moreover,
fibroblasts are quite different from myoblasts and do not form syncytia.
In very recent times a popular proposal has been that telomeres, the
sequences of noncoding DNA located at the end of chromosomes, shorten each
time a normal cell divides and that in some way this shortening “counts” the
number of cell divisions that a cell population has experienced, perhaps owing
to the loss of essential genes that have critical functions for cell viability
(30,31). What is not clear is how the documented process of replicative senes-cence in vivo leads to the development of physiological malfunction and the
onset of age-related pathologies in vivo. Changes in the expression of a num-ber of gene functions, including increases in the expression of genes coding for
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growth factors and extracellular matrix components, have been found by study-ing cells in replicative senescence in vitro. Researchers have been able to detect
relatively small numbers of senescent fibroblasts and epithelial cells in older
animals and human tissues in vivo using  β-galactosidase staining (pH > 6).
They have postulated that even such small numbers of cells, exuding various
entities because of activated genes etc., might be sufficient to alter tissue
homeostasis and so lead to physiological effects. This suggestion has yet to
be proven and the role of replicative senescence in aging remains an area of
intense research activity.
5. Are errors in the transcription and/or translation of DNA a key source of
aging? Or, alternatively, are changes in the rate of transcription or translation
of the information in DNA a key cause?
Medvedev (32) was the first to propose that the stability of DNA was respon-sible for the length of life of different species. Orgel then proposed his “error
theory of aging” in which he proposed that errors in DNA replication, tran-scription of RNA, and translation on the products might be responsible for the
deterioration of function during aging (33). Over a number of years a major
effort was made in this author’s laboratory to test the idea that development
and aging were caused by changes in the specific codons different kinds of
cells were able to translate. Initial studies showed that the aminoacylated
tRNA’s for a variety of amino acids differed from one kind of cell to another
and a theory called the “Codon Restriction Theory of Development and Aging”
was published in  Journal of Theoretical Biology  (34). The theory was then
tested against the actual codon usage of about 100 different messenger RNAs
and it was indeed found that certain kinds of gene products (e.g., the globin
parts of hemoglobins) do in fact have very similar patterns of codon usages and
codon dis-usages in messages ranging from birds (chickens) to mice and rats to
humans! On these bases, the inability to translate specific codons in specific
kinds of tissues may indeed turn out to be important in the control of gene
expression (at least in some tissues).
6. Are changes in RNA qualities responsible for aging?
Whether the kinds of RNA present in cells is important in controlling differ-entiation and aging is an issue that has arisen when it was discovered that cer-tain RNA molecules possess catalytic activity, e.g., are able to generate
themselves by catalytically transforming their precursors (35). I have recently
read evidence that even the transfer of growing polypeptide chains to the amino
acid on the a tRNA to the “next” position is catalyzed in the ribosomes by a
particular kind of RNA. Whether changes in catalytic RNA populations cause
certain disabilities during aging has not yet been tested, to my knowledge.
Understanding Aging 11
7. Do long-lived cells selectively fail in humans?
The answer to this question is certainly yes. The main sites in which clear
age changes take place are in cells that cannot be replenished without a disrup-tion in their functions in the body. Key cell types are neurons, heart muscle,
skeletal muscle, and certain hormone producing cells. The important precursor
of both androgens and estrogens, DHAE, declines linearly with age in men and
women and may well be a product of cells that are not replenishable. But even
more obvious is the postmitotic nature of cells in the nervous system and other
nonreplenishing tissues such as skeletal and heart muscle. Thus, damage to the
cells making up these organs generally cannot be repaired through replacement
because such postmitotic cells cannot be made to divide. In the case of
the brain, continual replacement of old cells by new ones might preserve reflex
brain function, but most such newly incorporated nerve cells would replace
neurons in whose facilitated synapses useful memories had been stored. Thus,
paradoxically, higher animals, particularly humans, age because some key
kinds of cells they possess have long, but not indefinitely long, lifetimes.
(Although it is fairly obvious I would like it to be called “The Strehler Para-dox,” so that way I might be remembered for something unless a different
version of the perpetual motion machine I proved unworkable actually gener-ated useful energy!)
8. What are the underlying causes of the age-related decline in the immune
The immune system consists of two major forms: innate and acquired. Innate
immunity comprises polymorphonuclear leukocytes, natural killer cells, and
mononuclear phagocytes and utilizes the complement cascade as the main soluble
protein effector mechanism. This type of immunity recognizes carbohydrate struc-tures that do not exist on eukaryotic cells; thus foreign pathogens can be detected
and acted against. Lymphocytes are the major cells involved in the system of
acquired immunity, with antibodies being the effector proteins. The T-cell receptor
(TCR) and antibodies recognize specific antigenic structures.
Deterioration of the immune system with aging (“immunosenescence”) is
believed to contribute to morbidity and mortality in man due to the greater
incidence of infection, as well as possibly autoimmune phenomena and cancer
in the aged. T lymphocytes are the major effector cells in controlling patho-genic infections, but it is precisely these cells that seem to be most susceptible
to dysregulated function in association with aging.
Decreases in cell-mediated immunity are commonly measured in elderly
subjects. By most parameters measured, T-cell function is decreased in elderly
compared to young individuals. Moreover, prospective studies over the years
have suggested a positive association between good T-cell function in vitro and
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individual longevity. The numbers and/or function of other immune cells are
also altered with age: antigen-presenting cells are less capable of presenting
antigen in older age; the number of natural killer cells increases in older age, and
these cells are functionally active; there is some evidence that granulocyte func-tion may be altered with age; B lymphocyte responses also alter with age, as
responses against foreign antigens decline whereas responses against self-anti-gens increase (36,37). Currently much effort is being directed toward elucidat-ing the processes leading to the phenomenon of immunosenescence. The reader
is encouraged to read a special issue of Mechanisms of Ageing and Develop-ment that was dedicated to publishing the proceedings of a recent international
meeting on immunosenescence (38).
One positive aspect of immunosenescence, however, is that the risk of trans-plant rejection is reduced with age.
9. Are ordinary mutations a major cause of aging? Or, alternatively, is the
instability of tandemly repeated DNA sequences a major cause of aging?
In 1995 a Special issue of Mutation Research entitled “Somatic Mutations
and Ageing: Cause or Effect?” was published, with an overview from this
author highlighting the history of this field of science (39).
Much of the early results from the experiments on the effects of ionizing
radiation and chemical mutagens on the life-span of Drosophila and other ani-mals were inconsistent with a simple mutation theory of aging. However, the
research papers presented in the special issue of Mutation Research, and else-where, do suggest an involvement of somatic and mitochondrial mutation in
the physiological and pathological decline associated with the aging process. I
also believe that some other kind of DNA change, the occurrence of which was
not accelerated by radiation proportionally to dose (as are ordinary mutations),
could be responsible for aging. This kind of postulated change in DNA might
well occur sufficiently frequently, even in unirradiated animals, to cause aging!
In humans the nucleolar organizing regions (NORs), which can be detected
by silver staining, are regions containing rDNA which is the template on which
rRNA is formed. There are about five or six pairs of chromosomes that possess
such NOR regions. It has been shown that the number of NORs decreases with
time in a variety of human cells. Perhaps, I thought, losses of such tandemly
duplicated regions takes place at a relatively high rate in nondividing human
cells during aging, but is not appreciably increased by exposure to moderate
amounts of radiation. After all, radiation affects all kinds of DNA and the rDNA
genes may well be able to repair most of the damage they receive either during
aging or as a result of chemical or electromagnetic radiation such as UV light
and X-rays or by neutrons. I postulated that mutations that cause the loss of
rDNA might be responsible for human aging because the more severe such loss
Understanding Aging 13
is, the greater should be the loss of function of any cell in manufacturing pro-teins. Such mutations could be the kind that cause the linear decrease in func-tion of various parts of the body observed by Shock. Although I thought this
unlikely to occur, particularly in postmitotic cells, we were eager to disprove it,
because loss of important genetic material would be very difficult to reverse
(e.g., through the use of a “clever” virus), whereas a defect in the regulation of
gene expression which had been the focus of our research should require sim-pler, but presently unknown, treatments to modify the rate of aging—which at
that time seemed to be on the horizon.
To test the possibility that rDNA loss is a major cause of aging, I asked a
very talented postdoctoral trainee, the late Roger Johnson, to work to study the
rDNA content of various mammalian tissues. He owned a small airplane that
made it possible for him to fly to Davis, California to obtain a variety of tissues
of control beagle dogs of different ages that were killed as part of an ongoing
study by the Atomic Energy Commission to determine the pathological effects
of radiation. We obtained fresh samples from the following organs: brain, heart,
skeletal muscle, kidney, spleen, and liver. When we compared the rDNA con-tent of the brains of beagles of various ages we found that the results were not
what we had hoped for and expected—namely, that no difference would be
found between young and old animals. Instead, the findings were that the rDNA
content decreased by about 30% in brains of dogs from approx 0–10 yr of age
(40). We then proceeded to compare the effect of age on the DNA of heart,
skeletal muscle, kidneys, spleen, and liver. Decreases in rDNA of about the
same magnitude were found in the other two postmitotic tissues, heart and
skeletal muscle, but were not detected in liver DNA or kidney DNA. A small,
probably insignificant, loss of these gene sequences was detected in dog spleens
(41,42). After the work on dogs was completed, we began to study human heart
and found a substantial loss of rDNA of aged humans (43). We later studied
two different areas of the human brain, the somatosensory cortex and the hip-pocampus. The fresh autopsy samples were kindly supplied by the Los Ange-les coroner. We discovered that the rate of loss of rDNA from human brain and
heart was about 70% per 100 yr. This rate is only about 1/7th of the rate
observed in dogs and thus is inversely proportional to the maximum longevity
of these two species (approx 120 yr and approx 16 yr). The ratio of these two
life-spans is very close to 7:1 and the ratio of loss of rDNA/yr is about 1:7. The
two parts of the human brain measured were almost identical in their rDNA
content, although the loss was of course greater in old tissues than in young
ones. This indicates that the measurements are reliable or at least that, if errors
were made, the errors must be very small. Over a period of about 10 yr we
continued to publish studies on humans. Most of these studies were reported in
Mechanisms of Aging and Development.
14 Strehler
A very interesting class of mutants in  Drosophila are called the  minute
mutants. My former dear friend Kimball Atwood (who has departed to the great
genetics lab in the sky) noted that there are many different minute mutants and
that they are found in various places on essentially all of the four chromosomes
this animal possesses. He suggested that the mutants might reflect the loss of at
least part of the tRNA coding regions for specific tRNAs. I don’t know whether
this hypothesis has been critically tested—if it hasn’t it certainly should be.
Deficiency (but not total absence of specific tDNAs that decode specific amino
acids) would be expected to interfere with the normal growth rate of all parts of
the developing fly embryo—hence the name, minute.
10. What causes Alzheimer’s disease and cancers—and what means are now
available to control these tragic diseases of the elderly (and of certain younger
persons as well)?
I spent considerable time and effort recently studying another major sci-entific question: Is a specific temporal code used in transmitting, decoding,
and storing information (memories) in the mammalian brain? I had pub-lished a theory on this concept in Perspectives in Biology and Medicine in
1969 (44). Knowledge of such a coding system could be quite interesting
and probably useful in understanding the familial forms of Alzheimer’s dis-ease. I studied the patterns in time of nerve discharges in response to spe-cific stimuli to the eyes of monkey brains. In the meantime I had constructed
an electronic memory system I thought might mimic the brain. I made some
progress and wrote a program that serially mimicked how I thought the brain
might store and recognize patterns. I also constructed an electronic analogue
that worked quite well. But, I made little progress in obtaining a clear answer
regarding the validity of my hypothesis until a brilliant French scientist, Dr.
Remy Lestienne, wrote to ask whether he could spend a year working with
my “group” (at that time only me!). After we had worked together for only a
month we discovered that the brain really did produce extremely precise
copies of doublets, triplets, quadruplets, and even sextuplets of pulses. Then
we analyzed various parameters, including the decay time for the occur-rence of repeating patterns. The patterns we used were precisely repeated
with variances between copies of the same pattern of less than 1/7th of a
millisecond for each of the three intervals that make up a pattern. This was
most surprising, because the duration of a nerve impulse is about 1 ms. Per-haps the most important discovery we made was that each repeating triplet
was surrounded by about seven doublets that were part of the repeating pat-tern and equally precisely replicated. Thus, we had not disproved my theory,
but rather found evidence that it was probably correct, at least for short-term
Understanding Aging 15
While this research was going on I also developed an electronic simulation
of the basic concepts and obtained a U.S. Patent on this device in 1993. I also
received a second patent that proposed a means to recognize different vowels
on the basis of the differences in logarithms of frequencies generated within
the mouth and nasopharyngeal cavities. Because the absolute frequencies that
children and women and men use to produce vowels are quite different a puzzle
existed as to how different vowels are understood despite the fact that the abso-lute frequencies generated are much different from person to person. A large-scale implementation of the content addressable temporal coding has not been
implemented although a very simple version was constructed by me and an
improved version was created by a most ingenious Japanese engineer named
Yuki Nakayama (sponsored by my friend H. Ochi, who has a consuming inter-est in aging research and is quite wealthy.) Perhaps the very new CD recorder
that Sony has recently marketed may be modified to construct a new and inex-pensive way to implement a device able to store the 1014 bits the human brain
evidently can store and retrieve upon proper cueing.
Alzheimer’s disease is manifested by the loss of memory, initially that
involving the recent past. One can remember minuscule details of the more
distant past, but sometimes forgets what day of the week it is and what one
wanted to get from the kitchen when one gets there. This realization of defects
in remembering recent events can be quite disconcerting to those of us who
have enjoyed the use of memory, logic, and analogy in solving scientific prob-lems and important problems generated by the process of getting older.
Alzheimer’s is also called presenile dementia, which means that it can occur as
early as the late 40s or 50s, long before other signs of senility manifest them-selves. As the disease progresses victims may even lose the ability to recognize
family members or even their spouses or their own names. When the brains of
persons who die of various diseases are autopsied, it is possible to recognize
those who have advanced stages of Alzheimer’s degeneration by looking for
the many “plaques” characteristic of Alzheimer’s. Similar plaques are found
in the brains of essentially all very elderly persons, but they are markedly more
numerous in the brains of true inheritors of the acute form of this age change in
brain anatomy—persons with Alzheimer’s. The plaques are visible on the sur-face of the brain and consist of localized patches of changed brain tissue vis-ible to the naked eye. When the plaques are examined microscopically at least
three characteristics are obvious: (1) the plaques contain many dead or dying
cells; (2) most of the cells that are still alive in a plaque possess long tangles of
fibers that are not found in profusion in “normal” neurons elsewhere in the
same individual’s brain biopsy; and (3) the cells are surrounded by very large
accretions of antibody-like substances called amyloid. These deposits often
encase the entire cell body of a neuron. It is important to note that these amyloid
16 Strehler
deposits are evidently different from most other kinds of amyloid found in the
brain and elsewhere. The key difference appears to be that a cleavage product
of the amyloid characteristic of Alzheimer’s causes cell death by opening Ca2+
channels in the neural membranes’ neuroreceptor regions. This causes a per-manent depolarization of the cells and evidently is the cause of their death and
the loss of memories the cells or cell groups store. The most exciting research
on this subject of which I am aware is that the drug Flupirtine now used in
Europe for the treatment of Alzheimer’s, reportedly with some success, pre-vents the influx of Ca2+ into cells that are pretreated with this substance when
Alzheimer’s amyloid is presented to them. This work, recently published in
Mechanisms of Ageing and Development, was carried out by my good friend,
Werner Mueller, who will become Editor-in-Chief of the journal when I cease
my editorial responsibilities at the end of this year (45). I believe this is the
most significant finding to be published on possible treatment of a very sad
disease of the elderly.
Cancer is a common cause of morbidity and mortality in the elderly. The
spectrum of the major types of cancers occurring in the early years of life (leu-kemias and sarcomas) is different from that occurring in later life (carcinomas
and lymphomas). The most frequent cancers in women in Western societies are
breast, ovarian, and colorectal, and in men prostate, lung, and colorectal. The
multistep theory of carcinogenesis (46) predicts the age-related increased risk
(5th power of age in both short-lived species such as rats and long-lived species
such as humans) for the development of a wide range of different types of
cancer (with the exception of the familial forms of the disease). The underlying
molecular cause of cancer is the accumulation of mutations within a number of
genes associated with the control of cell growth, division, and cell death.
Despite the great variety of cells that can give rise to cancer there are now
somewhat effective treatments for many of them (surgery, radiotherapy, and/or
chemotherapy). Optimal treatment for many cancers is more likely the earlier
the diagnosis is made. Among the most promising of new treatments for some
cancers is the use of radioactively labeled antibodies to the surface antigens
present on some cancer cells but not on normal cells. The labeled antibody
seeks out the surface of the cancer cell and the radioactivity attached to it selec-tively radiates and destroys the tumor cells. Another recent treatment that
appears to have at least some success is the use of substances that prevent angio-genesis, thereby effectively “asphyxiating” the dangerous tumor.
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University Press, Clarendon, London, and New York.
Understanding Aging 17
2. Pearl, R. (1921) The biology of natural death, public policy and the population
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3. Carrel, A. (1935) Man the Unknown, Harper, New York.
4. Comfort, A. (1956) The Biology of Senescence, Rinehart, New York.
5. Strehler, B. L. (1962) Time, Cells and Aging, 1st ed., Academic Press, New York.
6. Strehler, B. L. (1977) Time, Cells and Aging, 2nd ed., Academic Press New York.
7. Strehler, B. L. (1999) Time, Cells and Aging, 3rd ed., Master Print Demetriades
Bros., Cyprus.
8. Timeras, P. (1972) Decline in homeostatic regulation, in Developmental Physiol-ogy and Aging (Timeras, P. S., ed.), MacMillan Press, New York, pp. 542–563.
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10. Norris, A., Shock, N., and Wagman, I. (1953) Age changes in the maximum con-duction velocity of motor fibers of human ulnar nerve. J. Physiol. 5, 589–593.
11. Shock, N. and Yiengst, M. (1955) Age changes in basal respiratory measurement
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Biology of Aging  (Strehler, B. L., et al., eds.), Publ. No. 6, Am. Inst. Biol. Sci.,
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14. Strehler, B. L. and Mildvan, A. (1960) A general theory of mortality and aging.
Science 132, 14–21.
15. Strehler, B. L. (1961) Studies on the comparative physiology of aging. II. On the
mechanism of temperature life-shortening in Drosophila melanogaster. J. Gerontol.
16, 2–12.
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melanogaster. J. Gerontol. 17, 347–352.
17. Loeb, J. and Northrop, J. H. (1916) Is there a temperature coefficient for the dura-tion of life? Proc. Natl. Acad. Sci. USA 2, 456.
18. Loeb, J. and Northrop, J. H. (1917) On the influence of food and temperature on the
duration of life. J. Biol. Chem. 32, 103–121.
19. Sinex, F. M. (1964) Cross linkage and aging. Adv. Gerontol. Res. 1, 167–178.
20. Houuck, G. Dehesse, C., and Jacob, R. (1967) The effect of aging upon collagen
catabolism. Symp. Soc. Exp. Biol. 21, 403–426.
21. Franzblau, C. and Lent, R. (1968) Studies on the chemistry of elastin. Brookhaven
Symp. Biol. 21, 358–377.
22. Gallop, P., Blumenfeld, O., Henson, E., and Schneider, A. (1968) Isolation
and identification of alpha amino aldehyde and collagen.  Biochemistry  7,
23. Franzblau, C. Fariz, P., and Papaioannou, R. (1969) Lysenonorleucine. A new amino
acid from hydrolysate of elastin. Biochemistry 8, 2833–2837.
18 Strehler
24. McElroy, W. (1947) The energy stored for bioluminescence in an isolated system.
Proc. Natl. Acad. Sci. USA 33, 342–345.
25. McElroy, W. and Strehler, B. L. (1949) Factors influencing the response of the
bioluminescent reaction to ATP. Arch. Biochem. 22, 420–433.
26. Strehler, B. L. and Totter, J. R. (1962) Firefly luminescence in the study of energy
transfer mechanisms. 1. Substrate and enzyme determination.  Arch. Biochem.
Biophys. 40, 28–41.
27. Fleming, J. E., Miquel, J., Cottrell, S. F., Yenguyan, L. S., and Economas, A. S.
(1982) Is cell aging caused by respiration dependent injury to the mitochondrial
genome? Gerontology 28, 44–53.
28. Hayflick, L. (1965) The limited in vitro lifetime of human diploid strains. Exp. Cell
Res. 37, 614–638.
29. Hay, R. L., Menzies, R. A., Morgan, H. P., and Strehler, B. L. (1967) The division
potential of cells in continuous growth as compared to cells subcultured after main-tenance in stationary phase. Exp. Gerontol. 35, 44.
30. Chang, E. and Harley, C. B. (1995) Telomere length and replicative aging in human
vascular tissues. Proc. Natl. Acad. Sci. USA 92, 11,190–11,194.
31. Bodnar, A. G., Ouellette, M., and Frolnis, M. (1998) Extension of lifespan by intro-duction of telomerase into normal human cells. Science 279, 349–352.
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34. Strehler, B. L., Hirsch, G., Gusseck, D., Johnson, R., and Bick, M. (1971) The
codon restriction theory of aging and development. J. Theor. Biol. 33, 429–474.
35. Andron, L. and Strehler, B. L. (1973) Recent evidence on tRNA and tRNA acylase-mediated cellular control mechanisms. A review. Mech. Ageing Dev. 2, 97–116.
36. Pawelec, G. and Solana, R. (1997) Immunosenescence. Immunol. Today 18, 514–516.
37. Pawelec, G., Remarque, E., Barnett, Y., and Solana, R. (1998) T cells and aging.
FIBS 3, 59–99.
38. Special edition of Mechanisms of Aging and Development (1998), 102.
39. Special issue of Mutation Research (1995) Somatic Mutations and Aging: Cause or
Effect? 338, 1–234.
40. Johnson, R. and Strehler, B. L. (1972) Loss of genes coding for ribosomal RNA in
aging brain cells. Nature (Lond.) 240, 412–414.
41. Johnson, R., Chrisp, C., and Strehler, B. L. (1972) Selective loss of ribosomal RNA
genes during the aging of post-mitotic tissues. Mech. Ageing Dev. 1, 183–198.
42. Johnson, L. K., Johnson, R. W., and Strehler, B. L. (1975) Cardiac hypertrophy,
aging and changes in cardiac ribosomal RNA gene dosage in man. J. Mol. Cell.
Cardiol. 7, 125–133.
43. Strehler, B. L., Johnson, L. K., and Chang, M. P. (1979) Loss of hybridizable ribo-somal DNA from human postmitotic tissues during aging: I Age-dependent loss in
human myocardium. Mech. Age. Dev. 11, 371–378.
44. Strehler, B. L. (1969) Information handling in the nervous system: an analogy to
molecular genetic coder-decoder mechanisms. Perspect. Biol. Med. 12, 548–612.
Understanding Aging 19
45. Perovic, S., Böhm, M., Meesters, E., Meinhardt, A., Pergande, G., and Müller, W.
F. G. (1998) Pharmacological intervention in age-associated brain disorders by
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46. Vogelstein, B. and Kinzler, N. W. (1993) The multistep nature of cancer. TIG 9,
Fibroblast Model for Cell Senescence Studies 23
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Use of the Fibroblast Model in the Study
of Cellular Senescence
Vincent J. Cristofalo, Craig Volker, and Robert G. Allen
1. Introduction
In this chapter, we present standard procedures for the culture of human
cells that exhibit a finite proliferative capacity (replicative life-span). The use
of a cell culture model has the advantage of providing a controlled environ-ment to study a wide variety of cellular phenomena. It also has the inherent
limitation of isolating cells from the regulatory elements that might be pro-vided by other types of cells in vivo. Nevertheless, cell culture models have
been crucial to our current understanding of mechanisms of growth, differen-tiation, development, and neoplasia and numerous other disease states. In this
chapter we present procedures for human fibroblast culture including serum-free cultivation of cells, which is necessary when the cellular environment must
be fully defined. In addition, we present procedures for the determination of
replicative life-span, saturation density, and assessment of replicative capacity
from labeled thymidine incorporation in fibroblasts. The methods described
here have been well tested and provide highly reproducible results (1,2).
1.1. Cellular Senescence
Phenotypically and karyotypically normal human cells exhibit a limited
capacity to proliferate in culture (3,4). This finite proliferative potential of nor-mal cells in culture is thought to result from multiple changes (5) and has fre-quently been used as one model of human aging. Although most replicative
life-span data are derived from fibroblasts, other types of cells such as glial
cells (6), keratinocytes  (7), vascular smooth muscle cells  (8), lens cells  (9),
endothelial cells (10), lymphocytes (11), liver (12), and melanocytes (13) are
also known to exhibit a limited replicative life-span in culture. Both environ-
24 Cristofalo, Volker, and Allen
mental and genetic factors appear to influence the proliferative life-span of
fibroblasts from normal individuals (5,14,15). Not all of the determinants of
proliferative capacity are known; however, a variety of changes are associated
with the decline of proliferative capacity including changes in gene expression,
telomere shortening, and signal transduction. These are all thought to be impor-tant factors that influence replicative life-span (15–20).
1.1.1. Telomere-Shortening
Loss of telomeric repeats is tightly linked to the cessation of mitotic activity
associated with cellular senescence  (16,17,21,22). The telomeres of human
chromosomes are composed of several kilobases of simple repeats
(TTAGGG)n. Telomeres protect chromosomes from degradation, rearrange-ments, end-to-end fusions, and chromosome loss (23). During replication DNA
polymerases synthesize DNA in a 5′ to 3′ direction; they also require an RNA
primer for initiation. The terminal RNA primer required for DNA replication
cannot be replaced with DNA, which results in a loss of telomeric sequences
with each mitotic cycle (21,23). Cells expressing T antigen are postulated to
exhibit an increase in their proliferative life-span because they are able to con-tinue proliferating beyond the usual limit imposed by telomere length  (24).
Immortalized and transformed cells exhibit telomerase activity that compen-sates for telomere loss by adding repetitive units to the telomeres of chromo-somes after mitosis  (23,25–27). Cultures derived from individuals with
Hutchinson–Gilford syndrome  (28) often exhibit decreased proliferative
potential, albeit results with these cell lines are variable (29). Fibroblast cul-tures established from individuals with Hutchinson–Gilford progeria syndrome
that exhibit a lower proliferative capacity than cells from normal individuals
also exhibit shorter telomeres; however, the rate of telomere shortening per cell
division appears to be similar in progeria fibroblasts and normal cells (16). It
has recently been demonstrated that proliferative senescence can be delayed
and possibly eliminated by transfection of normal cells with telomerase to pre-vent telomere loss (30). It is also interesting to note that other repetitive DNA
sequences become shorter during proliferative senescence (31,32)
1.1.2. Mitogenic Responses and Signal Transduction
As a result of senescence-associated changes, cells assume a flattened mor-phology and ultimately cease to proliferate in the presence of serum  (5).
Numerous factors may contribute to the senescent phenotype; however, the
principal characteristic of cellular senescence in culture is the inability of the
cells to replicate DNA. Paradoxically, the machinery for DNA replication
appears to remain intact, as indicated by the fact that infection with SV-40
initiates a round of semiconservative DNA replication in senescent cells (33).
Fibroblast Model for Cell Senescence Studies 25
Nevertheless, senescent cells fail to express the proliferating cell nuclear anti-gen (PCNA), a cofactor of DNA polymerase δ, apparently as a result of a post-transcriptional block  (34). Furthermore, senescent fibroblasts fail to
complement a temperature-sensitive DNA polymerase α mutant (35,36). This
may contribute to the failure of senescent cells to progress through the cell
cycle because it is known that a direct relationship exists between the concen-tration of DNA polymerase α and the rate of entry into S phase (37). It has also
been observed that replication-dependent histones are also repressed in senes-cent cells and that a variant histone is uniquely expressed (18).
It might also be noted that the senescence-dependent cessation of growth is
not identical to G0 growth arrest that occurs in early passage cells that exhibit
contact inhibited growth or that are serum starved. Several lines of evidence
suggest that senescent cells are blocked in a phase of the cell cycle with many
characteristics of late G1. For example, thymidine kinase is cell cycle regu-lated; it appears at the G1/S boundary. Thymidine kinase activity is similar in
cultures of proliferating young and senescent WI-38 cells  (38,39). It should
also be noted that thymidine triphosphate synthesis, which normally occurs in
late G1, is not impaired in senescent cells (39). Furthermore, the nuclear fluo-rescence pattern of senescent cells stained with quinacrine dihydrochloride is
also typical of cells blocked in late G1 or at the G1/S boundary  (33,40). In
addition, Rittling et al.  (41) demonstrated that 11 genes expressed between
early G1 and the G1/S boundary are mitogen inducible in both young and senes-cent cells. On the other hand, growth-regulated genes such as cdc2, cycA, and
cycB, which are expressed in G1, are repressed in senescent cells (42). These
observations suggest the possibility that senescent cells are irreversibly arrested
in a unique state different from the normal cell cycle stages.
As cells approach the end of their proliferative potential in culture they
become increasingly refractory to mitogenic signals  (15,43,44). The signal
transduction pathways that convey these mitogenic signals play significant
roles in the regulation of cell proliferation and adaptive responses; hence,
decline in the activity of elements in these pathways may contribute signifi-cantly to the senescent phenotype. For example, there is a senescence-associ-ated loss in the capacity of cells to activate protein kinase C (45) or to increase
interleukin-6 (IL-6) mRNA abundance (46) following stimulation with phorbol
esters. Furthermore, transcriptional activation of c-fos following stimulation of
cultures with serum is also diminished in senescent cells (18,47). Other genes
such as  Id1 and Id2, which encode negative regulators of basic helix–loop–
helix transcription factors, fail to respond to mitogens in senescent cells (48)
Although signal transduction efficiency declines with replicative age, the
members of affected pathways are seldom influenced uniformly by senescence.
For example, both the number of receptors (per unit cell surface area) and
26 Cristofalo, Volker, and Allen
receptor affinities for epidermal growth factor (EGF), platelet-derived growth
factor (PDGF), and insulin-like growth factor-one (IGF-one) remain constant
throughout the proliferative life of fetal lung WI-38 fibroblasts (49–51); how-ever, senescent WI-38 cells produce neither the mRNA nor the protein for IGF-I
(52). Similarly, young and senescent WI-38 fibroblasts have similar baseline
levels of intracellular Ca2+ and exhibit similar changes in cytosolic Ca2+ fluxes
following growth factor stimulation  (53); however, the expression of
calmodulin protein is uncoupled from the cell cycle and exists in variable
amounts in senescent WI-38 cells (53). The calmodulin-associated phosphodi-esterase activity also appears to be diminished in late-passage cells (Cristofalo
et al., unpublished results). At least some of the changes in signal transduction
associated with senescence may also stem from alterations in the cellular redox
environment, because the rate of oxidant generation increases during senes-cence (54) and some steps in various signal transduction pathways are highly
sensitive to changes in redox balance. The protein abundances of protein kinase
A (PKA) and various isoforms of protein kinase C (PKC) are unchanged or
slightly increased by senescence (20,55); however, PKC translocation from the
cytoplasm to the plasma membrane is impaired in senescent fibroblasts (45,56).
Changes in signal transduction efficiency associated with senescence are
not necessarily the result of any decrease or loss of components of signaling
pathways. Experiments performed in various types or immortal and normal
cells reveal that increases in signal transduction components can also impede
signaling pathways. This is most clearly seen in the case of the extracellular
signal-regulated kinase (ERK) pathway where the correct sequence and dura-tion of activation and inactivation of ERKs at the G1/S boundary  (57–59) is
required for entry into S phase. Indeed, constitutive ERK activation has an
inhibitory effect on cell cycle progression, both in NIH 3T3 fibroblasts (58)
and in Xenopus oocytes (60). Furthermore, overexpression of oncogenic ras in
human fibroblasts leads to a senescent-like state rather than to an immortal
phenotype (61). Thus, increases as well as decreases in individual components
of pathways may contribute to senescence-associated changes in signal trans-duction. Taken together, senescence-associated changes in mitogenic signaling
pathways occur for a variety of reasons that may include any imbalances in or
dysregulation of controlling pathways. Interestingly, these effects are largely
confined to proliferation and noncritical functions because, if maintained, sub-populations of cells can survive indefinitely in a senescent state.
1.2. Relevance to Aging
Before beginning our discussion of methods for the propagation of human
fibroblasts and determination of replicative life-span, we digress briefly to dis-cuss interpretation of this type of data. We shall also consider the relationship
Fibroblast Model for Cell Senescence Studies 27
between changes observed during senescence in vitro and aging in vivo. Finally,
we will examine a second hypothesis that suggests that senescence in vitro
recapitulates at least some aspects of developmental changes associated with
The finite replicative life-span for normal cells in culture is thought to result
from multiple environmental and genetic mechanisms (5) and has frequently
been used as a model of human aging. Historically the use of replicative life-span of cell cultures as a model for aging has been accepted because (1) fibro-blast replicative life-span in vitro has been reported to correlate directly with
species maximum life-span potential (62), and most importantly (2) cultures of
normal human cells have been reported to exhibit a negative correlation
between proliferative life-span and the age of the donor from whom the culture
was established (8,16,63–68). Other types of evidence also appear to support
the strength of the model. For example, the colony-forming capacity of indi-vidual cells has also been reported to decline as a function of donor age (69,70).
Various disease states of cell donors have been found to significantly influence
the proliferative life-spans of cells in culture. For example, cell strains estab-lished from diabetic (68,71) and Werner’s patients exhibit diminished prolif-erative potential  (19,28,65,72,73). Cultures derived from individuals with
Hutchinson–Gilford syndrome  (28) and Down’s syndrome  (28,74) may also
exhibit decreased proliferative potential, albeit results with these cell lines are
more variable (29). Collectively, these observations have been interpreted to
suggest that the proliferative life-span of cells in culture reflects the physi-ological age as well as any pathological state of the donor from which the cells
were originally obtained.
It must be noted that interpretation of replicative life-span data is often dif-ficult owing to large individual variations and relatively low correlations. For
example, one large study  (75) determined replicative life-span in more than
100 cell lines, yet obtained a correlation coefficient of only–0.33. Hence, it is
difficult to assess whether the reported negative correlations between donor
age and replicative life-span indicate any compromise of physiology or prolif-erative homeostasis in vivo  (75,76). A major factor that has influenced the
results of most studies is the health status of donors when tissue biopsies were
taken to establish the cell cultures (68,75). Most studies include cell lines estab-lished from donors who were not screened thoroughly for disease, as well as
cell lines derived from cadavers to determine the effects of donor age on prolif-erative potential. Variations in the biopsy site have also been a factor that prob-ably influenced the results of many studies (68,75).
Studies of rodent skin fibroblasts appear to support the existence of a small,
but significant, inverse correlation between donor age and replicative life-span
(67,77,78). Furthermore, it has also been observed that treatment of hamster
28 Cristofalo, Volker, and Allen
skin fibroblasts with growth promoters can extend the proliferative life of cul-tures established from young donors but has negligible effects on cultures estab-lished from older donors (79). Aside from inherent species differences and the
effects of inbreeding that may influence these results, it is also apparent that
rodent skin is better protected from some types of environmental injury such as
light exposure. However, even in rodents, the relationship between donor age
and proliferative potential is not entirely clear. For example, an examination of
hamster skin fibroblast cultures established from the same donors at different
ages reveals no age-associated changes in proliferative potential in animals
older than 12 mo (78).
To address these issues, we recently examined the proliferative potential of
124 human fibroblast cell lines from the Baltimore Longitudinal Study of Aging
(BLSA) (80). All of these cell lines were established from donors described as
healthy, at the time the biopsy was taken, using the criteria of the BLSA. This
study revealed no significant change in proliferative potential of cell lines with
donor age, nor did we observe a significant difference between fetal and post-natally derived cultures (80). Goldstein et al. (68) also reported that no rela-tionship between proliferative life-span and donor age could be found in healthy
donors but did observe a relationship in diabetic donors. In addition, we per-formed a longitudinal study by determining the replicative life-span of cell
lines established from individuals sampled sequentially at different ages. As in
the case of the cross-sectional analysis, no relationship between donor age and
replicative potential was found in this longitudinal study. Indeed, cell lines
established from individuals at older ages frequently exhibited a slightly greater
proliferative potential than the cell lines established from the same individuals
at younger ages (80).
1.2.1. Relationship of In Vitro and In Vivo Models
One of the underlying assumptions of in vitro aging models is that the
changes observed during proliferative senescence bear at least some homology
to those observed during aging in vivo. In fact, both similar (concordant) and
dissimilar (discordant) changes have been observed between aging-associated
changes observed in vivo and in vitro; these are summarized in Ta ble 1.
Although the results presented in  Ta ble 1 clearly demonstrate that some
similarities do exist between aging in vivo and replicative senescence, it
remains unclear whether these changes arise through a common mechanism or
via distinct pathways. As seen in  Ta ble 1, senescence in tissue culture and
aging in the intact organism are not homologous. Others have noted that pro-gressive morphological changes begin to develop in diploid cell cultures shortly
after they are established regardless of the donor age; no cells are found in vivo
Fibroblast Model for Cell Senescence Studies 29
at any age that exhibit the morphological phenotype of cells, in vitro, at the end
of their replicative life-span (106).
Rubin (76) suggests that the limited replicative life-span in vitro may be an
artifact that reflects the failure of diploid cells to adapt to the trauma of dissocia-tion and the radically foreign environment of cell culture. However, that hypoth-esis ignores factors such as telomere shortening that appear to influence
proliferative life and that are not dependent on the culture environment. Pres-ently, it is possible to state that the loss of proliferative potential in vitro does
not directly reflect changes in replicative capacity that occur in vivo during aging
and that changes in gene expression associated with replicative senescence are
not completely homologous with changes observed during aging in vivo.
1.2.2. Relationship Between Senescence and Development
One view of the limited proliferative capacity of cells in culture is that it
stems from the effects of the culture environment on the state of differentiation
of the cells  (107–113). Although the state of differentiation may change in
Table 1
Aging in Cell Culture Replicative Senescence vs Donor Age
Concordant features Discordant features
Collagenase (↑)a (81,82) c-fos induction (↓) (20,83,84)
Stromelysin (↑) (85) EPC-1 mRNA (↓) (86,87)
PAI-1 (↑) (88,89) H-twist mRNA (↓)(90;
IGF-BP3 (↓) (91) G-6-PDH mRNA (=)b (54,92)
TIMP-1 (↓) (85) Fibronectin (↑) (93)
HSP 70 (↓) (94,95) ND-4 mRNA (↑) (96,97)
Response to Ca2+ (↓) (98,99) p21 mRNA (↑) (100,101)
MnSOD mRNA (↑?) (102,103)
β-Galactosidase (↑) (Cristofalo,
Chemiluminescence (↑) (54,96)
H2O2 Generation (↑) (54,96)
Collagen a(1)I mRNA (↓) (100,104,105)
Proliferative capacity (↓) (80)
Saturation density (↓) (80)
aArrow indicates direction of change in replicative senescence.
bIndicates no change.
G-6-PDH, glucose-6-phosphate dehydrogenase; HSP 70, heat shock protein 70;  IGF-BP3,
insulin-like growth factor binding protein-3; PAI-1, plasmogen activator inhibitor-1; SOD,
superoxide dismutase; TIMP-1, tissue inhibitor of metalloproteinase-1.
30 Cristofalo, Volker, and Allen
cells that senesce in vitro, there is, in fact, no evidence that the changes in gene
expression observed in fetal cells as they senesce in vitro, are tantamount to
differentiation, in vivo. While some analogous changes can be found they are
greatly outnumbered by the discordant differences that characterize these two
distinct phenomena. Hence, a comparison of senescence-associated changes
and differences that exist between fetal and postnatal cells reveals little simi-larity (Ta ble 2).
At least some analogous similarities exist between senescence in fetal fibro-blasts and developmental changes that occur in vivo. For example, it has been
observed that addition of PDGF-BB stimulated an increased mRNA abundance
of the transcript encoding the PDGF-A chain in fetal and newborns; however,
the response was greatly decreased in adult cells. Senescence in vitro of new-born fibroblasts appears to result in the acquisition of the adult phenotype (116).
In contrast, there are a number of differences reported between fetal- and adult-derived cell lines related to growth factor requirements for proliferation and
migration (117,119–121) that remain disparate even as these cultures become
Table 2
Comparison of Replicative Senescence of Fetal Cells In Vitro with
Differences Between Fetal and Adult Cells?
Concordant Discordant
ND-4 mRNA (↑) (96,97) c-fos induction (=)b (84)
MnSOD activity (↑) (103,114) EPC-1 mRNA (↑) (86,87,97)
Catalase activity (↑) (92) Cu/Zn SOD mRNA (↑) (102)c
IL-1α (↑) (103,115) MnSOD mRNA (↑) (102,114)c
IL-1β (↑) (103,115) Cu/Zn SOD activity (↑) (102)c
Response (↑) (116) COX-1 mRNA (↑) (96)c
SDd mRNA (↑) (96)c
COX activity (↑) (96)c
NDd activity (↑) (96)c
SD activity (↑) (96)c
G-6-PDH mRNA (↑) (92)
PDGF requirement (↑) (117)
Collagen a(1)I mRNA (Ø) (100,118)
β-Actin (Ø) (100;
aArrow indicates direction of difference between proliferatively young fetal and adult cells.
bIndicates no change.
cBased on observations of changes during proliferative senescence, made in this laboratory
that will be presented elsewhere (54).
dND=NADH dehydrogenase; SD=succinate dehydrogenase.
Fibroblast Model for Cell Senescence Studies 31
senescent. For example, Wharton (119) has shown that fetal dermal fibroblasts
will proliferate in plasma or serum while adult dermal fibroblasts require serum.
It is also noteworthy that the expression of some genes, such as  SOD-2,
increases during proliferative senescence but only in some types of fibroblasts
(114); in other types of fibroblasts no change is observed (54,114). It might be
expected that cells placed in culture will be deprived of those signals that direct
the normal sequence of developmental pathways and that differentiation, if it
occurs, is to an aberrant state. Alternatively, fetal cell lines may arise from
different precursor cells than do adult fibroblasts and thus merely differentiate
to a different fibroblast type.
1.2.3. Limitations and Strengths of the System
Although the loss of proliferative potential in vitro may not directly reflect
changes in replicative capacity that occur in vivo during aging, cell cultures
remain a powerful tool for a variety of aging-related studies. These include
studies of heritable damage to cell populations that simulate the effects of aging
in vivo (76), a variety of chemical and molecular manipulations used to induce
a senescence phenotype, the effects of stress (61,76,122–125), and as a system
to study abnormal growth or quiescence (5). The model may also help to fur-ther elucidate the effects of diseases that alter proliferative life-span
(19,28,65,68,71–73,126). Loss of capacity for senescence is a necessary step
for immortalization and transformation to a malignant phenotype. The model
may also prove useful in studies of the relationship between differentiation and
replicative aging (117,119–121).
2. Materials
The serum-supplemented and serum-free, growth factor-supplemented for-mulations presented each give optimal growth of human diploid fibroblast-like
cells. We also present methods for growth of cells in a defined medium using a
defined growth factor cocktail (2,127). All reagents and materials for cell cul-ture must be sterile, and all manipulations should be performed in a laminar
flow hood. Serial propagation is generally performed in serum-supplemented
media, yet serum is a complex fluid with numerous known and unknown
bioactive components. For many studies, it is often desirable if not crucial to
use a serum-free growth medium of defined composition.
2.1. Serum-Supplemented Medium
Suppliers and more detailed information on the items required for the prepa-ration of serum-supplemented media are listed in Table 3.
1. Incomplete Eagle’s modified minimum essential medium: Cells are grown in
Eagle’s modified minimum essential medium (MEM). Although the medium can
32 Cristofalo, Volker, and Allen
be purchased in liquid form, it is considerably less expensive to prepare the
medium from a commercially available mix. In our laboratory incomplete MEM
is prepared by dissolving Auto-Pow™ powder (9.4 g) and 100× basal medium
Eagle vitamins (10 mL) in 854 mL of deionized, distilled water. After the incom-plete medium has been mixed and dissolved, it should be divided into two equal
portions (432 mL each) and placed in 1-L bottles (see Note 2). The caps are
screwed on loosely, autoclave tape is applied, and the bottles are autoclaved for
15 min at 121°C (see Note 3). As soon as the sterilization cycle is finished, the
pressure is quickly released and the bottles are quickly removed from the auto-clave. The bottles are allowed to cool to room temperature in a laminar flow
hood. When the bottles have cooled, the caps are tightened. Incomplete medium
is stored at 4°C in the dark.
2. 100× Basal medium Eagle vitamins: Filter-sterilized 100× basal medium Eagle
vitamins are purchased in 100-mL bottles and stored at –20°C. When first thawed,
using sterile procedures, the vitamin solution is divided into 10-mL portions and
stored in sterile 15-mL centrifuge tubes at –20°C until use.
3. L-Glutamine (200 mM): L-Glutamine (14.6 g) is dissolved in 500 mL of deion-ized, distilled water without heating. This solution is then sterilized in a laminar
flow hood using a 0.2 µm pore size bottletop filter. Aliquots (50 mL) are added to
sterile 100-mL bottles that are then capped and stored at –20°C until use. When
thawed for use, the glutamine solution is divided into 5-mL portions and stored at
–20°C in sterile 15-mL centrifuge tubes until use.
4. Sodium bicarbonate (7.5% w/v): Sodium bicarbonate (37.5 g) is dissolved in
500 mL of deionized, distilled water. This solution is then filter sterilized using
a 0.2-µm pore size bottletop filter. The sterile solution is stored at 4°C.
5. Fetal bovine serum (FBS): Prior to purchase, various lots of fetal bovine serum
(FBS) are tested for 3 consecutive weeks to determine their effects on the rate of
cell proliferation and saturation density. The serum lot that gives the best growth
response is chosen, and quantities that will last about 1 yr are purchased. The
serum is stored at –20°C until use. Once thawed, serum is stored at 4°C for sub-sequent use; it should not be refrozen.
Table 3
Components of Standard Growth Medium
Component Amount/L Supplier Cat. no.
Auto-Pow™, autoclavable powder Eagle
MEM with Earle’s salts without glutamine
and without sodium bicarbonate 1 pkg ICN 11-100-22
100× Basal medium Eagle vitamins 10 mL ICN 16-004-49
200 mM L-Glutamine 10 mL Sigma G3126
Sodium bicarbonate (7.5% solution) 26 mL Sigma S5761
FBSa 100 mL Various
aFBS is from a variety of suppliers and tested on a lot-by-lot basis. See Note 1.
Fibroblast Model for Cell Senescence Studies 33
6. Standard serum-supplemented growth medium (complete medium with 10% v/v
FBS): To prepare the standard serum-supplemented growth medium (complete
medium with 10% v/v FBS), add 13 mL of filter-sterilized 7.5% (w/v) sodium
bicarbonate to 432 mL of sterile, incomplete Eagle’s MEM. The sodium bicarbon-ate must be added first because low pH can affect glutamine and serum compo-nents. After addition of the sodium bicarbonate add 50 mL of sterile FBS. Just
before use the medium is prewarmed to 37°C in a warm water bath, then trans-ferred to a laminar flow hood where 5 mL of a 200 mM solution of filter-sterilized
L-glutamine is added. Complete medium is generally prepared fresh for each use.
If this medium must be stored for periods exceeding 1 wk, additional filter-steril-ized L-glutamine (1 mL/100 mL of complete medium) is added just before use.
2.2. Serum-Free Medium
Suppliers and more detailed information on the items required for the prepa-ration of serum-free media are listed in Ta ble 4.
1. Serum-free growth medium: This medium is prepared by dissolving a packet of
powdered MCDB-104 medium (with  L-glutamine, without CaCl2, without
Table 4
Components of Serum-Free Growth Medium
Component Amount Supplier Cat. no.
MCDB-104, a modified basal medium
with L-glutamine, without CaCl2,
without Na2HPO4, without NaHCO3,
and without HEPES, and with sodium
pantothenate substituted for calcium
pantothenate 1 pkg/L Gibco-BRL 82-5006EA
Sodium phosphate, dibasic 0.426 g/L Sigma S5136
Sodium chloride 1.754 g/L Sigma S5886
Calcium chloride dihydrate 1 mM Sigma C7902
Sodium bicarbonate 1.176 g/L Sigma S5761
HEPESa 11.9 g/L Sigma H9136
1 M Sodium hydroxidea 25 mL/L Sigma S2770
EGF), human recombinant 25 ng/mL Gibco-BRL 13247-010
IGF-I, human recombinant 100 ng/mL Gibco-BRL 13245-014
Insulin 5 µg/mL Sigma I6634
Ferrous sulfate heptahydrate 5 µM Sigma F8633
1 M Hydrochloric acid Trace Sigma H9892
Dexamethasone 55 ng/mL Sigma D4902
95% Ethanol (not denatured) Trace Pharmco 111000-  190CSGL
aNot used in growth medium. See Note 1.
34 Cristofalo, Volker, and Allen
Na2HPO4, without NaHCO3, and without  N-[2-hydroxyethyl]piperazine-N’-[2-ethanesulfonic acid] (HEPES), with sodium pantothenate substituted for calcium
pantothenate) in 700 mL of deionized, distilled water. The packet is also rinsed
several times to dissolve any medium powder that may have adhered to it. The
following additional components are then added in the order listed: 0.426 g of
Na2HPO4, 1.754 g of NaCl, 1.0 mL of a 1  M CaCl2 solution, and 1.176 g of
NaHCO3. For most studies HEPES is not used. The final volume is brought to 1 L
with deionized, distilled water. Incomplete medium is sterilized by filtration
through a 0.2-µm bottletop filter into sterile glass bottles. Using sterile proce-dures in a laminar flow hood, a 5% CO2/95% air mixture is passed through a
sterile, cotton-filled CaCl2 drying tube, through a sterile pipet, and bubbled into
the medium (see Note 4). As the medium becomes saturated with the gas mix-ture, its color changes from pink to a salmon color. The final pH is 7.3–7.5.
Incomplete medium is generally prepared fresh for each use, but it may be stored
for up to 3 wk at 4°C. If unused complete medium is stored longer than 1 wk,
additional L-glutamine (1 mL/100 mL of complete medium) should be added
before use.
2. HEPES-buffered incomplete medium for stock solutions: The pH of carbon diox-ide/bicarbonate-buffered MCDB-104 solutions rises during thawing, resulting in
Ca2PO4 precipitate formation. Thus, growth factor and soybean trypsin inhibitor
solutions that are stored frozen are prepared in HEPES-buffered solutions. To
prepare 1 L of HEPES-buffered incomplete medium, mix medium as described
previously except 11.9 g of HEPES free acid and 25.0 mL of 1  M NaOH are
added instead of sodium bicarbonate. The pH of the medium is adjusted to 7.5 by
titration with additional 1 M NaOH and the volume is brought to a final volume
of 1 L with deionized, distilled water. The medium is sterilized by filtration
through a 0.2-µm bottletop filter into sterile glass bottles. The HEPES-buffered
incomplete medium may be stored at –20°C until needed.
3. Concentrated growth factor stock solutions: For these procedures, use sterile plas-tic pipets and perform all manipulations in a laminar flow hood. Stock solutions
of growth factors (100×) are prepared in HEPES-buffered incomplete medium at
the following concentrations: EGF (2.5 µg/mL) and either IGF-I (10 µg/mL) or
insulin (500 µg/mL) (see Note 5). All stock solutions are dispensed with sterile
plastic pipets into sterile 1.0-mL cryogenic vials. The stock solutions may be
stored at –20°C for short periods (up to 4 wk) or at –70°C for longer periods
(3–4 mo). Dexamethasone (5 mg/mL) is prepared in 95% nondenatured ethanol.
This solution is then diluted into HEPES-buffered incomplete medium to give a
100× stock solution (5.5 µg/mL). Stock dexamethasone is stored in sterile, sili-conized test tubes. Ferrous sulfate is prepared fresh, just prior use. After prepara-tion 5 µL of 1 M hydrochloric acid is added to each 10 mL of the ferrous sulfate
100× stock (0.5 mM). This solution is sterilized by filtration through a 0.2-µm
4. Complete serum-free growth medium: For 100 mL of complete serum-free
growth medium, 1 mL of each of the 100× stock solutions are added to 96 mL of
Fibroblast Model for Cell Senescence Studies 35
incomplete medium (MCDB-104). The resultant concentrations in the serum-free medium are: 25 ng/mL of EGF, 100 ng/mL of IGF-I, or 5 µg/mL of insulin
(see Note 5);  55 ng/mL of dexamethasone; and 5 µM of ferrous sulfate.
5. Soybean trypsin inhibitor solution for serum-free propagation: Soybean trypsin
inhibitor (100 mg) is added to 100 mL of HEPES-buffered incomplete medium.
This solution is sterilized by filtration through a 0.2-µm bottletop filter into a
sterile bottle. The sterile solution is then dispensed into sterile 15-mL centrifuge
tubes in 7-mL portions and stored at –20°C. When needed, the solution is thawed
and diluted 1:1 with bicarbonate-buffered incomplete medium.
2.3. Trypsinization
Suppliers and more detailed information on the items required for the prepa-ration of trypsinization solution are listed in Ta ble 5.
1. Ca2+/Mg2+-free medium: Cells tend to aggregate in media containing calcium; it
is thus desirable to use a medium that is low in Ca2+ and Mg2+ for mixing trypsin
solution. To prepare Ca2+/Mg2+-free medium, the following ingredients are added
to 900 mL of deionized, distilled water with magnetic stirring: 6.8 g of NaCl,
0.4 g of KCl, 0.14 g of NaH2PO4 · H2O, 1 g of glucose, 20 mL of 50× MEM
amino acids without glutamine, 10 mL of 100× basal medium Eagle vitamins,
and 10 mL of a 0.5% (w/v) solution of phenol red. The solution is then diluted to
1 L with deionized, distilled water and sterilized by filtration. The Ca2+/Mg2+-free medium is stored at 4°C until use.
2. Trypsin stock solution (2.5%): Filter-sterilized trypsin (2.5%) in Hanks’ buffered
salts solution is purchased in 100-mL bottles and stored at –20°C. Repeated
Table 5
Components of Trypsinization Solution
Component Final amount Supplier Cat. no.
Sodium chloride 6.8 g/L Sigma S5886
Potassium chloride 0.4g/L Sigma P5405
Sodium phosphate monohydrate,
monobasic 0.14g/L Sigma S5655
Glucose 1 g/L Sigma G6152
50× MEM amino acids, without
glutamine 20 mL/L Gibco-BRL 11130-051
100× basal medium Eagle vitamins 10 mL/L ICN 16-004-49
0.5% Phenol red 10 mL/L ICN 16-900-49
Sodium bicarbonate (7.5% solution) 5 mL/50 mL Sigma S5761
Trypsin, 2.5% in Hanks’ balanced
salt solution 5 mL/50 mL Sigma T4674
Soybean trypsin inhibitor, type I-S 1 mg/mL Sigma T6522
36 Cristofalo, Volker, and Allen
freeze–thaw will very rapidly decrease activity. The bulk trypsin solution should
be thawed only once, dispensed in 5-mL portions in sterile 15-mL centrifuge
tubes and then stored at –20°C until use.
3. Trypsin solution (0.25%): Five milliliters of sterile sodium bicarbonate (7.5%) is
added to 40 mL of ice-cold Ca2+/Mg2+-free medium. Subsequently, 5 mL of
freshly thawed 2.5% trypsin stock is added to the solution. This solution should
be prepared just before the cells are treated and should be kept on ice.
2.4. Thymidine  Incorporation
Suppliers and more detailed information on the items required for measure-ment of thymidine incorporation are listed in Table 6.
1. [3H-methyl]-thymidine stock solution: Under sterile conditions, [3H-methyl]-thy-midine (2 Ci/mmol, 1 mCi/mL) is diluted to a concentration of 5 µCi/mL in ster-Table 6
Items for Thymidine Incorporation
Item Supplier Cat. no.
[3H-methyl]-Thymidine, 2 Ci/mmol;
1 mCi/mL Dupont NEN NET-027A
Coverslip, No. 1, 22 mm × 22 mm Thomas 6662-F55
Coverslip rack, ceramic Thomas 8542-E30
Coverslip rack, glass Fisher 08-812
Chloroform Sigma C5312
95% Ethanol, not denatured Pharmco 111000190CSGL
95% Sulfuric acid Sigma S1526
70% Nitric acid Sigma 25,811-3
Sodium hydroxide Sigma S5881
Petri dish, glass, 100 mm Thomas 3483-K33
NTB-2 Emulsion Eastman Kodak 165 4433
D-19 Developer Eastman Kodak 146 4593
Acid fixer Eastman Kodak 197 1746
Hematoxylin, Harris Modified Fisher SH30-500D
Permount Fisher SP15-100
Microscope slide, 3 in × 1 in Thomas 6684-H61
Lab-Tek® Chamberslide™, two-chamber Nalge Nunc 177380
Lab-Tek® Chamberslide™, four-chamber Nalge Nunc 177437
Lab-Tek® Chamberslide™, eight-chamber Nalge Nunc 177445
Sodium phosphate, dibasic Sigma S5136
Potassium phosphate, monobasic Sigma P5655
Methanol Fisher A408-1
Slide mailer, polypropylene Thomas 6707-M27
Slide box, polypropylene Thomas 6708-G08
Fibroblast Model for Cell Senescence Studies 37
ile medium. This stock solution is aliquoted (5-mL portions) in a laminar flow
hood using sterile procedures into sterile, 15-mL centrifuge tubes and stored at
–20°C until use.
2. Phosphate-buffered saline (PBS) solution: dissolve 8 g of NaCl, 0.2 g of KCl,
1.44 g of Na2HPO4, and 0.24 g of KH2PO4 in 900 mL of H2O with magnetic
stirring. The pH is adjusted to 7.4 with HCl, the volume adjusted to 1 L, and the
solution is autoclaved for 20 min at 121°C.
3. Emulsion: Kodak NTB-2 emulsion is purchased in a lightproof container. The
emulsion is stored at 4°C (see Note 6).
4. Developer and Fixer
a. Kodak D-19 developer is purchased in packets that make 1 gal when reconsti-tuted. The entire packet is used at one time and the solution is stored in a
brown bottle in the dark. The developer remains useable for 1–3 mo. When
the developer turns yellow, it is discarded.
b. Acid fixer is made and stored in the same manner as the D-19 developer.
3. Methods
3.1. Cell Propagation in Serum-Supplemented Medium
Cells may be grown in a variety of culture vessels (see Note 7). Amounts
described in the following procedure are for a T-75 flask. Proportional amounts
are used for other size vessels; i.e., for a T-25 flask, one third of all of the
amounts given is used. Trypsinization and seeding of flasks should be per-formed in a sterile environment (see Note 8).
To propagate adherent cells:
1. Prepare fresh trypsin solution (0.25%) and place it on ice; prepare fresh growth
medium and warm it to 37°C.
2. Using sterile procedures in a laminar flow hood, remove spent growth medium
from the culture vessel. For flasks and bottles, the medium should be removed by
aspiration or decanting from the side opposite the cell growth surface. For cell
culture plates and dishes, the medium should be removed by aspiration from the
edge of the growth surface.
3. Gently wash the monolayers of adherent cells twice with 0.25% trypsin solution
(4 mL).
4. Remove residual trypsin solution by aspiration from the side opposite the cell
growth surface (flasks) or from the edge of the growth surface (plates, dishes, and
slides) as appropriate.
5. Add enough trypsin solution (0.25%) to wet the entire cell sheet (2 mL/T-75).
6. The culture vessel should be tightly capped to maintain sterility and placed at
7. The cells will assume a rounded morphology as they are released from the growth
surface. Detachment of the cells should be monitored using a microscope. As a
general rule, detachment will be complete within 15 min. The trypsinization pro-cess may be speeded up by gently tapping the sides of the flask. Care should be
38 Cristofalo, Volker, and Allen
taken to not splash cell suspension against the top and sides of the flask, because
this will lead to errors in the determination of the number of cells in the flask.
8. When all of the cells have detached from the growth surface, as determined
by inspection with a microscope, the flask is returned to the laminar flow
hood. Complete medium with 10% v/v FBS is carefully pipeted down the
growth surface of the vessel to neutralize the trypsin and to aid in pooling the
cells. For a T-75 flask, 8 mL of complete medium is used. The final harvest
volume is 10 mL.
9. Cell clumps should be dispersed by drawing the entire suspension into a 10-mL
pipet and then allowing it to flow out gently against the wall of the vessel. The
process is repeated at least three times. The procedure is then repeated with a
5-mL pipet. Until the procedure becomes routine, a sample is withdrawn and
examined under the microscope to ensure that a suspension of single cells has
been achieved. During this process, the cells should be kept on ice to inhibit cell
aggregation and reattachment.
10. Using sterile procedures, remove an aliquot from the cell suspension, then dilute
it into Isoton II in a counting vial. Typically, 0.5 mL of the cell suspension is
diluted into 19.5 mL of Isoton II.
11. Count the sample with a Coulter Counter.
12. Calculate the number of cells in the harvest. Calculate the volumes of cell sus-pension and complete medium needed for new cell culture growth vessels. In
most cases, cells are seeded at a density of 1 × 104 cells/cm2 of cell growth sur-face, and the total volume of cell suspension plus complete medium added to the
culture vessels is maintained at 0.53 mL/cm2 of cell growth surface.
13. In the laminar flow hood, add the calculated amounts of complete medium to new
culture vessels.
14. Dissolved CO2 in equilibrium with HCO3
– is the principal buffer system of the
medium, although serum also has some buffering capacity. Because CO2 is vola-tile, the gas phases in the flasks are adjusted to the proper pCO2 to maintain the
pH of the medium at 7.4. Using sterile procedures in a laminar flow hood, a 5%
CO2/95% air mixture is passed through a sterile, cotton-filled CaCl2 drying tube,
through a sterile pipet, and into the gas phase of the cell culture flask with the
growth surface down. As the gas mixture is flushed over the medium surface, the
color of the medium will change from a dark red toward a red-orange. The flask
is flushed until the medium no longer changes color. At this point, the gas above
the medium is 5% CO2 and the pH of the medium is 7.4 (see Note 4). The flask is
then tightly capped to prevent gas exchange with the outside environment. Cells
grown in culture plates, dishes, and Lab-Tek® slides, which are not gas-tight, are
not equilibrated with the gas mixture in this manner; instead they must be grown
in incubators that provide a humidified, 5% CO2 atmosphere.
15. The cell harvest is resuspended with 10-mL and 5-mL pipets, as before. Inoculate
each culture vessel to a final density of 1 × 104 cells/cm2 of growth surface.
16. Briefly flush the culture vessel a second time with the 5% CO2/95% air mixture
to replace the CO2 lost when the vessel was opened. Cap the flask tightly and
Fibroblast Model for Cell Senescence Studies 39
incubate at 37°C. Periodically, examine the color of the medium to ensure that
the seal is gas tight.
17. The cumulative population doubling level (cPDL) at each subcultivation is calcu-lated directly from the cell count (see Note 7).
One week after seeding a T-75 flask with the standard inoculum of 7.5 × 105
cells at a cPDL of 37.2, the cells are harvested. One doubling would yield
2 × 7.5 × 105 = 1.5 × 106 cells; two doublings would result in 4 × 7.5 × 105 =
3.0 × 106 cells; three doublings would yield 8 × 7.5 × 105 = 6.0 × 106 cells, etc.
Thus, the population doubling increase is calculated by the formula:
NH/NI = 2X
or [log10 (NH) – log10 (NI)]/Log10 (2) = X
where NI = inoculum number, NH = cell harvest number, and X = population
doublings. The population doubling increase that is calculated is then added to
the previous population doubling level to yield the cPDL. For example, if
9.1 × 106 cells were harvested, then the population doubling increase can be
calculated from the expression:
9.1 × 106 cells = 2 (X) × 7.5 × 105 cells
X log10 2 = log10 (9.1 × 106) – log10 (7.5 × 105)
X = 3.6
The population doubling increase is added to the previous cPDL to give the
new cPDL of the cell population. For this example, the new cPDL is 37.2 + 3.6
= 40.8. The end of the replicative life-span was defined by failure of the popu-lation to double after 4 wk in culture with 3 wk of consecutive refeeding.
3.2. Cell Propagation in Serum-Free Medium
1. Because undefined mitogens and inhibitors present in serum complicate the inter-pretation of cell growth response results, soybean trypsin inhibition solution
should be used to stop trypsin instead of complete medium with 10% v/v FBS to
wash and collect the cells from the growth surfaces of flasks. Otherwise, cells
are released from the surface of their culture vessel exactly as described previ-ously for propagation of cells in serum-supplemented medium (Subheading 3.1.,
steps 1–12).
2. Wash the cells to remove residual mitogens and trypsin inhibitor, rather than using
them directly to inoculate the culture flasks:
a. Under sterile conditions, the cells are pelleted by centrifugation at 75g for
5 min at 4°C.
40 Cristofalo, Volker, and Allen
b. The centrifuge tubes are placed in ice, transferred to a laminar flow hood, the
supernatant is removed, and the cells are resuspended in 10 mL of incomplete
serum-free growth medium (Subheading 2.1.).
c. Under sterile conditions, the cells are again pelleted by centrifugation, and
after removal of the supernatant, the cells are resuspended in 10 mL of com-plete serum-free growth medium (Subheading 2.2.).
3. Determine the cell number with the Coulter Counter as before, using an aliquot
of the cell suspension (0.5 mL)
4. Cells are then seeded exactly as described in  Subheading 3.1., steps 13–17,
except that serum-free cell growth medium is used.
3.3. Replicative Life-Span
As noted previously, cells in culture exhibit a finite number of replications.
At the end of their in vitro life-span substantial cell death occurs; however, a
stable population emerges that can exist in a viable, though nondividing, state
indefinitely (128). Furthermore, small subpopulations of cells may retain some
growth capacity even after the vast majority of cells in a culture are no longer
able to divide. As a practical matter, cultures of cells may be considered to have
reached the end of their proliferative life-span when the cell number fails to
double after 4 wk of maintenance in growth medium with weekly refeedings.
The maximum proliferative capacity of the cells is determined as follows:
When cell cultures are near the end of their proliferative life-span, at least
four identical sister flasks are prepared. One flask is harvested each week. If
the number of cells harvested is at least double the number inoculated, cells are
subcultivated as usual. One of the sister flasks may also need to be harvested to
provide enough cells for subcultivation into four flasks. If the number of cells
harvested is not at least double the number inoculated, all of the sister flasks
are refed by replacement of the spent medium with fresh complete medium and
equilibration with 5% CO2/95% air mixture. This process is repeated three
times. When cultures fail to double during this period, the culture may be con-sidered to have reached the end of proliferative life or to be “phased out.”
3.4. Saturation Density
Cultures are grown until the cells are densely packed and no mitotic figures
are apparent. This usually requires from 7 to 10 d after seeding for early pas-sage cells, and more than 9 d for later passage cells. To estimate the saturation
density, these confluent and quiescent cells are then harvested and counted as
described previously.
3.5. Microscopic Estimate of Cell Density
It is often desirable to obtain an estimate of cell density without harvesting
the cells. A stage micrometer is used to calibrate the eyepiece micrometer and
Fibroblast Model for Cell Senescence Studies 41
determine the diameter of the field of view for each objective and ocular lens
used. The area of the field of view is calculated as Area = π r2, where r is the
radius of the field of view.
Scan the sample to ensure that the cells are uniformly distributed. Then count
at least 400 cells using random fields. Since the standard deviation of a Poisson
distribution is the square root of the number, 400 cells are counted. The square
root of 400 (20) is 5%, which is the limit of statistical reliability for most bio-logical work. Record the number of cells and the number of fields counted. The
cell density is then calculated as follows:
cell density = (no. of cells counted)/([no. of fields counted] · [area per field])
3.6. Thymidine Incorporation
3.6.1. Coverslips
1. Place coverslips in a clean, glass rack using forceps.
2. Lower the rack containing the coverslips into a solution of chloroform/95% etha-nol (1:1) and allow to soak for 30 min.
3. Rinse the coverslips with deionized water.
4. Submerge the coverslips in a 95:5 solution of concentrated sulfuric acid (95%)/
concentrated nitric acid (70%), previously prepared in a fume hood and allowed
to cool to room temperature. Soak the coverslips in this solution for 30 min.
5. Rinse the coverslips thoroughly in deionized water.
6. The rack containing the coverslips should then be lowered into a solution 0.2 M
NaOH and allowed to soak for 30 min.
7. Remove the coverslips from the NaOH solution and rinsed at least three times in
deionized water.
8. Remove the coverslips from the rack and allow to air-dry on lint-free disposable
9. When completely dry, bake the coverslips for 3 h at 180°C for sterilization.
3.6.2. Cell Slides
1. In a laminar flow hood, under sterile conditions, cells are harvested and counted
in the usual manner.
2. Cells are seeded at a density of 1 × 104 cells/cm2 on Lab-Tek® slides or in cell
culture dishes that contain coverslips (Subheading 3.6.1.). If using coverslips
use sterile forceps to arrange them in the dish so that they do not overlap one
3. Immediately after seeding, the slides and dishes are placed in an incubator at
37°C in an atmosphere of 5% CO2/95% air.
4. Twenty-four hours later, add the stock solution of [3H-methyl]-thymidine (spe-cific activity 2 Ci/mmol; Subheading 2.4.) to the cultures to a final concentration
of 0.1 µCi/mL.
5. After 30 h  (129), the labeling medium is removed, and cells are immediately
washed twice with PBS (Subheading 2.4.), fixed in 100% methanol for 15 min,
42 Cristofalo, Volker, and Allen
and air-dried. If cells are grown on coverslips, remove the coverslips from the
dishes and place in a clean ceramic or glass rack using forceps prior to washing
and fixing. If a Lab-Tek® slide is used, the plastic container and gasket must
be removed prior to washing and fixing. These procedures should be done rapidly
to limit damage to the cells. The cells must not be permitted to dry before they
are fixed.
6. Mount coverslips with the cell surface up using mounting resin. Allow the resin
to dry overnight.
3.6.3. Autoradiography
1. Remove the Kodak NTB-2 emulsion from storage at 4°C and place it in a warm
room at 37°C. The emulsion will liquefy in 3–4 h. The emulsion may also be
melted by placing it in a 40°C water bath in the dark for about 1–1.5 h. Do not
shake the bottle because the resultant bubbles may cause irregularities in the final
emulsion thickness.
2. In a dark room, the desired amount of emulsion is gently, but thoroughly
mixed in a 1:1 ratio with deionized, distilled water.
3. Add 15–20 mL of the 1:1 emulsion/water solution to a container (a slide
mailer works well for this) previously set up in a 40°C water bath in the dark.
4. Dip each slide individually into the slide mailer. One dip is sufficient to coat
the slide.
5. Place each dipped slide in a standing (vertical) position in a wire test tube rack to
drain off excess emulsion. The slides are allowed to dry for 30 min in the dark.
6. The dipped slides are placed into a slide box with a desiccant. The box is covered
and sealed with black electrical tape. The box is placed inside a second light-tight
container that also contains a desiccant and this is also sealed with electrical tape.
7. The container is placed at 4°C for 4 d.
Development of Cell Slides
1. Pour Kodak D-19 developer and acid fixer into large glass dishes.
2. Open the slide containers in a dark room (photo-safe light can be used), and
remove the slides and place them in racks.
3. Place the slides in developer for 5 min.
4. Transfer the slides fixer for 5 min.
5. At this point, the room light may be turned on, if desired. Gently rinse the slides
for 15 min in cold running water. The slides should next be lightly stained with
Harris” modified hematoxylin stain to enhance nuclear visualization.
3.6.5. Staining Slides
1. Place the developed slides in staining dishes containing Harris” modified hema-toxylin stain for 5–10 min. This amount of time is sufficient to produce light
2. Drain slides in slide racks on paper towels.
Fibroblast Model for Cell Senescence Studies 43
3. Rinse the slides continuously with deionized, distilled water until the excess stain
is removed and then drain them on paper towels.
4. Excess emulsion should be wiped from the back of slides while they are
still damp
5. Air-dry the slides.
3.6.6. Counting Labeled Nuclei
1. For ease in identifying the limits of individual chambers under the microscope, if
Lab-Tek® slides are used, the stain between the individual chambers can be
removed with the end of a paper clip or push pin.
2. Silver grains over nuclei where [3H-methyl]-thymidine has been incorporated into
the DNA will be readily visible at 400× magnification. Nuclei with five or more
grains are considered labeled.
3. To determine the percentage of labeled nuclei, at least 400 cells are counted per
coverslip or chamber using random fields. Typically, determinations are done in
4. Notes
1. It is important that the highest quality deionized, distilled water is used to prepare
growth medium and all other reagents used for cell culture.
2. It is important that the bottles not be filled to more than one half volume to pre-vent overflow during sterilization.
3. Prolonged heat destroys some medium components.
4. Cells in a culture environment require carbon dioxide for growth and survival,
and we have found that well controlled CO2/bicarbonate buffered media gives
superior growth when compared with media containing synthetic buffers, such as
5. Insulin and IGF-I both stimulate growth through the IGF-I receptor, although
insulin has lower affinity for the IGF-I receptor and 50-fold higher concentra-tions are required to achieve comparable growth. Insulin is less expensive
than IGF-I, and despite the reduced specificity, insulin is satisfactory for most
6. Emulsion should never be stored near high-energy sources of radioactivity.
7. Cell cultures are typically subcultivated weekly. Multiple identical sister flasks
are prepared at subcultivation, as a hedge against potential contamination or other
anomalies. Because a substantial fraction (25–60%) of the cells do not survive
subcultivation (130), the number of cells does not generally increase above the
seeded cell number until approx 24 h after subcultivation.
8. Cultures should routinely examined microscopically for contamination, and
tested for mycoplasma at 5-wk intervals (131).
This work was supported by the National Institutes of Health Grants
AG00378 and AG00532.
44 Cristofalo, Volker, and Allen
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Fibroblast Model for Cell Senescence Studies 49
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Human T-Cell Clones 53
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Human T-Cell  Clones
Graham Pawelec
1. Introduction
Techniques for generating human T-cell clones (TCCs) were first described
nearly two decades ago (1,2). This was a direct consequence of the discovery
of T-cell growth factor and the subsequent ability to propagate T-cells over
extended periods  (3). Early on, numerous publications in immunology indi-cated an apparently unlimited growth potential of normal mammalian T lym-phocyte cultures; however, even at this time, other investigators challenged this
conclusion (4,5). Nonetheless, the possibility remained that at least some TCCs
represented an exception to the rule of the Hayflick Limit for growth of normal
somatic cells. If this were the case, the real relevance of replicative senescence
as a universal phenomenon would be highly questionable. On the other hand, if
those T-cells surviving apparently indefinitely were endowed with the proper-ties of stem cells rather than differentiated cells, this quandary would be
resolved. However, as far as could be judged, the apparently immortal TCCs
described in the literature seemed to possess all the attributes of normal T-cells,
not stem cells. Several explanations for this apparent paradox have been pro-posed, the most likely of which may be that such immortal lines are in fact
abnormal. Few clones were tested for karyotypic or other abnormalities. Such
analyses, when performed, often revealed genetic aberrations in human as
well as murine clones (6,7). In the case of murine cells, continuous cultures
often transform spontaneously in culture, but in humans this is rare or absent.
We and others have systematically approached the question of longevity of
normal human TCCs using variations of the original interleukin-2 (IL-2)-dependent cloning and propagation protocol (1,2). This procedure involves lim-iting dilution of the cell suspension to be cloned, and microwell culture of the
54 Pawelec
diluted cells on an irradiated feeder cell layer in the presence of chemically
defined media supplemented with growth factors (e.g., IL-2). We wished to
establish whether culture aging of TCCs did occur, how it could be character-ized, and whether it depended on the source and nature of the T-cells studied.
The first question approached was to what extent culture conditions affected
cloning efficiencies and longevity of the TCC (8); most of our early data were
then obtained using a standard culture system employing medium supple-mented with human serum (HS) and natural IL-2  (9) and using feeder cells
consisting of a pool of peripheral blood mononuclear cells (PBMCs) from >20
random healthy donors. The T-cells to be cloned were derived from young adult
donors and were mostly prestimulated with alloantigens. Under these condi-tions, the type of T-cell predominantly derived was CD4+ and carried the α/β
T-cell receptor (TCR2). This chapter therefore focuses on this type of TCC,
and not on CD8 or TCR1 cells, which seem to behave somewhat differently in
culture, but which we have not studied so extensively.
For meaningful studies of T-cell aging in vitro, it is essential to know the in
vivo age of the starting T-cell population. This is impossible in a mixture of
T-cells from an adult individual, as separation of subsets according to naive
cell and memory cell markers is a crude and inaccurate method. Assessing age
by measuring telomere lengths, even of individual cells, is also not satisfac-tory, as it cannot take into account whether telomerase has been activated at
some point in these cells (10). The only way to be sure that all T-cells being
studied are of the same age at the beginning of the experiment is to isolate
precursors and cause them to differentiate into T-cells in vitro (11,12). Longev-ity comparisons between these pre-T-cell-derived precursors and TCCs from
mature T-cells of the same donors suggested that the latter have a shorter life
expectancy corresponding to the time required for the precursors to develop
into T-cells in vitro (13). We therefore hypothesized that the T-cell “clock” was
first set at the time when fully mature T-cells were generated and not at some
time beforehand at the precursor or stem cell level (14). Going further back in
the T-cell differentiation pathway, CD34+ stem cells have the potential to
develop into T-cells in in vitro culture systems (15), and this property could be
exploited to study T-cell aging. Thus far, cumbersome thymic organ culture or
thymic stromal cell culture systems have been required for this; here we present
a variant TCC culture protocol that allows the generation of mature T-cells from
isolated CD34+ stem cells in liquid culture in the absence of thymic components.
2. Materials
1. T-cell growth factors (TCGFs): The quality and purity of the T-cell growth fac-tors employed is critical. The main and most commonly used TCGF is IL-2 in the
form of purified recombinant protein. Nowadays, many companies offer high-
Human T-Cell Clones 55
quality IL-2; the investigator should pretest a batch for suitability for the
cells being cultured. Mixtures of TCGFs may be useful in some circumstances,
especially IL-2 + IL-4 or IL-2 + IL-7 (see Subheading 3.). Major suppliers
are Genzyme, Endogen, PeproTech, R and D Systems, Boehringer-Mannheim,
and so on.
2. Monoclonal antibodies (MAbs): Again, there are many suppliers of different an-tibodies, and companies favored will be different in different parts of the world.
Major suppliers are Becton-Dickinson, Coulter-Immunotech, DAKO, and so on.
However, for certain purposes, such as cell isolation with magnetic beads (see
Subheading 2.3.1.), it may be economically desirable to obtain hybridoma cells
and produce MAb oneself. The hybridoma cells are easy to grow, and for the
purposes of cell separation, culture supernatants do contain enough MAbs.
Obtaining hybridomas may be a problem, but cell banks such as the American
Type Culture Collection (ATCC) can provide hybridomas secreting MAbs against
common antigens sufficient for most cell separation purposes.
3. Magnetic beads:
a. Dynabeads (Dynal, Oslo, Norway): In this method, the T-cell population is
negatively selected after the cells are labeled with cocktails of MAbs against
B cells (e.g., CD19), natural killer (NK) cells (CD16), monocytes (CD14),
major histocompatibility class (MHC) II, etc. Because of the amount of MAbs
required it is recommended that hybridoma supernatants are used. Dynabeads
M450 coated with sheep anti-mouse IgG can be employed for most negative
cell separations.
b. Miltenyi CD34 Progenitor Kit (Miltenyi Biotec, Bergisch-Gladbach, Ger-many) is supplied with the necessary reagents for CD34 cell isolation by posi-tive selection. The magnetic particles are precoated with CD34 MAbs directed
against a particular exposed epitope of CD34 (class II epitope); purity of the
derived population can then be checked with a MAb directed at a different
epitope (e.g., anti-class I MAb My10 from Coulter-Immunotech).
4. Culture medium: Human serum for supplementing media such as RPMI 1640
or IDMEM to support long-term growth of T-cells cannot be reliably obtained
commercially. It is necessary to prepare and screen the serum on T-cells in the
laboratory. Serum can be obtained or purchased from blood banks, but it is
difficult to obtain enough male nontransfused AB donors for regular use. It
may be satisfactory to use male nontransfused donors of any blood type, as we
do. Serum is separated from coagulated blood by centrifugation and one ali-quot from each of at least 20 sera prepared at the same time is heat inactivated
(30 min at 56°C). The bulk of the sera are frozen separately. The test samples
are then separately tested for their ability to support T-cell proliferation. Those
that are judged satisfactory are then thawed and pooled. Culture medium must
be supplemented with 10–20% of such serum pools to support long-term T-cell
Alternatively, use X-Vivo 10 or X-Vivo 15 serum-free medium (BioWhit-tacker), formula unknown.
56 Pawelec
3. Method
3.1. Source of Cells to Be Cloned
3.1.1. Purification of T Cells (see Note 1)
1. Selectively deplete non-T-cells from PBMC using a cocktail of antibodies for
CD14 (expressed by monocytic cells, macrophages, dendritic cells), CD16 (on
NK cells), CD19 (B cells and B-cell precursors), and HLA-DR (monocytes, B
cells, activated T-cells, and dendritic cells).
2. Incubate 107 cells/mL at 4°C for 30 min with approx 10 µg/mL of each antibody,
centrifuge, wash twice, and resuspend in 1.5 mL of phosphate-buffered saline
(PBS) with 0.1% bovine serum albumin (BSA).
3. Add approx 108 washed Dynabeads in 0.5 mL and incubate at room temperature
for 1 h. Gently shake occasionally. Add 2 mL of PBS and put the tube into the
magnetic field for 1–2 min. Gently aspirate the supernatant. This contains
the negatively selected cells not held by the magnet. Wash twice and control
purity with anti-T-cell antibody by immunofluorescence.
4. Remove any possible remaining functional accessory cells by treating the popu-lation with  L-leucyl-L-leucine methyl esther (LME). Incubate at 2.5 × 106/mL in
10 mM LME for 45 min at room temperature in culture medium without serum.
5. Wash twice and then check absence of functional accessory cells. This can be
done by stimulating with T-cell mitogens such as phytohemagglutinin in the
absence of added cells. There should be no response. After reconstitution of acces-sory function with B-lymphoblastoid cell lines (B-LCLs) the response should be
measurable. A typical protocol is to incubate 2.5 × 104 T-cells per round-bottom
microtiter plate well in culture medium together with 1% phytohemagglutinin
(PHA, M form; Gibco-BRL) in triplicate. A duplicate set of wells receives in
addition 2.5 × 104 B-LCL cells (irradiated at 80 Gy). Proliferation can be assessed
3 d later, for example, by addition of 37 kBq/well of tritiated thymidine and
assessing incorporated nuclear radioactivity after 8–16 h.
3.1.2. Purification of CD34+  Cells
1. Separate low-density mononuclear cells (MNCs, <1.077 g/mL) from buffy coats
from healthy adult donors by isopycnic centrifugation (e.g., 25 min, 400g over
Lymphoprep). Wash twice and centrifuge through a cushion of 10% BSA in PBS
to remove platelets.
2. Separate cells using the MiniMACS Multisort system (Miltenyi Biotec, Bergisch-Gladbach): incubate 5 × 108 MNCs for 5 min at 12°C in 500 µL of FcR-blocking
reagent (Miltenyi Biotec) and then with the same volume of CD34 Multisort
microbeads without washing. Then incubate at 4°C for 45 min, centrifuge, and
resuspend in PBS–EDTA.
3. Rinse MS+ separation columns (Miltenyi Biotec) and place in the magnetic field.
Load 108 cells onto each column in 500  µL. Wash thrice and discard eluate.
Remove column from the magnetic field and elute CD34+ cells with 1.5 mL of
Human T-Cell Clones 57
buffer. Load this eluate onto a second column and isolate the CD34+ cells iso-lated by repeating the above procedure.
4. Incubate the microbead-labeled CD34+ cell population with Multisort Release
Reagent (Miltenyi Biotec) for 10 min at 12°C to release beads from the cells.
5. Then load the suspension onto a third MS column in a magnetic field. Centrifuge
the eluate containing bead-free CD34+ cells through a cushion of 10% BSA in
PBS for 10 min at 600g. Resuspend pellet by adding 30 µL of stop reagent and
10 µL of streptavidin-conjugated microbeads (Miltenyi Biotec) and then incu-bate at 6°C for 30 min.
6. Resuspend cells in 500 µL of buffer and load onto freshly prepared MS columns
in a magnetic field. Collect eluate containing the CD34+ cells and check purity.
For this, incubate the cells for 30 min at 4°C with phycoerythrin (PE)-conjugated
CD34 MAb directed against an epitope other than that used for bead separation.
Greater than 98% of the cell population must be CD34+.
3.2. LD Cloning, Propagation, and Longevity Assessment
1. Resuspend cells for cloning in culture medium (see Note 2) and adjust the con-centrations so that 10 µL contain 45, 4.5, or 0.45 cells. Pipet 10 µL of the 0.45
suspension to 60 × 1 mm diameter wells of culture trays (“Terasaki plates”) and
leave in a vibration-free area for 1 h. Check distribution of cells in the wells
visually using an inverted microscope (being careful to look around the edges of
the wells). Only 37% of the wells should contain cells according to the Poisson
distribution. Readjust dilutions if necessary, replate, and check again.
2. Plate multiple trays (at least five) with the 0.45 cells/10 µL suspension, one with
4.5 and one with 45. Add feeder cells to each well. Irradiated (30 Gy) pooled
PBMCs can most flexibly be used as feeder cells at 1 × 104/well (see Note 3).
3. Stack plates wrapped in aluminum foil for ease of handling and as a precaution
against contamination. Incubate at 37°C in 5% CO2 in air in a humidified incuba-tor for up to a week and then examine the plates using an inverted microscope.
4. Transfer contents of wells containing viable growing cells (> one third full) to
7 mm diameter flat-bottom microtiter plate wells with fresh medium and 1 × 105
feeder cells. Retain Terasaki plates for up to 2–3 wk and examine again at inter-vals to identify any late positive wells. Transfer these to microtiter plates
as well.
5. Examine microtiter plates every few days. Split those becoming overcrowded
with growing cells 1:1 into new culture wells and re-feed with medium (but not
feeder cells). After about a week in microtiter plates, transfer contents of wells
with growing cells into 16 mm diameter cluster plate wells with 2–5 × 105 feeder
cells, and fresh medium (see Note 4).
6. Observe after 3–4 d. Divide wells that are full or nearly full into four, the others
into two, with fresh media, but no more feeder cells. After a total of about a week in
cluster plates, count the number of cells in each clone and subculture to 2 × 105/well,
again with 2–5  × 105 feeders/well and fresh medium. Supplement with fresh
medium after 3–4 d and subculture again if necessary. Continue to propagate by
58 Pawelec
weekly or fortnightly subculture with new feeder cells and fresh medium (see
Note 5).
7. Estimate longevity in terms of population doublings (PDs). Score initial limiting
dilution cloning wells as positive if at least one third full of growing cells. One
third of the surface of a Terasaki well is equivalent to about 1000 cells (= 10 PDs).
Use the number of clones derived at this stage to calculate the average life-span
of all clones derived, that is, do not include clones achieving less than this num-ber of PD in the analysis. The number of cells per microtiter plate well prior to
cluster plate transfer is approx 1 × 105 (= approx 17 PDs). Assume that clones
dying between the Terasaki and microtiter plate stages have undergone 17 PDs,
and use this figure in calculations of average longevity. After clones are trans-ferred to 16 mm diameter cluster wells, the number of PDs undergone can be
estimated for each clone from the exact number of cells counted at each subcul-ture. Cryopreserve cells at any point of their life-spans. Continue calculations of
PDs undergone on the basis of the number of viable cells replated after thawing,
not the number of cells originally present. Take maximum life-span of cells in
each cloning experiment to be the PDs corresponding to the time point of the
death of the longest living clone in each case (see Note 6).
4. Notes
1. For many applications where mature T-cells are to be cloned, it is not necessary
to purify them beforehand. PBMCs as the starting population can be so stimu-lated that only T-cells can grow (e.g., with T-cell mitogens or antigens). When
using CD34+ cells, purity is, however, critical, because contaminating non-CD34
cells most likely have a growth advantage over the CD34+ cells.
2. Clearly the culture medium employed is a critical aspect of the technique.
For many years, we and others found that although T-cells could be grown for
limited periods in completely chemically defined serum-free media, cloning
and long-term propagation in such media was not possible. We and others
were forced to use a serum supplement, most commonly FCS or HS. In our
experience, very few batches of FCS prove suitable for human T-cell cloning
and extensive propagation. Unfortunately, the same was true for commer-cially available HS. We have therefore always obtained material from blood
banks and prepared the serum ourselves. Because of paucity of AB blood
donors, we have always used nontransfused male blood. Each serum is sepa-rately heat inactivated (56°C, 30 min) and individually tested for its ability
to support lymphocyte proliferation. Sera supporting acceptable levels of pro-liferation (usually around 80% of tested samples) are pooled and used at 10–
20% v/v with culture medium (RPMI 1640 or IMDMEM). However, note that
CD34+ cells cannot be grown in either FCS- or HS-containing medium. Two
factors recently enabled us successfully to grow these cells and facilitate
their differentiation into mature T-cells. The first was to use the serum-free
medium X-Vivo 10 without adding any other serum supplement. This medium
is also suitable for the cloning and long-term propagation of TCC derived
Human T-Cell Clones 59
from mature T-cells. The second factor was to use a suitable cytokine cocktail
that supported the viability of the stem cells and also allowed T-cell growth to
take place. This consisted of stem cell factor (SCF), flt3-L, and IL-3, together
with IL-2 and either oncostatin M (OM) or IL-7.
3. Use autologous PBMCs, a mixture of autologous PBMCs and autologous
B-lymphoblastoid line cells, or other appropriate antigen-presenting cells (APC),
in the presence of specific antigen. Alternatively, use an antigen-non-specific
stimulus such as 50 ng/mL of the anti-CD3 monoclonal antibody OKT3 or
2 µg/mL of purified or 1% crude PHA, together with the same number of alloge-neic or autologous PBMCs, or pooled PBMCs (irradiated at 30 Gy).
4. Clones successfully propagated in cluster plate wells for 2 wk can be taken to be
established. They can at this point be cryopreserved, although it is advisable to
retain some of each clone in culture to test different conditions to establish opti-mal parameters for each particular clone. Human TCCs can be readily cryo-preserved using the same protocols as are suitable for freezing resting T-cells.
Having a frozen stock enables the different culture conditions to be tested to
optimize growth, without risking the loss of the whole clone. Restimulation
parameters should be established for each clone. T-cells require periodic reacti-vation through the T-cell antigen receptor to retain responsiveness to growth fac-tors. This can be accomplished either specifically or nonspecifically. All clones
can be propagated with weekly restimulation; some but not all can be propagated
with restimulation only every 2 wk. It should be established whether each clone
can be propagated with the most convenient feeder cells (80 Gy-irradiated
B-LCL) instead of PBMC feeders. Most TCCs flourish on B-LCLs alone, but
some need the presence of PBMCs as well (this is especially true during cloning).
Propagation of the TCCs on PBMC feeders can also be continued, but for practi-cal reasons it may often be more convenient and easier to grow large amounts of
B-LCLs than to isolate the PBMCs.
5. For convenience, it is also easier to grow TCCs in scaled-up culture vessels than
in 16 mm-diameter culture wells. However, not all clones can be adapted to
growth in flasks. This has to be tested for each clone, using between 1 × 105 and
5 × 105/mL of TCCs with an equal number of feeders in tissue culture flasks.
Clones not growing under these conditions can rarely be adapted to growth in
flasks by altering the amounts or concentrations of TCCs or feeders or by increas-ing or decreasing the frequency of stimulation and/or feeding.
6. Longevity estimation in PD is an extremely conservative measurement indicating
the absolute minimum number of cell divisions achievable by each cell. This is
because it simply assumes that all daughter cells at each cell division are viable
and themselves capable of dividing and generating two viable progeny. In reality,
it is highly likely that this is not the case.
Work in the author’s laboratory is supported by the Deutsche Forschungs-gemeinschaft, the Dr. Mildred Scheel Foundation, the Dieter Schlag Founda-
60 Pawelec
tion, the VERUM Foundation, the Novartis Foundation for Gerontological
Research, the ƒortüne Program of University of Tübingen Medical Faculty,
and the European Commission (see http://www.medizin.uni-tuebingen.de/
1. Bach, F. H., Inouye, H., Hank, J. A., and Alter, B. J. (1979) Human T-lymphocyte
clones reactive in primed lymphocyte typing and cytotoxicity.  Nature 281,
2. Pawelec, G. and Wernet, P. (1980) Restimulation properties of human alloreactive
cloned T cell lines. Dissection of HLA-D-region alleles in population studies and in
family segregation analysis. Immunogenetics 11, 507–519.
3. Morgan, D. A., Ruscetti, F. W., and Gallo, R. C. (1976) Selective in vitro growth of
T-lymphocytes from normal human bone marrows. Science 193, 1007–1008.
4. Effros, R. B. and Walford, R. L. (1984) T cell cultures and the Hayflick limit. Hum.
Immunol. 9, 49–65.
5. Pawelec, G. (1985) Functions and changing activities of interleukin 2-dependent
human T lymphocyte clones derived from sensitization in mixed leukocyte cul-tures, in T Cell Clones (von Boehmer, H. and Haas, W., eds.), Elsevier, Amsterdam,
Holland, pp. 311–322.
6. Johnson, J. P., Cianfriglia, M., Glasebrook, A. L. and Nabholz, M. (1982) Karyo-type evolution of cytolytic T cell lines, in Isolation, Characterisation, and Utilisation
of T Lymphocyte Clones (Fathman, C. G., Fitch, F. W., eds.), Academic Press, New
York, pp. 183–191.
7. Kaltoft, K., Pedersen, C. B., Hansen, B. H., and Thestrup-Pedersen, K. (1995)
Appearance of isochromosome 18q can be associated with in vitro immortalization
of human T lymphocytes. Cancer Genet. Cytogenet. 81, 13–16.
8. Kahle, P., Wernet, P., Rehbein, A., Kumbier, I., and Pawelec, G. (1981) Cloning of
functional human T lymphocytes by limiting dilution: impact of feeder cells and
interleukin 2 sources on cloning efficiencies. Scand. J. Immunol. 4, 493–502.
9. Pawelec, G., Schwuléra, U., Blaurock, M., Busch, F. W., Rehbein, A., Balko, I., and
Wernet, P. (1987) Relative cloning efficiencies and long-term propagation capacity
for T cell clones of highly purified natural interleukin 2 compared to recombinant
interleukin 2 in man. Immunobiology 174, 67–75.
10. Effros, R. B. and Pawelec, G. (1997) Replicative senescence of T lymphocytes:
does the Hayflick Limit lead to immune exhaustion? Immunol. Today 18, 450–454.
11. Pohla, H., Adibzadeh, M., Buhring, H. J., Siegels-Hubenthal, P., Deikeler, T.,
Owsianowsky, M., Schenk, A., Rehbein, A., Schlotz, E., Schaudt, K., and Pawelec,
G. (1993) Evolution of a CD3+CD4+ alpha/beta T-cell receptor+ mature T-cell clone
from CD3-CD7+ sorted human bone marrow cells. Dev. Immunol. 3, 197–210.
12. Preffer, F. I., Kim, C. W., Fischer, K. H., Sabga, E. M., Kradin, R. L., and Colvin, R.
B. (1989) Identification of pre-T cells in human blood. Extrathymic differentiation
of CD7+CD3- cells into CD3+ gamma/delta or alpha/beta + T cells. J. Exp. Med.
170, 177–190.
Human T-Cell Clones 61
13. Adibzadeh, M., Pohla, H., Rehbein, A., and Pawelec, G. (1995) Long-term culture
of monoclonal human T lymphocytes: models for immunosenescence? Mech. Age-ing. Dev. 83, 171–183.
14. Pawelec, G., Rehbein, A., Haehnel, K., Merl, A., and Adibzadeh, M. (1997) Human
T cell clones as a model for immunosenescence. Immunol. Rev. 160, 31–43.
15. Freedman, A. R., Zhu, H. H., Levine, J. D., Kalams, S., and Scadden, D. T. (1996)
Generation of human T lymphocytes from bone marrow CD34+ cells in vitro. Nat.
Med. 2, 46–51.
Telomeres and Replicative Senescence 63
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Telomeres and Replicative Senescence
Hector F. Valenzuela and Rita B. Effros
1. Introduction
Telomere length measurement can be used both to monitor the proliferation
of long-term cultures of somatic cells as well as to determine the replicative
history of in vivo-derived cells. The most frequently used technique for telom-ere length measurement is Southern hybridization (1,2). The method consists
of isolating total genomic DNA, digesting the DNA with restriction enzymes
so as to isolate the undigested telomere restriction fragments (TRFs), and sepa-rating these fragments by gel electrophoresis. The DNA is denatured and trans-ferred from the gel to a membrane or filter, and the DNA samples are then
hybridized to radiolabeled complementary probe. However, when blotting TRF
DNA to the membrane, differential transfer may occur owing to inefficient
transfer of larger fragments of DNA (>10 kb) to a membrane. As the mean
length of the TRF is based on the assumption that the amount of telomeric
DNA (TTAGGG repeats) in a given TRF is proportional to the length (3,4), this
would lead to possible error in calculating the mean length of the telomeres.
The method that we present here avoids these potential problems by eliminat-ing the membrane blot step altogether and probing the gel directly.
The following protocol has been refined for measuring telomeric DNA
length from human cells. Similar protocols can be adapted to measure
telomere  lengths in cells from other species. However, researchers should
adjust the probe sequence for hybridization (not all species have the same
telomere sequence) and optimize the restriction enzymes to obtain TRF within
the resolvable molecular weight range of the gel because some species may
have extremely long telomeres. For more information regarding telomeres, we
suggest reading Kipling’s The Telomeres (5).
64 Valenzuela and Effros
The TRF assay method that we present here can be divided into three stages:
(1) isolation and digestion of genomic DNA; (2) gel electrophoresis, drying,
and hybridization; and (3) analysis of TRF length. We will emphasize in this
protocol the measurements of the TRF after the DNA isolation. As mentioned
previously, the protocol improves on the standard Southern blot procedure by
eliminating the DNA transfer from gel to membrane, thereby reducing the time
and labor involved. After digestion, the DNA fragments are separated in an
agarose gel, which is then dried. The gel is then denatured, neutralized, and
hybridized in a manner similar to the membrane in the usual Southern blot
method. Once the gel is washed, it can be analyzed directly by densitometry of
an autoradiograph or by using a phosphorimager (3).
2. Materials
1. Denaturing buffer solution: 1.5 M NaCl, 0.5 M NaOH. Dissolve 43.83 g of NaCl
and 10 g of NaOH in 400 mL of distilled water, then raise volume to 500 mL.
Store at room temperature.
2. Neutralization buffer solution (1.5 M NaCl, 0.5 M Tris-Cl): Dissolve 43.83 g of
NaCl and 39.4 g of Tris-Cl in 450 mL of distilled water. Adjust pH to 8.0 (with
approx 2 g of NaOH). Raise volume to 500 mL. Store at room temperature.
3. Hybridization buffer solution: Mix 64 mL of distilled water, 25 mL 20× saline
sodium citrate (SSC), 10 mL of Denhardt’s reagent (50×), and 1 mL of sodium
pyrophosphate (stock 1 M). Sterile filter with 0.22 µm filter. Store at 4°C. The
50× Denhardt’s reagent (Sigma Chemical, St. Louis, MO, USA) can be prepared
using 5 g of bovine serum albumin, 5 g of Ficoll, and 5 g of polyvinylpyrrolidine.
4. 20× SSC washing solution (3 M NaCl, 0.3 M sodium citrate): Dissolve 175.3 g of
NaCl and 88.2 g of sodium citrate in 800 mL of H2O. Adjust pH to 7 with a few
drops of NaOH. Raise volume to 1 L with distilled water. Prepare 1.5L of 0.5×
SSC for washes. Sterilize by autoclaving, and store at room temperature. The
solution is stable for several months.
5. Probe: The probe sequence can be either (TTAGGG)3 or (CCCTAA)3. Prepare an
aliquot concentration of 40 pmol/µL. Store at –20°C (see Subheading 3., step 4).
6. Ladders: A ladder that ranges from 1 to 20 kb is required. Alternatively, we advise
mixing two ladders, a 1 kb ladder at 1 µg/µL (Gibco-BRL) and λDNA HindIII
digest at 1 µg/µL (New England BioLabs). Store at –20°C.
7. Enzymes: Restriction enzymes  HinfI and  RsaI (New England BioLabs). Use
appropriate buffer to ensure maximum activity. Klenow polymerase (Gibco-BRL)
and T4 polynucleotide kinase (Gibco-BRL) are used for labeling ladder and
probe, respectively.
8. Quick Spin Columns Sephadex G25 Fine (Boehringer Mannhein cat. no.
9. Polyethylene bags for hybridization step. (Fisher bags cat. no. 01-812-10E; Fisher
International Headquarters, 50 Fadem Rd, Springfield, NJ 07081-3193, USA:
Tel: 201-467-6400; Fax: 201-379-7415).
Telomeres and Replicative Senescence 65
10. Isotopes: require [α-32P]ATP and [γ-32P]ATP for labeling ladder and probe,
respectively. Alternatively one can use 33P isotopes, but do not mix these differ-ent isotopes in the same gel. 33P is safer to handle but requires twice the exposure
time of 32P.
11. Gel apparatus: Gel cast of 15–20 cm long and thin combs (2 mm).
12. Whatman 3MM paper cut out to 2.5 cm longer than gel in both length and width.
3. Methods
1. Isolation of genomic DNA: The isolation of the genomic DNA can be performed
by any number of standard protocols in the literature. We recommend “DNAzol”
DNA Isolation reagent (Molecular Research Center, Inc., 5645 Montgomery
Road, Cincinnati, OH 45212, USA; Tel: 513-841-0900; Fax: 513-841-0080). The
main consideration in selecting a protocol should be to choose a method that
yields DNA fragments larger than 60 kb; otherwise results may be skewed (see
Note 1).
2. Digestion of genomic DNA: For the digestion of 10 µg of high molecular weight
DNA use the following recipe:
10 µg of Genomic DNA
10 µL of 10× reaction buffer
2 µL of HinfI
2 µL of RsaI
X µL dH2O
100 mL (final volume). Incubate at 37°C for 2–3 h.
Run 2 mL of digested DNA with undigested DNA on a mini-gel to test for comple-tion of digestions. Digestion is incomplete if there are fragments larger than 50 kb. The
amount of DNA loaded per lane must be at least 1–2 µg; a larger amount of DNA
increases the sensitivity of detection, especially for short telomeres.
3. Agarose gel: Pour a 0.5% agarose/0.5× TBE gel. The gel must be at least 10 cm
long (we recommend 15–20 cm) and must be approx 3/4 cm thick. The longer gel
allows good separation of large fragments of DNA and the thin combs prevent the
DNA from diffusing. Run gel for a total of 750 V/h. Do not run gel faster than
50 V, as this prevents good resolution of long telomere fragments. We recomend
30 V, which should then be run for 25 h (for a total of 750 V/h).
4. Loading DNA onto gel: Load at least 1–2 µg of DNA per lane. Labeling of 1 kb
and λDNA HindIII digest ladders is performed as follows in a 1.5-mL Eppendorf
tube. Ladders should be loaded last onto the gel to minimize exposure to radiation.
0.5 µL of 1 kb ladder
0.5 µLof λDNA HindIII digest ladder
4 µL of 10× Klenow buffer
3 µL of [α-32P]ATP
31 µL dH2O (or add dH2O to 40 µL final volume)
1 µL Klenow fragment
40 µL (final volume)
66 Valenzuela and Effros
Incubate 2 min at 37°C, and put on ice while you prepare the quick spin col-umn to remove the unincorporated [α-32P]ATP. For a freshly prepared ladder use
0.5 µL/lane (see Notes 2 and 3).
5. Gel drying: Place two sheets of 3MM Whatman chromatography paper in gel
drier, then place gel on top leaving 2.5 cm around the margins of the gel. Finally,
place Saran wrap over gel and Whatman paper (Fig. 1). Dry gel under vacuum at
60°C for 45–75 min.
Note: Start preparing the probe (step 7) and prewarm hybridization buffer
(step 8) (see Note 4)
6. Gel washes: Remove gel from vacuum dryer by holding gel at the opposite
end from the wells. Gels occasionally tear during this step, and by handling
the end furthest from the wells, the chance of tearing the gel in the area near
the DNA is reduced. One should keep in mind that at this point the gel is
radioactive from the labeled ladder, so all proper precautions should be main-tained. If Whatman paper sticks to gel, use water to peel them apart. Place the
gel in a Pyrex dish container and add enough denaturing reagent (500 mL) to
completely submerge gel. Let sit at room temperature for 10 min with gentle
shaking. Dispose of buffer by pouring into proper radioactive waste. Repeat
washing with neutralization buffer (500 mL) in the same Pyrex container for
10 min at room temperature with gentle shaking. Upon completion of this
wash, dispose of buffer in the same manner.
7. Probe labeling reaction can be prepared the following way in a 1.5-mL
Eppendorf tube:
1 µL of 40 pmol/µL of (TTAGGG)3 oligo
5.3 µL of 10× T4 polynucleotide kinase buffer
6.5 µL of [γ-32P]ATP
1.5 µL of T4 polynucleotide kinase
38.7 µL of dH2O
53 µL (final volume)
Fig. 1. Setup for drying the agarose gel in a gel dryer.
Telomeres and Replicative Senescence 67
Incubate reaction at 37°C for 30 min, then use a quick spin column to separate
unincorporated [γ-32P]ATP.
8. Hybridization of gel: Prewarm the hybridization buffer (15 mL) to 37°C. For a
gel 10–20 cm in length use 15 mL (but no more than 20 mL) of hybridization
buffer. Add 1 µL of label probe for every 1 mL of hybridization buffer. Keep at
37°C while you prepare the gel.
Prepare the gel for hybridization by placing the gel in a hybridization bag.
Perform this by cutting open two sides of the bag, so that it opens up like a book.
This method will prevent the gel from sticking to plastic that may lead to tearing
of the gel. Place gel inside and seal ends of plastic bag once again with an Impulse
Sealer so that there is only one opening. (There should be 1–2 cm of margin
space between the gel and the plastic bag.) Through this opening add the
prewarmed hybridization buffer with the added probe. Check the seals for leaks
over the Pyrex container before proceeding. Caution: The hybridization solution
will be highly radioactive.
Before sealing the open end of the bag, remove any bubbles that may be in
the bag. (Remember to do this over a Pyrex container in case of leaks.) It may be
helpful in removing the final bubbles to seal the bag at the very edge, then squeeze
the remaining bubbles to one edge, sealing them off with a second seal. You must
remove most of the bubbles before the second seal for this double sealing to
9. Incubate the gel at 37°C for at least 6 h (although we recommend overnight). For
very weak probes, 2–3 d may be necessary.
10. Washing with SSC: Cut open plastic bag, remove the hybridization solution, and
dispose in a proper radioactive waste receptacle. Carefully place gel in a Pyrex
container and add 500 mL of 0.5× SSC (prewarmed at 37°C) for 6–7 min. Repeat
the same wash two more times. Remove all SSC, then enclose the gel in Saran
Wrap before exposing to imaging film (see Note 5).
11. Analysis: Analysis of the TRF length can be done either by densitometric scan-ning of the autoradiogram or by using a phosphoimager (see Note 6)
4. Notes
1. For the isolation of genomic DNA, we recommend using a guanidium based
method, such as DNAzol® (Molecular Research Center). This method is fast
(30 min) and reliable; however, we emphasize the importance of using wide-mouth
tips to prevent DNA shearing during the isolation steps.
2. Along with the samples, we recommend including two control DNA fragments,
one from cells with long telomeres and the other from cells with short telomeres.
For the long telomere source, one can use a subline of the 293 tumor, which has
stable TRF length of approx 10.5 kb. As source of short telomeres, Daudi cells,
with a mean TRF length of approx 3.9 kb, can be used. Prepare a large batch of
genomic DNA from the control cell lines, aliquot into small amounts, and use
these as standards to compare gel-to-gel variations in the sizes of TRF that may
occur between experiments. It may be necessary to add more than 2 µg of DNA
68 Valenzuela and Effros
for cells that have short telomeres. Longer telomeres, because they have more
TTAGGG repeats, give a stronger signal than short telomeres.
The term “TRF length” is not synonymous with “telomere length.” The TRF
includes both the telomere repeats and the adjacent subtelomeric region that con-tains both repetitive and nonrepetitive sequences .
3. We suggest using ladders at both ends of the gel. Use 0.5 µL/lane of ladder label-ing reaction mix when using freshly prepared material. If the ladder was prepared
at an earlier time, add up to 1 µL/lane, but beware of adding too much lest the
signal be overexposed, making it impossible to quantitate TRF length. If this
problem occurs, one may either add more (>2 µg/lane) genomic DNA, or a smaller
quantity of labeled DNA ladder per lane (but not less than 5 µg/lane).
Gel orientation can be marked in a number of ways, such as by cutting away a
corner of the gel, loading two lanes with the ladder on one side of gel, or loading
control TRF DNA on one side of gel.
4. Note that a 0.5% agarose gel is extremely fragile, and we suggest using a spatula
or the back of a Pyrex dish during all transfers. Gels can be stored for up to 2 d at
this stage, for example, if the probe is not ready. This can be done by placing the
gel in a sealed plastic bag with 2 mL of 2× SSC at 4°C, as described in Subhead-ing 3.
5. If the background radiation is too high, place gel once for 1 h (or twice for 30 min
each) at 48°C in 0.1× SSC. Shortening exposure time may also reduce back-ground. If the problem continues, one should increase the amount of DNA, which
reduces the noise by reducing the background. Adding lower concentrations of
probe will also help reduce the background.
If there is no signal, lengthen exposure time. Verify that probe was properly
labeled. Verify that hybridization solution has no precipitates. If precipitate is
seen, it is necessary to prepare fresh hybridization solution.
6. The mean TRF length is calculated by integrating signal intensity over TRF dis-tribution on gel as a function of mol wt. Divide a scanned TRF image into a grid
in which columns cover the entire length of TRF sample analyzed and there are at
least 30 boxes dividing each column (Fig. 2). The following equation can then be
used to calculate the mean TRF length:
Mean TRF length = Σ(ODi·Li) / Σ(ODi)
where ODi is the intensity signal and Li is the mol wt at a particular (i) box in the
grid as compared to a size marker from a ladder. Before using above equation for
TRF length analysis, background must be subtracted from ODi. For each sample
the background can be calculated by averaging the OD from the top two boxes
and bottom two boxes adjacent to the smear. The average background OD is then
subtracted from each box in the grid for that particular sample. Alternatively,
some phosphorimagers (Packard, Instant Imager) come included with software
that can quantitate the intensities of signals to obtain mean values that are plotted
to preassigned values from a ladder.
Telomeres and Replicative Senescence 69
The authors wish to thank Dr. Choy-Pik Chiu (Geron Corporation, Menlo
Park, CA) for providing comments on the manuscript, and the Geron Corpora-Fig. 2. Calculation of mean TRF length. Exposed image has been divided into col-umns and rows where OD and L values can be measured. For column 3, the boxes used
for calculating the background are indicated.
70 Valenzuela and Effros
tion for their generous help in establishing the telomere assays in our labora-tory. This work was supported by the National Institute on Aging (AG10415),
the UCLA AIDS Institute, and the Seigel Life Project/UCLA Center on Aging.
1. Southern, E. M. (1975) Detection of specific sequences among DNA fragments
separated by gel electrophoresis. J. Mol. Biol. 98, 503.
2. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning A Labora-tory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, pp.
3. Allsopp, R. C., Vaziri, H., Patterson, C., Goldstein, S., Younglai, E. V., Futcher,
A. B., Greider, C. W., and Harley, C. B. (1992) Telomere length predicts replicative
capacity of human fibroblasts. Proc. Natl. Acad. Sci. USA 89, 10,114–10,118.
4. Harley, C. B., Futcher, A. B., Greider, C. W. (1990) Telomere shorten during ageing
of human fibroblasts. Nature 345, 458–469.
5. Kipling, D. (1995)  The Telomere, Oxford University Press, pp. 1–12, 78–96,
130–142, 146–163.
Detection of Molecular Events 71
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Detection of Molecular Events
During Apoptosis by Flow Cytometry
Ruaidhri J. Carmody,* Ana P. Costa-Pereira,* Sharon L. McKenna,
and Tom G. Cotter
1. Introduction
Apoptosis describes an intrinsic cell suicide program that may be activated
by both endogenous and exogenous stimuli. This method of cell death is char-acterized by specific morphological features including chromatin condensa-tion, nuclear fragmentation, cell shrinkage, membrane blebbing, and the
formation of membrane-bound vesicles termed apoptotic bodies (1). Apoptosis
has come to be referred to as the physiological mode of cell death, as it allows
cellular destruction in the absence of an associated inflammatory response.
In contrast, necrosis is a pathological mode of cell death that occurs under
circumstances of severe cellular injury/trauma. Necrotic cell death involves
cell swelling and organelle disruption, followed by lysis and release of cellular
debris. This form of cell death may cause damage to surrounding tissue due to
the inappropriate triggering of an inflammatory response (2).
Apoptosis occurs during normal mammalian development and also plays an
important role in subsequent tissue homeostasis by balancing cell division with
cell death (3). Apoptosis has also been ascribed a role in several disease states
including malignancy and neurodegenerative disorders, in which deregulation
or inappropriate activation of the apoptotic program contributes to the observed
pathology. The physiological importance of apoptosis in the maintenance of
tissue homeostasis, and the observation of apoptotic cell death in age-related
degenerative disorders such as Alzheimer’s disease and Parkinson’s disease
(reviewed in ref. 4), suggests that the regulation of apoptosis may be a critical
parameter for consideration in studies of aging.
*These authors have contributed equally to this work.
72 Carmody et al.
This chapter outlines some current procedures for the detection of apoptosis
and the analysis of intracellular molecular events important in apoptotic path-ways. Biochemical events include the generation of reactive oxygen species
(ROS) and disruption of mitochondrial transmembrane potential (∆ψm). The
methods described in this chapter all utilize a flow cytometer for quantitative
analysis of data. Several techniques (e.g., propidium iodide [PI] or terminal
deoxyuridine triphosphate [dUTP] nick end labeling [TUNEL]) assay may, how-ever, be adapted for use with a fluorescent microscope. Flow cytometric analy-sis has the advantage of a rapid assessment of large numbers of cells in a highly
quantitative, nonsubjective manner. In addition, flow cytometry enables the par-allel assessment of two and possibly three parameters in the same cell. Cell
sorting may also be an option for some users (see also subsequent sections).
1.1. Translocation of Phosphatidylserine
Several studies have shown that phosphatidylserine (PS) is asymmetrically
distributed and is preferentially located in the inner leaflet of the plasma mem-brane (PM) (5). In the early stages of apoptosis in virtually all cell types, redis-tribution of membrane phospholipids results in the exposure of PS on the outer
membrane (6). PS exposure has been shown to be tightly coupled to other
apoptosis-associated changes (7), and seems to be stimulus independent (6).
Externalization of PS, following the induction of apoptosis, can be readily
detected using recombinant Annexin V, a protein that binds with high affinity
to PS in a Ca2+-dependent manner (8). Because necrotic cells have lost their
membrane integrity, they may also stain positive with Annexin V. However,
dual staining with PI enables membrane-disrupted cells to be readily distin-guished. Secondary necrotic cells are apoptotic cells which have subsequently
lost membrane integrity (see Subheading 4.1.1.), and therefore cannot be
distinguised from primary necrotic cells using this method.
As Annexin V detects a cell surface marker, no fixation or permeabilization
is required for the procedure. The cells are therefore analyzed “alive” and may
be recovered by fluorescence-activated cell sorter (FACS) sorting for further
analytical purposes. This method also facilitates dual labeling for surface anti-gens that recognize native antigen conformations.
1.2. DNA Fragmentation
The degradation of DNA into oligonucleosomal sized fragments of 180–200
basepairs by specific endonuclease activity is a major biochemical event dur-ing apoptosis in most cell types (9). This DNA fragmentation was originally
observed as a ladder pattern using agarose gel electrophoresis. However, this
method of detection is essentially qualitative and does not allow for the identi-fication of subpopulations of apoptotic cells. Several flow cytometric methods
Detection of Molecular Events 73
have been described that allow the quantitative analysis of DNA fragmentation
as well as the parallel measurement of other parameters such as cell cycle and
antigen expression (10,11). The most commonly used of these methods is the
TUNEL assay.
The TUNEL assay is based on the addition of biotinylated dUTP nucleotide
to 3′ hydroxyl termini at DNA strand breaks. This reaction is catalyzed by the
enzyme terminal deoxynucleotidyl transferase that repetitively adds deoxyri-bonucleotide to the 3′ hydroxyl termini of DNA. Fluoresceinated avidin is then
employed in a second step reaction to fluorescently label the DNA strand
breaks, thus allowing the detection of DNA strand breaks by flow cytometry.
1.3. Reactive Oxygen Species
The generation of ROS and alterations in the cellular redox state have been
reported to be a common biochemical event in apoptosis (12). Moreover, ROS
have been proposed to be putative second messengers in the initiation of
apoptosis. The production of ROS during cytotoxic drug induced apoptosis
and inhibition of apoptosis by antioxidants supports this view (12). Oxidative
stress is also believed to play a role in Parkinson’s disease and amylotrophic
lateral sclerosis disease states, the latter of which has been linked to genetic
lesions in a cellular antioxidant pathway (4). Possible sources of intracellular
ROS include the depletion of cellular antioxidants such as glutathione, disrup-tion of mitochondrial respiration, and the activation of oxidant-producing
enzymes such as NADPH oxidase.
The fluorescent probes 2′,7′-dichlorofluorescein diacetate (DCFH/DA) and
dihydorethidium (DHE) may be used for the measurement of intracellular per-oxide and superoxide anion levels, respectively. DCFH/DA is cell permeant
and is nonfluorescent until the acetate groups are removed by cellular esterase
activity and a peroxide group is subsequently encountered. Hydrolyzed, oxi-dized DFCH/DA fluoresces at 529 nm (FL-1, log scale; see Subheading 3. and
Ta ble 1) (Fig. 1A,B) and is unable to leave the cell, thus allowing the measure-ment of intracellular peroxides by flow cytometry.
DHE is also cell permeant and is oxidised to ethidium by superoxide anion.
Once oxidised, ethidium is free to intercalate with DNA in the nucleus where-upon it emits fluorescence at 605 nm (FL-2) (see Fig. 1A,C).
1.4. Mitochondrial Transmembrane Potential Alterations
Emerging evidence suggests a central role for mitochondria during apoptosis
induced by a diverse range of stimuli in a number of cell types. Indeed, several
groups have proposed mitochondria as the central executors of apoptosis
(reviewed in ref. 13). Mitochondrial events during apoptosis include release of
cytochrome-c and disruption of the transmembrane potential (∆ψm) (13). Dis-
74 Carmody et al.
ruption of  ∆ψm is believed to occur through permeability transition (PT), a
process that involves the opening of the mitochondrial PT pore, allowing
release of solutes 1.5 kDa and smaller and subsequent disruption of  ∆ψm.
Importantly, inhibitors of PT also inhibit apoptosis in several models of
apoptosis, supporting the view that disruption of mitochondrial function is
central to the apoptotic process.
The cell permeant fluorescent probe JC-1 can be employed to monitor
changes in ∆ψm in intact cells. In the presence of a high ∆ψm JC-1 forms what
are termed J-aggregates that fluoresce strongly at 590 nm (FL-2). Reduced
∆ψm results in a reduced FL-2 signal in JC-1-stained cells (Fig. 2A,B). This
method has been demonstrated to be quantitative in addition to qualitative and
allows subpopulations of cells with different mitochondrial properties to be
identified (14).
2. Materials
2.1. Annexin V Assay
1. Fluoresceinated Annexin V (Annexin V-FITC) (e.g., Bender MedSystems,
Heidelberg, Germany). Protect from light and store at –20°C.
2. Binding buffer: 10 mM N-[2-hydroxyethyl]piperazine-N’-[2-ethanesulfonic acid]
(HEPES)/NaOH, pH 7.4, 140 mM NaCl, 2.5 mM CaCl2. Store at 4°C.
3. PI. Protect from light and store at 4°C.
4. Phosphate-buffered saline (PBS): 8.06 mM Na2HPO4, 1.47 mM KH2PO4, pH 7.4;
2.27 mM KCl, and 137 mM NaCl.
2.2. Terminal dUTP Nick End Labelling (TUNEL) Assay
1. Fixation buffer: 2% (w/v) p-Formaldehyde in PBS, pH 7.4 (see Subheading 2.1.
for PBS formulation).
Table 1
Summary of Assays and Probes Described in this Chaptera
Probe/Assay Parameter measured Emission (nm) Channel
Annexin-V PS translocation 515 FL-1
TUNEL DNA fragmentation 515 FL-1
Antigen analysis Antigen expression 580 FL-2
PI DNA content 620 FL-2
DHE Superoxide anion 605 FL-2
DCFH/DA Peroxide 529 FL-1
JC-1 Mitochondrial membrane 590 FL-2
aThe table includes the emission peak of probes and the channel through which data should
be collected and analyzed.
Detection of Molecular Events 75
Fig. 1.  (A) Schematic diagram illustrating the analysis of intracellular peroxides
and superoxide using the fluorescent probes DCFH/DA and DHE. Hydrolysis and oxi-dation of DCFH/DA causes it to fluoresce in FL-1, while the oxidation of DHE to
ethidium results in an increase in fluorescence in FL-2 owing to the intercalation of
ethidium with cellular DNA. (B) Production of peroxides in DU145 prostate cancer
cells treated with camptothecin. Peroxide levels were assessed in untreated DU145
prostate cells (dashed line) treated with 10 µg/mL of camptothecin (solid line) and in
cells treated with 1 mM H2O2 (dotted line) for 1 h, as described in Subheading 2.3.
After 1 h of treatment with 10 µg/mL of camptothecin, there is an increase in peroxide
production that can be seen as a shift to the right in relative fluorescence.  (C)
Measurment of superoxide anion in retinal cell primary cultures after 24 h. Retinal
cells cultures display high levels of apoptosis after 24 h in culture (see Fig. 4-2). Stain-ing cells with DHE reveals a significant increase in superoxide levels at 24 h (solid
line) relative to immediately isolated cells (dashed line).
76 Carmody et al.
Fig. 2.  (A) Schematic diagram of mitochondrial events during apoptosis. Intact
mitochondria display high transmembrane potential (∆ψm) and fluoresce strongly in
FL-2 when stained with JC-1. Apoptotic mitochondria undergo permeability transition
resulting in a release of solutes 1.5 KDa and smaller and a loss transmembrane poten-tial, consequently displaying reduced fluorescence in FL-2 when stained with JC-1.
(B) Alterations in  ∆ψm in NSO myeloma cells upon camptothecin treatment. NSO
myeloma cells were treated with 30 mM ammonia for 18 h. Depolarization in ∆ψm was
assessed as described in  Subheading 2.3., using JC-1.  ∆ψm depolarization can be
monitored by measuring the fluorescence in FL-1. Membrane depolarization results in
an increase in FL-1 flourescence. Treated cells (solid line) show an increase in FL-1
fluorescence relative to untreated cells (dashed line).
Detection of Molecular Events 77
2. Elongation buffer: 0.2 M Potassium cacodylate; 25 mM Tris-HCl, pH 6.6; 2.5 mM
cobalt chloride; 0.25 mg/mL of bovine serum albumin (BSA); 100 U/mL of terminal
deoxynucleotidyl transferase (TdT) (e.g., Boehringer Mannheim, East Sussex, UK);
0.5 nM biotin-16-dUTP (e.g., Boehringer Mannheim). Make fresh as required.
3. Staining buffer: Dilute 20× saline citrate buffer (0.3 M sodium citrate; 3 M NaCl,
pH 7.0) to 4×; add 2.5 mg/mL of fluoresceinated avidin, 0.1% (v/v) Triton X-100; 5% (w/v) nonfat dried milk, to give 0.6 M NaCl and 0.06 M sodium citrate.
Staining buffer is freshly made and protected from light.
2.3. Detection of Intracellular ROS
1. DCFH/DA (Molecular Probes, Leiden, The Netherlands) prepared as a 5 mM
stock in dimethyl sulfoxide (DMSO). Protect from light and store at –20°C.
2. DHE (Molecular Probes) prepared as a 10 mM stock in DMSO. Protect from
light and store at –20°C.
2.4. Measurement of Mitochondrial Transmembrane Potential
1. JC-1 (Molecular Probes), made as a 5 mg/mL stock in DMSO. Protect from light
and store at –20°C.
3. Methods
3.1. Annexin V Assay
1. Harvest 1–2.5 × 105 cells and resuspend in 200 µL of binding buffer.
2. Dilute Annexin V–flourescein isothiocyanate (FITC) as recommended by the
3. Add diluted Annexin V–FITC and incubate for 10 min at room temperature, in
the dark.
4. After a wash in PBS, resuspend cells in binding buffer and add PI (see Subhead-ing 4.1.2.) (to a final concentration of 5 µg/mL).
5. Quantitate Annexin V binding and PI staining by flow cytometry (FL-1 and
FL-2 respectively) (see Note 3) (see Subheading 3. and Table 1) (Fig.3).
3.2. TUNEL  Assay
1. Fixation and permeabilization (see Note 4): Harvest approx 1  × 106 cells and
suspend in 1 mL PBS. Add 1 mL of 2% (w/v) p-formaldehyde fixation buffer
(see Note 5). Leave on ice for 15 min. Wash once in PBS and resuspend in
2mL of –20°C 70% (v/v) ethanol. Leave at –20°C for at least two hours or up
to two weeks.
2. Elongation: Wash fixed/permeabilized cells twice in PBS and resuspend in 50 µL
of elongation buffer. Incubate at 37°C for 30 min. Wash twice in PBS.
3. Staining: Resuspend cells in 100 µL of staining buffer and incubate in the dark at
room temperature for 30 min.
78 Carmody et al.
4. Wash twice in PBS and keep on ice and in the dark until read on a flow
5. Measure green fluorescence (labeled DNA strand breaks) following excitation
at 488 nm using a 525 nm band pass filter (FL-1, log scale; see Subheading 3.
and Ta ble 1).
3.2.1. Analysis of Cell Cycle in Conjunction with TUNEL Assay
After the TUNEL assay procedure has been completed resuspend cells in
1mL of PBS containing 5 µg/mL of PI, and 0.1% DNase-free RNase A. Cell
cycle analysis can then be carried by measuring the red fluorescence of PI at
>600 nm (FL-2, linear scale; see Subheading 3. and Table 1) (Fig. 4-1).
3.2.2. Analysis of Antigen Expression in Conjunction
with TUNEL Assay
The combination of TUNEL staining with immunofluorescence labeling of
a cell-specific antigen allows the subsequent analysis of apoptosis in specific
subpopulations. Cells may be labeled for antigen expression prior to or follow-ing fixation/permeabilization depending on the nature of the epitope. It is rec-ommended that it first be determined whether epitope labeling is affected by
the fixation/permeabilization process. The following procedure detects an intra-Fig. 3. Annexin V/PI dual staining of human T cells treated with Actinomycin
D. Jurkat cells were treated for 4 h with 15 mg/mL of Actinomycin D. Cells were
stained with Annexin V-FITC (FL-1, y-axis) and PI (FL-2, x-axis) as described in
Subheading 2.1. Normal cells are negative for both Annexin and PI and appear in
the lower left quadrant. Apoptotic cells with intact membranes stain with Annexin,
but not with PI and therefore appear in the upper left quadrant. Primary and sec-ondary necrotic cells have disrupted membranes and stain with both Annexin V
and PI (upper right quadrant).
Detection of Molecular Events 79
Fig. 4-1. Analysis of apoptosis by TUNEL and cell cycle in human leukemic cells
treated with Etoposide. HL-60 cells (myeloid leukemia cell line) were treated for 3
and 6 h with 40 µM Etoposide (VP16). Cells were stained with FITC using the TUNEL
procedure as described in Subheading 2.2. Dual staining with PI enables cell cycle
parameters to be assessed in parallel with apoptosis. This data shows that after a 3-h
incubation with VP16 the majority of S-phase cells have initiated endonucleolytic DNA
cleavage. Some cleavage is also evident in G0/G1 cells, whereas G2/M cells are rela-tively resistant. After 6 h, most of the unlabeled G2/M cells have disappeared. Some of
these cells may have labeled with FITC, although most appear to have cycled through
to G0/G1, where the cell cycle is blocked.
80 Carmody et al.
Fig. 4-2. The detection of rod photoreceptor apoptosis in a primary retinal cell cul-ture using dual labeling for rhodopsin expression (rod specific protein) and TUNEL.
(A) Retinal culture stained with anti-rhodopsin antibody that was subsequently labeled
with a phycoerytherin-conjugated secondary antibody. The encircled rhodopsin-posi-tive population (high FL-2, y-axis) can be readily distinguished from nonrod cells in the
retinal culture. (B) TUNEL of rhodopsin-positive cells after 24 h in culture displaying
high levels of apoptosis. The dashed line represents a negative control of cells stained
without the TdT enzyme while the  solid line  represents TUNEL stained rhodopsin-positive cells.
cellular antigen. The cells must therefore be labeled with the specific antibody
following fixation.
1. Once the TUNEL assay protocol is completed, incubate cells in appropriately
diluted (using PBS/1% BSA, w/v) primary antibody for 60 min.
2. Wash twice in PBS and once in PBS/BSA (0.1%, w/v).
3. Incubate cells in appropriately diluted (using PBS/1% BSA) phycoerythrin-conjugated secondary antibody for 30 min.
Detection of Molecular Events 81
4. Antigen expression is analyzed by measuring red fluorescence at >600 nm (FL-2,
log scale; see Subheading 3. and Ta ble 1) (Fig. 4-2).
3.3. Measurement of ROS
3.3.1. Measurement of Intracellular Peroxide Levels
1. Incubate cells (see Notes 6 and 7) (5 × 105/mL) with 5 µM DCFH/DA, for 1 h at
37°C, in the dark.
2. Assess peroxide levels using a FACScan flow cytometer with excitation and emis-sion settings of 488 nm and 530 nm, respectively (FL-1, log scale; see Subhead-ing 3. and Ta ble 1) (Fig. 1B).
3.3.2. Measurement of Intracellular Superoxide Levels
1. Incubate cells (see Subheadings 4.3.1. and 4.3.2.) (5  × 105/mL) with 10  µM
hydroethidine, for 15 min at 37°C, in the dark.
2. Superoxide levels are assessed using a FACScan flow cytometer with excitation
and emission settings of 488 nm and 600 nm respectively (FL-2) (see Subhead-ing 3. and Ta ble 1; Fig. 1C).
3.4. Measurement of Mitochondrial Transmembrane Potential
1. Incubate cells (5 × 105/mL) with 5 µg/mL of JC-1 for 15 min at 37°C, in the dark.
2. Ratiometric measurements are performed using a FACScan flow cytometer, in
FL-1 and FL-2 (log scales; see Subheading 3. and Table 1) (Fig. 2A).
3.5. Acquisition and Analysis of Flow Cytometric Data
The assays described in this chapter were performed on a FACScan flow
cytometer (Beckton Dickinson). An excitation source of 488 nm was obtained
using a 15-mW air-cooled argon ion laser. Fluorescence emission was collected
through a 530/30 band pass filter (FL-1) on a log scale for TUNEL, DCFH/DA,
JC-1, and Annexin-V assays and a 585/42 band pass filter log scale (FL-2) for
PI, DHE, JC-1, and antigen labeling assays, while cell cycle analysis (PI) was
conducted using a linear scale (FL-2 area). A minimum of 5000 events were
collected for each sample. CellQuest™ software version 1.1.1 was used for
both acquisition and analysis of data.
4. Notes
4.1. Annexin V Assay
1. Under physiological conditions apoptotic cells are phagocytosed prior to loss of
membrane integrity.
2. Following addition of PI, cells should be analyzed with minimal delay, as PI may
eventually leak into normal and apoptotic cells.
3. As some cells stain very brightly with FITC, it may be necessary to use FACS com-pensation (FL-2-FL-1) during data acquisition (see Subheading 3. and Ta ble 1).
82 Carmody et al.
4.2. TUNEL  Assay
4. If an assessment of necrosis is required in a given population this can be achieved
prior to fixation by employing the PI exclusion assay as described in Subhead-ing 2.1.
5. This avoids any clumping of cells that may occur if a 1% (w/v) p-formaldehyde
fixation buffer were added directly to samples.
4.3. Detection of Reactive Oxygen Species
6. Cells can be treated with apoptosis-inducing agents either before, after, or during
the incubation period, depending on the time point at which ROS levels are to be
7. As a positive control for peroxide production, cells may be treated with 1 mM
H2O2 for 30–60 min (see Fig. 1B).
This work was supported by the Foundation for Science and Technology
(Fundação para a Ciência e a Tecnologia), Lisbon, Portugal, RP Ireland Fight-ing Blindness, and the EU Biotech Programme.
1. Kerr, J. F. R., Wyllie, A. H., and Currie, A. R. (1972) Apoptosis, a basic biological
phenomenon with wider implications in tissue kinetics. Br. J. Cancer 26, 239–245.
2. Trump, B. F., Beresky, I. K., and Osornio-Vargas, A. R., eds. (1981) in Cell Death
in Biology and Pathology, Chapman and Hall, New York.
3. Raff, M. C. (1992) Social controls on cell survival and cell death. Nature 365, 397–400.
4. Gorman, A. M., McGowan, A. J., O’Neill, C., and Cotter, T. G. (1996) Oxidative
Stress and apoptosis in neurodegeneration. J. Neurol. Sci. 139, 45–52.
5. Devaux, P. F. (1991) Static and dynamic lipid asymmetry in cell membranes. Bio-chemistry 30, 1163–1173.
6. Martin, S. J., Reutelingsperger, C. P. M., and Green, D. R. (1996) Annexin V- a
specific probe for the detection of phosphatidylserine exposure on the outer plasma
membrane leaflet during apoptosis, in  Techniques in Apoptosis: A Users Guide
(Cotter, T. G. and Martin S. M., eds.), Portland Press, London, pp. 107–119.
7. Martin, S. J., Reutelingsperger, C. P. M., McGahon, A. J., Rader, J., van Schie, R.
C. A. A., La Face, D. M., and Green, D. R. (1995) Early redistribution of plasma
membrane phosphatidylserine is a general feature of apoptosis regardless of the
initiating stimulus: Inhibition by overexpression of Bcl-2 and Abl.  J. Exp. Med.
182, 1545–1556.
8. Swairjo, M. A., Concha, N. O., Kaetzel, M. A., Dedman, J. R., and Seaton, B. A.
(1995) Ca2+-bridging mechanism and phospholipid head group recognition in the
membrane-binding protein Annexin V. Nat. Struct. Biol. 2, 968–974.
9. Arends, M. J., Morris, R. G., and Wyllie, A. H. (1990) Apoptosis: the role of the
endonuclease. Am. J. Pathol. 136, 593–608.
Detection of Molecular Events 83
10. Cotter, T. G. and Martin, S. J., eds. (1996) in Techniques in Apoptosis, Portland
Press, London.
11. Carmody, R. J., McGowan, A. J., and Cotter, T. G. (1998) Rapid detection of rod
photoreceptor apoptosis by flow cytometry. Cytometry, 33, 89–92.
12. McGowan, A. J., Ruiz-Ruiz, M. C., Gorman, A. M., Lopez Rivaz, A., and Cotter, T.
G. (1996) Reactive oxygen intermediates (ROI): common mediators of poly(ADP-Ribose)polymerase (PARP) cleavage and apoptosis. FEBS Lett. 392, 299–303.
13. Kroemer, G. (1997) Mitochondrial implication in apoptosis. Towards an endosymbi-ont hypothesis of apoptosis in evolution. Cell Death Different. 4, 443–456.
14. Salvioli, S., Ardizzoni, A., Franceschi, C., and Cossarizza, A. (1997) JC-1, but not
DiOC6(3) or rhodamine 123, is a reliable fluorescent probe to assess ∆ψm change
in intact cells: Implications for studies on mitochondrial functionality during
apoptosis. FEBS Lett. 411, 77–82.
Raf-1 Protein Kinase Activity 85
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Raf-1 Protein Kinase Activity
in T Cells from Aged Mice
Christopher J. Kirk and Richard A. Miller
1. Introduction
Most of our models of signal transduction through the T-cell receptor (TCR)
involve components and pathways first described in T-cell clones and T-cell
lymphomas such as the Jurkat cell line (1). These studies, while providing valu-able insights, are not always reliable guides to the analogous biochemical
events in cells freshly isolated from live donors (2). Thus, studies of the effects
of aging on T-cell activation must frequently begin with a detailed study of the
responses of cells from young individuals. Studies of aging effects involve addi-tional challenges, including the difficulty of purifying sufficient numbers of
cells from specific subsets, and allowing for the inherent variability among
donors of any age.
The mitogen-activated protein kinase (MAPK) pathway involves the sequen-tial activation of three kinases—Raf-1, mitogen-activated protein kinase
(MEK), and extracellular-signal-regulated kinase (ERK)—and plays an impor-tant role in T-cell activation (3). Here we describe an in vitro kinase assay for
Raf-1, which utilizes Raf-1 specific antibodies and a recombinant substrate, to
assess age-related differences in Raf-1 activation in mouse splenic CD4+ T-cells
stimulated through the TCR. The problems involved in analyzing Raf-1 activ-ity levels in freshly isolated T-cells are similar to those that are likely to be
faced in the study of age-related alterations in the signal transduction of other
cell types.
1.1. Isolation of Primary Lymphocytes
T-cell populations are made up of many distinct subsets, each with different
activation requirements, some of which change systematically with age. Thus,
86 Kirk and Miller
experiments on unseparated T-cell pools are likely to confound age effects on
activation pathways with the effects of subset transitions. Aging leads to an
increase in memory cells in both the CD4 and CD8 pools as measured by the
increase in CD44hi cells (4). The naïve and memory pool can be further subdi-vided based on differences in expression of the membrane glycoprotein, P-gly-coprotein, which also shows increased expression with age (5). It thus seems
reasonable to study activation pathways first in separated CD4 and CD8 sub-sets, and to include studies of separated subsets wherever practical.
Purification of these subsets from spleens begins with the depletion of eryth-rocytes and B cells by differential centrifugation and panning on anti-IgG-coated plates, respectively  (6). These procedures typically produce 25–35  ×
106 T-cells from a single mouse spleen, among which 85–95% express the CD3
ε-chain characteristic of T-cells. For CD4+ T-cell purification, as is described
in this chapter, CD8 cells are depleted by incubation with anti-CD8 antibodies
followed by removal with anti-IgG coated magnetic beads. Multiple subsets
can be depleted simultaneously; for example, addition of anti-CD44 and anti-CD8 can be used to purify CD4 naïve cells (i.e., CD4+CD45RBhi). Yields of
15–20 × 106 CD4+ T-cells are usually obtained from a single spleen. The num-bers of memory and naïve cells that can be obtained from a single spleen vary
depending on the age of the mouse (see previous paragraph); experiments
involving cell types present only at low frequencies, such as memory cells from
young mice, may require pooling spleens of mice of the same age in individual
1.2. Stimulation of T-Cells
with Monoclonal Antibodies to Cell Determinants
Because the number of cells obtained from a single spleen is on the order of
5–20 × 106 depending on the subset isolated, it is necessary to develop experi-mental conditions that permit the stimulation and assay of samples as small as
2–5 × 106 cells. Therefore, choosing appropriate stimuli as well as optimizing
stimulation conditions will influence the level of activation of the target enzyme
and are important factors in developing reliable assays for aging studies.
Activation of T-cells can be achieved through the use of lectins, such as
concanvalin A or phytohemagglutinin, that bind to unknown cell surface deter-minants. T-cells can also be stimulated using monoclonal antibodies specific for
components of the TCR and for other cell surface markers (e.g., CD4 or CD28),
or stimulated using pharmacological agents such as phorbol esters and calcium
ionophores. We describe here the use of monoclonal antibodies to the ε-chain of
the TCR/CD3 complex and CD4 to stimulate CD4+ T-cells isolated from young
and old mice. We feel that this provides a physiologically relevant stimulation
for a polyclonal population of T-cells. Studies using T-cells from transgenic
Raf-1 Protein Kinase Activity 87
mice that express a rearranged antigen-specific TCR will, in future work, pro-vide analogous information about responses to peptides presented by accessory
cells, which may more closely mimic the in vivo activation of T-cells.
The T-cells are incubated on ice with the stimulating antibodies followed by
crosslinking at 37°C with anti-rat IgG to initiate activation. We use purified
ascites fluid as our source of stimulating antibodies, although commercially
available monoclonals can also be used. We have found that anti-rat IgG can
effectively crosslink anti-CD3ε, even when using hamster clone 145–2C11 as
the anti-CD3 reagent. Furthermore, anti-rat IgG has the advantage of being
able to crosslink the rat antibodies used as costimulators (such as anti-CD4).
The phorbol ester phorbol myristate acetate (PMA) (10 ng/mL) activates Raf-1
in mouse T-cells, and can be used as a positive control. The commonly used
human lymphoma cell line Jurkat will also activate Raf-1 in response to either
PMA or to anti-CD3 antibodies (7), although it should be noted that activation
of human peripheral blood lymphocytes (PBL) or Jurkat cells with anti-CD3ε
does not require the addition of a crosslinking reagent such as anti-rat IgG.
Antibodies to other cell surface markers have been shown either to augment
TCR-driven T-cell activation or to activate signaling pathways independently
(8,9). In the case of Raf-1 we have evidence that the costimulatory molecule
CD28 plays a role in Raf-1 activation of CD4+ T-cells (21) while others have
shown that CD38 ligation can activate Raf-1 in Jurkat  (10). Little is known
about whether aging affects signal transduction from these other cell surface
receptors, although our own data suggest that Raf-1 induction by anti-CD28
antibodies in CD4 cells is unaffected by aging in mice.
When stimulating T-cells with antibodies to the TCR or other surface mol-ecules, it is important to determine how time of stimulation and dose of anti-bodies affect activation of the target enzyme. We found that in splenic CD4+
T-cells, Raf-1 activation occurs within 1 min of crosslinking anti-CD3ε and
anti-CD4 (Fig. 1). Initial descriptive studies of aging effects should include
data on the kinetics of activation in both old and young samples, to see if appar-ent differences in the level of activation represent merely an alteration in the
time course for activation. Furthermore, concentration curves of the antibody
or antibodies used must be determined to find the optimal stimulation condi-tions. We titrated the amounts of anti-CD3ε and anti-CD4, which were purified
from ascites fluid, to determine the optimal stimulation conditions for Raf-1
activation in CD4+ T-cells. We found that anti-CD3 concentrations between
0.2 and 7 µg/mL could effectively stimulate Raf-1 in CD4+ T-cells. In our hands,
addition of anti-CD4 at concentrations between 2 and 8  µg/mL augments
stimulation by anti-CD3, while anti-CD4 costimulation with amounts above
or below this range inhibits Raf-1 activation by anti-CD3. Furthermore,
anti-CD4 antibodies by themselves could not stimulate Raf-1 (21).
88 Kirk and Miller
1.3. Measurement of Raf-1 Kinase Activity
Raf-1 is a serine/threonine kinase of 74 kDa that is activated by association
with activated p21Ras. Besides its association with Ras, Raf-1 activation also
Fig. 1. (A) Analyzing Raf-1 kinase activity in CD4+ T cells from young, middle-aged, and old mice. Raf-1 was immunoprecipitated from 3  × 106 CD4+ T cells that
were stimulated with crosslinked anti-CD3+ anti-CD4 for the indicated times or
crosslinked anti-DNP for 2 min (lanes marked C) and incubated with a KIMEK sub-strate and [32P]ATP in an in vitro kinase assay. Reaction products were resolved by
10% SDS-PAGE and visualized by a PhosphorImager.  Arrow  indicates the 50-kDa
migrating band of KIMEK. (B) Data from several experiments (N = 5 for 1 and 2 min
and N = 4 for 5 and 10 min) are presented as -fold increase in Raf-1 kinase activity
over anti-DNP-stimulated samples. *p < 0.05 for Young vs Old and Young vs Middle-aged by one-way ANOVA followed by a Student–Newman–Keul’s post hoc test.
Raf-1 Protein Kinase Activity 89
requires either tyrosine or serine/threonine phosphorylation (11). The impor-tance of Raf-1 activity in T-cell activation and interleukin-2 (IL-2) production
has been shown using Jurkat cells transfected with dominant negative forms of
Raf-1. In this system, inhibition of Raf-1 blocks IL-2 production through the
TCR (12,13). It has also been shown in Jurkat cells that Raf-1 activation
through the TCR involves a protein kinase C dependent step that may be unique
to T-cells (14).
As the first kinase activated in the MAPK pathway, Raf-1 phosphorylates
and activates the dual specificity kinases MEK1/2, which, in turn, activate
ERK1/2 (11). Early studies of Raf-1 in vitro kinase activity utilized either non-specific substrates, such as histone H1, or Raf-1 autophosphorylation (11,15).
Other measures of Raf-1 activity involved analyzing the retardation in mobility
of Raf-1 in sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), presumably due to activating phosphorylation. However, these assays
lack the specificity of analyzing in vitro phosphorylation of a substrate known
to be phosphorylated by Raf-1 in vivo. Furthermore, as Raf-1 contains multiple
sites of phosphorylation, including those that inhibit kinase function, the slowly
migrating form of Raf-1 in SDS-PAGE may contain a collection of differen-tially phosphorylated species of varying levels of activity (11).
With the cloning of MEK into plasmids that allow for protein production
and purification from  E. coli, a suitable substrate for in vitro Raf-1 kinase
assays is available  (16). We used a plasmid encoding a MEK construct that
contains a lysine to methionine substitution at residue 97 in the ATP binding
site, and therefore lacks kinase activity  (16). This kinase-inactivated MEK,
which we call KIMEK, has been used as a substrate to determine the enzymatic
activity of Raf-1 in several cell systems, including Jurkat  (7). In our hands,
plasmid-derived KIMEK contains multiple species that migrate closely to the
predicted molecular weight for KIMEK of 50 kDa (Fig. 1), each of which
is phosphorylated by Raf-1 in in vitro kinase assays. These bands are likely to
represent degradation products of the KIMEK, and have been noted in other
publications using this substrate (16,17). Another version of KIMEK is com-mercially available as a GST fusion protein from Upstate Biotechnology (Lake
Placid, NY, USA).
Raf-1 function is more difficult to quantify in freshly isolated lymphocytes
than in cell lines, in part because Raf-1 protein levels are lower in primary cell
isolates. T-cells isolated from spleens are in a quiescent state and contain a
small cytosolic compartment as compared to a T-cell clone or Jurkat cell, which
are the size of T-cell blasts. In fact, Raf-1 protein levels are approximately
tenfold lower per cell in quiescent T-cells than in T-cell blasts  (18), and we
have found that Raf-1 protein expression in Jurkat is approximately fivefold
greater than in splenic CD4+ T-cells (Kirk and Miller, unpublished results).
90 Kirk and Miller
Furthermore, limitations in the number of T-cells that can be obtained from
each donor make it critical to develop and validate methods to measure kinase
activity from the lysates of 2–5 × 106 cells.
The in vitro kinase assay for Raf-1 involves the immunoprecipitation of
Raf-1 from cell lysates followed by incubation with the KIMEK substrate and
[32P]ATP. Several precautions in Raf-1 isolation and kinase measurement help
to ensure high levels of kinase activity, low levels of background phosphoryla-tion, and maximal difference between age groups. Anti-Raf-1 antibodies can
currently be purchased in either mouse monoclonal or rabbit polyclonal forms.
Although we have achieved the best results with rabbit polyclonal sera, other
forms should be tested in different systems (e.g., using human cells, or differ-ent strains of mice or cell types). Increasing the stringency of a lysis buffer will
reduce coprecipitation of other kinases, which might otherwise increase the
background phosphorylation of bands other than KIMEK. Background phos-phorylation can also be reduced by preclearing the lysates with Protein G–Seph-arose prior to immunoprecipitation.
2. Materials
1. Mice. We use specific-pathogen free male (BALB/c  × C57BL/6)F1 mice pur-chased from the National Institute on Aging contract colonies at the Charles River
Laboratories (Kingston, NJ, USA). Initial work should use animals of several
different ages, avoiding the use of animals so old that they are likely to be ill.
In our own studies we typically use mice aged approx 3–6 mo, 12–14 mo, and
18–22 mo from strains in which 50% mortality is not reached until 24–26 mo of
age. Mice over the age of 15 mo should be examined at necropsy and those with
skin lesions, splenomegaly, or macroscopically visible tumors should not be used.
2. Hanks balanced salt solution is supplemented with 0.2% bovine serum albumin
(BSA) (HBSS–BSA). It should be used and stored at 4°C unless otherwise noted.
3. Lympholyte-M™ (Cedarlane Laboratory, Ontario, Canada).
4. Antibodies: Antibodies to CD3ε (clone 145-2C11), the dintrophenyl hapten
(DNP) (clone UC8), CD4 (clone GK1.5), and CD8 (clone 53.6-7) were produced
in our laboratory from cell lines purchased from the American Type Culture Col-lection (Rockville, MD, USA). Anti-CD62L (clone Mel-14) can be purchased
from Pharmingen (San Diego, CA, USA). Anti-rat IgG used in cell stimulation
was purchased from Sigma (St. Louis, MO, USA). We find that most lots of goat
anti-rat IgG are suitable for crosslinking hamster antibodies (such as anti-CD3ε),
and are often superior to anti-hamster antibodies in this system; anti-rat Ig sec-ondary antibodies also have the advantage that they provide crosslinking between
hamster anti-CD3 and rat anti-CD4 or anti-CD8 antibodies. Magnetic beads conju-gated to anti-rat IgG are obtained from PerSeptive Diagnostics, Cambridge, MA,
USA, and polyclonal anti-Raf-1 from Santa Cruz Biotech, Santa Cruz, CA, USA.
5. Preparation of anti-Ig-coated dishes for B-cell depletion. Plates of 150 mm diam-eter are incubated with 35 mL of phosphate-buffered saline (PBS) containing
Raf-1 Protein Kinase Activity 91
1 µg/mL of rabbit anti-mouse IgG (H + L) overnight at 4°C on a level surface.
Plates can be stored for 4–7 d at 4°C or kept for long-term storage at –20°C. Prior
to use the plates are washed 3× with PBS to remove unbound antibody and incu-bated for at least 1 h with 30 mL of HBSS–BSA at 4°C. Petri dishes of 100 mm
diameter can be prepared in the same way using one-third the volume of rabbit
anti-mouse IgG.
6. RPMI supplemented with 10% FCS (RPMI–FCS).
7. Protein G–Sepharose (Pharmacia, Piscataway, NJ, USA)
8. RIPA buffer: 10 mM sodium phosphate, pH 7.0; 150 mM NaCl, 2 mM EDTA, 1%
sodium deoxycholate, 1% Nonidet P-40 (NP-40), 0.1% sodium dodecyl sulfate
(SDS); 0.1% 2-mercaptoethanol, 50 mM NaF, 200  µM Na3VO4, 2  µg/mL of
aprotinin; 0.5  µg/mL of leupeptin; and 2  µg/mL of pepstatin. Add 0.1 mM of
phenylmethyl sulfonyl flourite (PMSF) just prior to use.
9. PAN buffer: 10 mM PIPES, pH 7.0, 100 mM NaCl, 20 µg/mL of aprotinin.
10. Recombinant kinase inactive MEK substrate (KIMEK). We used a plasmid encod-ing a KIMEK containing a 6 histidine tag (the generous gift of Gary L. Johnson,
National Jewish Center for Immunology, Denver, CO, USA), that was expressed
and purified as described (19). Commercially available KIMEK in the form of a
glutathione-S-transferase (GST) fusion protein is available from Upstate Biotech-nology (Lake Placid, NY, USA, cat. no. 14–159).
12. Kinase buffer: 20 mM PIPES, pH 7.0, 75 mM NaCl, 10 mM MnCl2, 20 µg/mL of
13. Nitrocellulose (Schleicher and Shuell, Keene, NH, USA).
3. Methods
3.1. Purification of Splenic CD4+ T-Lymphocytes
3.1.1. T-lymphocyte Purification
1. Kill mice by cervical dislocation or CO2 asphyxiation. Experiments should be
age balanced, with mice of different age groups killed in each experiment.
2. Wet down fur with 70% ethanol and remove spleens to separate Petri dishes
(6 mm diameter) containing 6 mL of HBSS–BSA at room temperature.
3. Rub spleens between frosted glass slides to obtain a single cell suspension.
4. Pass suspension through a sheet of nylon mesh to remove cell clumps and con-nective tissue. Wash plate once with 6 mL of HBSS–BSA and pass through the
nylon mesh sheet. Each splenic suspension should be in a volume of 12 mL.
5. Centrifuge splenocytes at ~20°C for 5 min at 500g.
6. To remove erythrocytes, resuspend splenocytes in 8 mL of HBSS–BSA warmed
to 37°C. Carefully overlay splenocyte suspension on 4 mL of Lympholyte-M™
that has also been warmed to 37°C. Four milliliters of Lympholyte-M™ has the
capacity for 100 × 106 lymphocytes (the approximate number of lymphocytes in
a single spleen). Centrifuge at room temperature for 15 min at 600g with the
brake off.
7. Remove buffy coat containing lymphocytes and wash with 15 mL of HBSS–
BSA. Centrifuge at 4°C for 5 min at 500g. If the Lympholyte-M™ has not been
92 Kirk and Miller
sufficiently diluted there will be incomplete cell pelleting. Dilute further in
HBSS–BSA if supernatant shows any turbidity following centrifugation.
8. Resuspend lymphocytes in 15 mL of ice-cold HBSS–BSA and add to anti-Ig-coated dishes. Incubate at 4°C on a level surface for 40 min. Swirl plates once at
the end of the first 20 min to redistribute the cells. Dishes of 150 mm diameter
should receive approx 120–150 × 106 erythrocyte-depleted spleen cells; using more
cells per dish diminishes cell purity, while using fewer cells diminishes cell yield.
9. Gently separate nonadherent from adherent cells by swirling the plate vigorously
and then slowly removing the cell suspension from the tilted dish. It is particu-larly helpful to remove the last 0.5–1 mL very slowly, allowing the meniscus to
collect at the low point of the dish. Wash plate once with 5 mL of ice-cold HBSS–
BSA to improve cell yield, again taking care to remove the suspension slowly.
Centrifuge cells at 4°C for 5 min at 500g.
10. Resuspend cells in ice-cold HBSS–BSA and count. From 25 to 30 × 106 T-cells
can be obtained from a single spleen. Remove an aliquot of cells to perform flow
cytometry with an antibody to mouse CD3ε to determine the proportion of T-cells
in the resulting population (typically 85–90%).
3.1.2. Negative selection of T cell subsets
1. Dilute T-cells, prepared as described in Subheading 3.1.1., into HBSS–BSA to a
concentration of approx 5 × 106 cells/mL.
2. Add antibodies to cell markers of subsets to be depleted (i.e., for CD4+ T-cell
purification add antibodies to CD8). The amount of antibody needed is based on
trial experiments in which CD8 cell contamination of resulting cells is deter-mined by flow cytometric analysis. In our experiments, we use 1 µg of antibody
for 107 cells at a concentration of 5 × 106 cells/mL.
3. Incubate on ice for 20 min with gentle shaking at 5- to 10-min intervals.
4. Prepare anti-rat IgG-coated beads according to the manufacturer’s specifications.
Prepare enough for a bead-to-cell ratio of 50:1. Resuspend beads in 0.4 mL of
ice-cold HBSS–BSA.
5. Centrifuge cells at 4°C for 5 min at 500g. Wash cells once in 10 mL of cold
6. Resuspend cells in 1 mL of cold HBSS–BSA and add 0.4 mL of bead solution.
7. Incubate on ice for 20 min with shaking every 5 min to ensure suspension of
8. Fill tube to 5 mL with cold HBSS–BSA and pass under magnetic field (PerSeptive
Diagnostics, Cambridge, MA, USA), which is kept at 4°C.
9. Carefully remove cell suspension without disturbing the beads aligned along the
side of tube.
10. Repeat steps 8 and 9 two to three more times to ensure complete removal of beads.
11. Centrifuge cells at 4°C for 5 min at 500g. Resuspend in cold RPMI supplemented
with 10% fetal calf serum (FCS) (RPMI–FCS) and count. An aliquot of ~0.5 × 106
cells should be removed at this stage for flow cytometric analysis to assess the
purity of selected cells.
Raf-1 Protein Kinase Activity 93
3.2. T-lymphocyte Activation
1. Aliquot equal numbers of cells into 1.5-mL tubes in a total volume of 1 mL. A
sample from each mouse should be saved for analysis of Raf-1 protein expression
levels. In our experience we get usable results from samples as small as 2 × 106
cells/sample, although samples of 5 × 106 cells/sample give stronger signals and
are thus preferable when cells are not limiting.
2. Incubate cells with antibodies to the TCR/CD4 complex (anti-CD3ε and anti-CD4)
or other cell surface determinants. Some aliquots should also be incubated, as a
control, with a nonstimulatory antibody such as monoclonal anti-DNP of the same
species and isotype as the antibodies used for cell activation. Incubate the samples
on ice for 30 min. Invert the tubes several times during the incubation period.
3. Prepare solutions of RPMI–FCS with 5  µg/mL of anti-rat IgG (crosslinking
reagent) and RPMI–FCS with PMA (10 ng/mL). Warm to to 37°C.
4. Centrifuge cells at 1000g for 30 s to pellet.
5. Resuspend in 1 mL of warm anti-rat Ig solution or PMA and incubate at 37°C for
the desired interval. Raf-1 activation in mouse CD4 T-cells stimulated by anti-CD3 and anti-CD4 is detectable within 1 min and typically reaches a plateau
level within 5–10 min before subsiding to lower levels over the next half hour.
Stimulation with PMA for 5 min serves as a positive control for Raf-1 activation.
6. Stop reaction by centrifugation at 10,000g for 10 s.
7. Wash cells in ice-cold PBS. Centrifuge cells at 10,000g for 10 s.
8. Lyse cells in RIPA buffer, using 0.8 mL for immunoprecipitation or 40  µL for
protein expression levels.
9. Incubate in RIPA for 15–30 min. Centrifuge cells at 4°C for 10 min at 13,000g.
10. Remove pellet or transfer supernatants to a new tube.
3.3. In Vitro Kinase Assay of Raf-1
3.3.1. Raf-1 Immunoprecipitation
1. Preclear samples with 20 µL of Protein G–Sepharose (50% slurry) for 0.5–1 h on
a rocker at 4°C. Wash the Protein G–Sepharose several times with lysis buffer
before use. Pellet beads and transfer supernatants to new tubes.
2. Add anti-Raf-1 antibodies at 1:100 dilution (i.e., 8 µL for 0.8 mL of lysate) and
25 µL of Protein G–Sepharose. Incubate overnight on a rocker at 4°C.
3. Wash samples twice with ice-cold RIPA using centrifugation at 10,000g for 10 s.
4. Wash twice with ice-cold PAN buffer containing 1% NP-40 and once in ice-cold
PAN buffer.
3.3.2. Raf-1 Kinase Assay
1. For each sample add 50–100 ng of purified KIMEK and 30 µCi of [γ-32]P-ATP to
30 µL of kinase buffer. The kinase buffer should be prepared for all samples in
the same tube to ensure equal loading of KIMEK and [32P]ATP to each sample.
2. Add 30 µL to each sample and incubate for 15 min at 30°C.
3. Stop reaction by addition of 15 µL of 4× reducing sample buffer and boiling.
94 Kirk and Miller
4. Run samples on 10% SDS-PAGE and transfer to nitrocellulose. Expose blot to
X-ray film or an intensifier screen for phosphor storage image analysis.
5. Densitometric values are corrected for background levels and can be expressed as
fold activation by expression of the ratio of the densitometric volumes in the
stimulated samples to the anti-DNP-stimulated sample.
6. In a separate 10% SDS-PAGE gel transferred to nitrocellulose, the whole cell
lysates should be analyzed by Western blot analysis using the anti-Raf-1 antibod-ies according to the manufacturer’s instructions.
4. Notes
1. Because of limitations in the yield of T-cell subsets from mouse spleens, it is impor-tant to minimize cell loss during the purification procedures. To that end, careful
notice should be paid to the overlay of splenocytes on Lympholyte-M™ and the
complete removal of the buffy coat following centrifugation. It is also important to
ensure complete removal of nonadherent cells from the T plates. The plates can
placed under a microscope to determine if nonadherent cells remain. Further washes
of the anti-Ig-coated dishes may be helpful. Lastly, it is important to be sure cells
are in a single cell suspension, that is, not clumped, prior to the addition of the
anti-rat-Ig-coated magnetic beads to minimize cell loss at that step.
2. The level of Raf-1 kinase activation by antibodies to cell determinants is depen-dent upon antibody dose and time of stimulation. Dose curves for stimulating
antibodies will be necessary, as ascites purification and antibody storage will
give varying levels stimulating efficacy.
3. The stringency (i.e., detergent type) of the lysis buffer will determine the amount
of nonspecific phosphorylation of bands other than the KIMEK substrate. We use
RIPA because it has been shown to bring down Raf-1 in the absence of associated
proteins such as 14-3-3 (20). Because we find the purity of our KIMEK is <75%,
we have tried to limit the amount of coprecipitating kinases that may phosphory-late nonspecific proteins in our substrate preparation. However, less stringent
buffers can be used if the purity of the KIMEK is high (i.e., commercially pro-duced GST-KIMEK, which is more pure than the 6 histidine tagged KIMEK).
4. It is not necessary to determine protein concentration in cell lysates prior to Raf-1
immunoprecipitation. Assays should be performed using equal cell numbers in
each sample. Because contamination by erythrocytes may differ from young and
old mice, the total amount of protein is likely to be higher in T-cells from old
mice, even though the same number of cells is present in each sample.
5. Because the amount of Raf-1 present in 2–5 × 106 freshly isolated T-cells is quite
low, it is important to reduce the signal-to-noise ratio in the kinase assay. We
transfer the kinase samples from the SDS-PAGE to nitrocellulose before autora-diography to remove free [32P]ATP that remains in the gel.
This research was supported by Research Grant AG09801 and by training
Grants AI-07413 and GM-07315 at the University of Michigan.
Raf-1 Protein Kinase Activity 95
1. Wange, R. L. and Samelson, L. E. (1996) Complex complexes: signaling at the
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R. N. (1995) Zeta phosphorylation without ZAP-70 activation induced by TCR
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Chem. 271, 27,564–27,568.
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ligation results in activation of the Raf-1/mitogen-activated protein kinase and the
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The ins and outs of Raf kinases. Trends Biochem. Sci. 19, 474–480.
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T. D. (1993) Raf-1 is required for T cell IL2 production. EMBO J. 12, 4367–4373.
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Med. 180, 401–406.
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The MEK kinase activity of the catalytic domain of RAF–1 is regulated indepen-dently of Ras binding in T cells. J. Biol. Chem. 270, 5594–5599.
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Differentially Expressed Genes 97
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Identification of Differentially Expressed Genes
in Young and Senescent T Cells
Andrea Engel, Mahdi Adibzadeh, and Graham Pawelec
1. Introduction
The need to analyze changes in gene expression patterns occuring in senes-cent cells has stimulated the search for proper methods to identify the actual
differences between young and senescent cells. In studies of aging, differential
display reverse transcriptase-polymerase chain reaction (DDRT-PCR) has
already been applied successfully by several groups. Altered gene expression
in young and senescent fibroblast cultures (1), human mammary epithelial cells
(2), and rat brain cells (3,4) has been detected with this technique.
In 1992, two different but related strategies were introduced for fingerprint-ing expressed RNAs as cDNA tags. One was called differential display (DD),
(5) or, according to the PCR nomenclature, DDRT-PCR (6,7). The other was
named RNA arbitrary primed PCR (RAP-PCR) (8–10). Both variants of RNA
fingerprinting are able to detect differences in gene expression of a certain
percentage of expressed genes. Under the appropriate conditions, the pattern of
fragments derived from one type of cells is reproducible and can be compared
with that of another cell type. In both methods, primers of arbitrary sequence
are used to generate cDNA tags. This already suggests that in principle any
primer could be used to amplify differentially expressed genes.
The first method, DDRT-PCR, uses combinations of 10-mer arbitrary prim-ers with anchored cDNA primers and generates fragments that originate mostly
from the poly(A) tail and extend about 50–600 nucleotides upstream (5–7,11).
The original differential display (DDRT-PCR) technique (5,6) should be able
to generate a largely complete pattern of all mRNAs expressed in a particular
cell using a reasonable number of primer pairs (6). In this technique, cDNA is
amplified in a PCR with low-stringency annealing (at 42°C) throughout. The
98 Engel, Adibzadeh, and Pawelec
PCR primers consist of 10-base arbitrary primer and the same locking primer
used in the first strand synthesis (T11VN; V can be A, G, or C; N can be any of
the four nucleotides). The process of the DDRT-PCR is the following:
1. During the first strand synthesis, the locking primer samples a sizeable portion of
the mRNA (potentially 1/16th) present in the cell.
2. The arbitrary primer permits amplification of reverse-transcribed cDNA species
using the locking primer.
3. Repeated PCR cycling with low-stringency results in amplification of products
of a size <500 basepairs that can be evaluated on standard acrylamide gels.
The second technology, termed RNA arbitrary primed PCR (RAP-PCR),
was developed by M. McClelland and J. Welsh (8,10). With this technique, an
18-base arbitrary primer is used in both the first and cDNA synthesis reaction
and PCR. In the PCR only one primer is used to amplify the cDNA. The PCR
consists of a single round of low-stringency amplification (at annealing tem-perature of 37–42°C) followed by multiple rounds with higher stringency (at
annealing temperature of >50°C). By structuring the PCR in this manner, the
amplification of genomic DNA (gDNA) should be minimized because there
should be a limited chance of incorporating the primer used in the first strand
synthesis into both ends of complementary portions of gDNA. In contrast, the
18-base arbitrary primer is incorporated into one end of the cDNA as a result of
the reverse transcription of the RNA and into the other end during the single
round of low-stringency PCR. Consequently, the authors conclude that the sig-nal should be derived from RNA, not from gDNA. The process of the RAP-PCR is the following:
1. During the first strand synthesis, the arbitrary primer anneals and extends from
sites contained within the mRNA.
2. Second strand synthesis proceeds in a similar manner during the single round of
low-stringency PCR.
3. PCR amplification at high stringency proceeds strongly, having incorporated the
arbitrary primer into both ends of the cDNA.
4. A template-dependent competition exists that determines which potential PCR
products will ultimately predominate. PCR products can be analyzed by
acrylamide gel electrophoresis.
Any primer of 18 bases can be used in this technique and the PCR products
can be larger than 500 basepairs.
Here we present a detailed protocol that uses a poly(T) primer in the reverse
transcriptase reaction and only one 18-base primer in the arbitrary amplifica-tion of cDNA. The advantage of using the T12AG primer in the reverse tran-scription is the amplification of mRNA throughout total RNA. Previous
experiments have shown that the RAP-PCR principle with one primer in the
Differentially Expressed Genes 99
random amplification reaction can be used as successfully as if the cDNA were
done with the random primer of the following PCR. This method is applicable to
any set of two or more eukaryotic cell types and will result in reproducible
patterns of PCR products that correspond to expressed genes. Interesting bands
are excised from the gel, reamplified, cloned, and sequenced. The differential
expression has to be confirmed by Northern blotting, Southern blotting, or
nuclear run-on analysis.
2. Materials
2.2 DNA Agarose Gel Electrophoresis
1. 10× TBE: 1 M Tris, 0.89 M boric acid, 20 mM EDTA. Store at room temperature.
2. DNA sample buffer: 0.5% bromophenol blue, 0.5% xylene cyanol, 5% saccha-rose. Freeze until used in portions of 1 mL, then store at 4°C.
2.2. RNA Gel Electrophoresis
1. 10× 4-morpholine propanesulfonic acid (MOPS): 200 mM MOPS, pH 7.0; 50 mM
sodium acetate, pH 4.8; 10 mM EDTA, pH 8.0. Store at room temperature.
2. RNA sample buffer: 17 µL 10× MOPS, 84 µL of formamide, 29 µL of formalde-hyde (37%). Prepare fresh prior to use.
2.3. DNA Acrylamide Gel Electrophoresis
1. Sample buffer: 13 mL of double-distilled H2O (ddH2O), 13 mL of rehydration
buffer (DELECT buffer system), 1% bromophenol blue, 1% xylene cyanol,
250 µL of 0.2 M EDTA-Na2. Store at 4°C.
2.4. Enzymes
1. Alkaline phosphatase (Boehringer Mannheim).
2. AmpliTaq DNA polymerase (Amersham, Braunschweig).
3. Klenow fragment DNA polymerase (Boehringer Mannheim).
4. Reverse transcriptase (Amersham, Braunschweig).
5. RNase (Boehringer, Mannheim).
6. RNasin (RNase inhibitor, Promega, Madison, WI, USA).
7. T4-DNA ligase (Boehringer Mannheim).
8. T4-Oligonucleotide kinase (New England Biolabs, Schwalbach).
9. Taq Long Plus DNA polymerase I (Stratagene, Heidelberg).
2.5. Reaction-Kits, Filters, and Gels
1. CleanGels, 15% (ETC, Kirchentellinsfurt).
2. DELECT Kit for DDRT-PCR electrophoresis (ETC, Kirchentellinsfurt).
3. Hyperbond N+ Nylon membrane (Amersham, Braunschweig, or any other nylon
4. Megaprime™ DNA Labeling Kit (Amersham, Braunschweig).
100 Engel, Adibzadeh, and Pawelec
5. QIAquick® Gel Extraction kit (Qiagen, Hilden).
6. Rapid DNA Ligation Kit (Boehringer Mannheim).
7. RAP-PCR Kit (Stratagene, Heidelberg).
8. RNeasy Mini Kit (Qiagen, Hilden).
2.6. Molecular Weight Markers
1. 1 kb DNA Ladder (Gibco-BRL, Eggenstein).
2. 100-basepair DNA Ladder (Gibco-BRL, Eggenstein).
2.7. Silver Staining
1. Fixation solution: 10% acetic acid or 15% EtOH/5% acetic acid. Prepare a fresh
solution prior to use.
2. Silver reacting solution: 0.1% AgNO3, 200 µL of formaldehyde (37%). Prepare a
fresh solution prior to use.
3. Developing solution: 2.5% Na2CO3, 200 µL of formaldehyde (37%), 200 µL of
Na-thiosulfate (2%). Prepare a fresh solution prior to use.
4. Stopping/desilver solution: 2.0% (w/v) Glycine. Prepare a fresh solution prior
to use.
5. Impregnating solution: 10% Glycerol. Prepare a fresh solution prior to use.
2.8. Cloning of DNA Fragments
1. 50 mM CaCl2. Store at 4°C.
2. 10 × Klenow buffer: 500 mM Tris-HCl, pH 7.5, 60 mM MgCl2, 10 mM dithio-threitol (DTT). Freeze until use, then store at 4°C.
2.9. Bacterial Growth Media
1. LB Medium: 10 g/L of bacto-tryptone, 10 g/L of NaCl, 5 g/L of glucose, 5 g/L of
yeast extract. Store at 4°C.
2. LB plate agar: LB medium, 15 g/L of agar. Store at 4°C, cover down.
2.10. Plasmid Isolation
1. Resuspension solution: 50 mM Tris-HCl, pH 7.5; 10 mM EDTA. Store at room
2. Lysis solution: 0.2 M NaOH, 1% sodium dodecyl sulfate (SDS). Store at room
3. Neutralization solution: 2.55 M K-acetate, pH 4.8. Store at room temperature.
2.11. Northern Blotting
1. 20× SSC: 3 M NaCl, 0.3 M sodium citrate, pH 7.0 (adjust with NaOH). Store at
room temperature.
2. Hybridization buffer: 1 mM EDTA, pH 8.0, 0.25 M Na2HPO4, pH 7.2, 7% SDS,
1% BSA. Can be stored at 4°C for a few days.
3. Washing buffer: 1 mM EDTA, pH 8.0, 20 mM Na2HPO4, pH 7.2, 1% SDS. Store
at room temperature.
Differentially Expressed Genes 101
2.12. General Equipment
1. Centrifuge.
2. Centrifuge tubes (15 mL, 50 mL).
3. Electrophoresis tank (for horizontal electrophoresis), gel casting and tray.
4. High-voltage power supply.
5. Incubator at 37°C with a shaker.
6. Centrifuge for microfuge tubes.
7. Multiphor II-chamber (Pharmacia) or any other gel electrophoresis system for
horizontal acrylamide gel electrophoresis.
8. PCR Cups Micro Amp
9. Pipet tips (0,1–10 µL, 10–200 µL, 100–1000 µL).
10. Thermocycler
11. Video-Printer Mitsubishi Copy Processor (Software Bioprint Version 5.04,
Fröbel, Lindau) or another gel documentation system.
12. Whatman 3MM filter paper.
3. Methods
3.2 RNA Preparation
Basically, RNA can be isolated with any kind of RNA preparation method.
We had the best results with the cytoplasmic RNA preparation protocol for the
RNeasy-Kit (Qiagen) following the protocols provided. The common CsCl2
isolating method (12), or total RNA isolation with TRIZOL™ from Biozol or
any other RNA isolating-method should also be useful.
The RNA should finally be diluted in RNase-free (diethylpyrocarbonate
[DEPC]-treated) water and can be stored at –80°C (see also Note 1).
3.2. Reverse Transcription of RNA
In this protocol the reverse transcription is done with a T12VN-Primer
(T12VN; V can be A, G, or C; N can be any of the four nucleotides) to amplify
different “RNA families” with a poly(A) tail, i.e., mRNA. The nucleotides VN
should allow the primer to bind at the end of the long poly(A) tail of mRNA
(see also Notes 1 and 2).
1. For the reverse transcriptase reaction, 200 ng of mRNA or 0.5 µg of total RNA
should be used. In the first step the RNA is denatured for 5 min at 65°C in a
volume of 6 µL. After a short centrifugation (30 s at full speed) the reagents listed
in Ta ble 1 are added.
2. To allow the T12VN primer to find its complementary sequence on the RNA, the
probe is incubated for 10 min at room temperature. After that the probe is incu-bated for 45 min at 42°C in a water bath for the reverse transcriptase reaction.
Lastly the enzyme should be denatured by heating the probe for 5 min at 95°C.
102 Engel, Adibzadeh, and Pawelec
3. After a short centrifugation step the cDNA can be stored for 2–4 wk at –20°C or
for a longer period at –80°C.
3.3. Random Amplification of cDNA
In this system, the first step of the amplification reaction is the binding of
the random primer at low stringency to one tail of the mRNA. The second
cycle, already at higher stringency, forms DNA fragments with primer binding
sites on both ends of the fragment. The amplification reaction is then started
(see also Note 3).
1. The PCR is done in a total volume of 50  µL with the components shown in
Table 2.
2. The PCR-cycles are described in Ta ble 3.
With this protocol, oligonucleotides of > 500 basepairs can be amplified.
The probes are analyzed by horizontal gel electrophoresis and are detected by
silver staining with a nonradioactive method. They also can be stored at –20°C.
3.4. Discontinuous Acrylamide DNA Electrophoresis
For DNA separation, CleanGels and the DELECT-buffer Kit can be used.
This system allows a size separation of the DNA in a native, nondenaturing
manner. The electrophoresis is done in a horizontal manner in the Multiphor II
chamber from Pharmacia. This is an easy and useful method to analyze the
PCR products. The gels and buffer systems are available from Pharmacia,
Heidelberg or ETC, Kirchentellinsfurt.
Different DNA electrophoresis gels are provided. For these applications 15%
gels with 36 slots and a probe volume of 10–15 µL have proven to be useful,
but other gel systems can also be used. Here we describe a native electrophore-sis in a discontinuous buffer system (DELECT) with 15% gels which are rec-ommended for the separation of small oligonucleotides. The DELECT buffer
Kit contains a rehydration buffer for dry gels and a cathode and an anode buffer
Table 1
Components for the Reverse Transcriptase
Reaction After Denaturation of the RNA
5 mM T12VN-Primer 2 µL
10 mM dNTPs 4 µL
5 × RT-buffer 4 µL
0.1 mM DTT 2 µL
RNasin 0.5 µL
Reverse transcriptase 1.5 µL
Differentially Expressed Genes 103
for electrophoresis. The following descriptions are always for whole gels of 36
slots. The gels can also be divided according to the number of probes, but the
volumes and running conditions then have to be adapted.
3.4.1 Sample Dilution
Four to six microliters of the sample should be diluted 1:1 in loading buffer
for acrylamide gel electrophoresis.
3.4.2. Rehydration of the Dry Gel
For rehydration of the gels special chambers are available, but any plain
chamber could be used.
1. Pipet 35 mL of rehydration solution in the chamber and lay the gel film — with
the gel surface facing down — into the buffer, avoiding air bubbles.
2. Shake the gel slowly for 90 min at room temperature.
3. Remove excess buffer with an absorbant paper (e.g., Whatman paper), dry the
sample wells, and wipe buffer off the gel surface with the edge of the filter paper
until you can hear a “squeaking.”
Table 3
PCR Conditions for the Random Amplification
Phase Temperature (°C) Time Cycles
94 5 min —
Denature 94 30 s
Anneal 36 30 s 1
Extend 72 1 min
Denature 94 30 s
Anneal 50 30 s 35
Extend 72 1 min
Cool 4 ∞ —
Table 2
Mixture for the Random Amplification of cDNA
10× Taq buffer 5 µL
25 mM MgCl2 2 µL
10 mM dNTPs 10 µL
5 U/mL of Taq Long Plus 0.2 µL
ddH2O20.8 µL
Primer (18 nM)2 µL
cDNA (ca. 0.25 ng/µL) 10 µL
104 Engel, Adibzadeh, and Pawelec
3.4.3 Application of the Gel and the Electrode Wicks
1. Switch on the thermostatic circulator of the Multiphor II chamber or of a compa-rable electrophoresis system; adjust to 21°C.
2. Lay two electrode wicks into the compartments of the paper pool, or any compa-rable chamber.
3. Apply 20 mL of the electrode buffer to each wick (for the anode wick use the
anode buffer and for the cathode wick the cathode buffer).
4. Apply a very thin layer of kerosine onto the cooling plate with a tissue paper, to
ensure good contact with the gel.
5. Place the gel (surface up) on the center of the cooling plate: the side containing
the wells must be oriented toward the cathode.
6. Place the cathodal strip onto the cathodal edge of the gel, and the anodal one onto
the other edge. For the DELECT buffer system the strips should be ca. 8 mm
away from the edges of the samples. Smooth out any air bubbles.
3.4.4. Sample Application and Gel Electrophoresis
1. Apply 10–15 µL of each sample to the sample wells. To determine the size of the
amplifed products it would be worthwhile in addition to apply a molecular weight
2. Clean palladium electrode wires with a wet tissue paper before each electro-phoresis run.
3. Move electrodes so that they will rest on the outer edges of the electrode wicks.
The running conditions for a whole 36-slot 15% gel are shown in Table 4. If gels
are divided the voltage should be kept constant and the mA and W should be
adjusted according to the gel size (e.g., the half for half-gels).
Depending on the fragment size, the second step has to be varied in length.
The larger the amplified oligonucleotides are, the longer this step has to run. The
xylene cyanol runs at a molecular range of ca. 100 basepairs.
4. After running, gels have to be fixed in 10% acetic acid or 15% EtOH/5% acetic
acid for at least 30 min. The fixation step can be prolonged overnight.
3.4.5. Silver Staining
1. After fixation, gels are washed 3  × 5 min in ddH2O. Then the silver staining
protocol shown in Ta ble 5 is performed (see also Note 4).
Gels can be stored at room temperature after they are shrink-wrapped into a
strong cling film.
Table 4
Running Conditions for a Whole 15% CleanGel
10 min 200 V 20 mA 10 W
60–120 min 375 V 30 mA 20 W
20–60 min 450 V 30 mA 20 W
Differentially Expressed Genes 105
3.5. Reamplification of the differentially
expressed oligonucleotides
1. The differentially expressed bands can be cut out of the acrylamide gel with a
scalpel and then reduced to small pieces with a pipet tip.
2. From 50 to 100 µL of sterile ddH2O is added and the probes then incubated for 30 min
at 65°C and overnight at room temperature to elute the DNA out of the gel.
3. For the reamplification reaction, 10  µL of a 1:10 dilution of the eluted oligo-nucleotides are used. The PCR mixture is the same as in the amplification reac-tion. The conditions are shown in Ta ble 6.
4. The PCR probes can be analyzed in a regular horizontal DNA electrophoresis
with 1.5–2% agarose gels and then detected with an DNA intercalating stain (e.g.,
ethidium bromide). They also can be stored at –20°C.
The reamplification reaction could amplify multimers of the original oligo-nucleotide or other bands, which are artefacts from the elution of the acrylamide
gel. For further cloning the fragments into a plasmid, only the interesting band
should be eluted from an agarose gel by a convenient method. The cloned frag-ments should further be sequenced, because one eluted band from the random
Table 6
PCR Conditions for the Reamplification of Differentially Expressed
Phase Temperature (°C) Time (min) No. of cycles
Denature 94 5 —
Denature 94 1
Anneal 55 2 35
Extent 72 1
Cool 4 ∞ —
Table 5
Silver Staining Protocol of a Whole 15% 36-Slot CleanGel
Silver reaction 20–30 min 200 mL 0.1% AgNO3 + 200 µL
Formaldehyde (37%)
Washing Thoroughly wash gel surface with ca. 250 mL of ddH2O
with a squeeze bottle
Developing 2–5 min 200 mL 2.5% Na2CO3 + 200 µL of
(visual control)   Formaldehyde (37%) +
200 µL of Na thiosulfate (2%)
Stop/desilver 10 min 250 mL 2.0% (g/v) glycine
Impregnate 10 min 250 mL 10% Glycerol
106 Engel, Adibzadeh, and Pawelec
amplification might contain some fragments of the same size, but with differ-ent sequences. This is based on the randomly amplified products in the PCR,
which can result in oligonucleotides coincidentally of the same size. The dif-ferential expression of the DNA fragments has to be confirmed, i.e., by North-ern blotting (see also Note 5).
3.6. DNA Gel Electrophoresis
1. Agarose gels are prepared with 1× TBE buffer and should have a percentage of
1.5–2.0% agarose.
2. DNA gel electrophoresis is done in a horizontal manner. The gels have to be
covered with running buffer (1× TBE). DNA probes are diluted 1:5 (v/v) in
sample buffer and loaded into the sample wells of the gels (depending on the
concentration and the volume of the gel wells, 10–40 µL of the DNA probe can
be taken). To determine the size of the products it would again be useful to apply
additionally a molecular weight marker. The orientation of the electrodes has to
be chosen so that the DNA can run in the direction of the anode (positively
charged electrode), because DNA is negatively charged.
The voltage should be set to 1–15 V/cm (distance between the electrodes). The
tracking dyes incorporated into the sample buffer serve as markers for the
progress of the run.
3. At the end, the DNA is stained with ethidium bromide (0.5 µg/mL) for 2–5 min in
a staining bath (Caution: wear gloves, because ethidium bromide is carcino-genic). After staining, the gels are briefly washed in ddH2O to remove the surplus
ethidium bromide (again wear gloves and collect ethidium bromide waste sepa-rately). The DNA bands can be detected in UV light and can be documented with
a suitable system (see also Note 7).
3.7 Purification of Nucleic Acids
3.7.1. Phenol/Chloroform Extraction
Phenol is also a carcinogen, so work under a fume hood. The extraction of
aqueous nucleic acids with phenol-chloroform allows the denaturing and
removal of proteins (eg., enzymes) from the probes.
1. PCR probes (45–40 µL, see also Notes) are diluted with ddH2O to a final volume
of 300–500 µL in a microfuge tube.
2. The same volume phenol/chloroform/isoamyl alcohol (25:24:1) is applied and
probes are mixed vigorously on a vortex mixer.
3. To separate the aqueous—DNA containing—and the nonaqueous phase the
probes are centrifuged at 12,000g for 2 min at room temperature.
4. The upper aqueous phase is transferred into a new microfuge tube and the same
volume chloroform/isoamyl alcohol (24:1) is added.
5. The probe is mixed and centrifuged again. The upper phase is carried over in a
new microfuge tube and the DNA is concentrated with the protocol for ethanol
Differentially Expressed Genes 107
3.7.2 Ethanol precipitation
This protocol can be employed should the sample require concentrating
and desalting.
1. To precipitate the DNA, the probe is treated with 1:10 vol 3 M NaAc, pH 4.8, and
2–3 vol of cold absolute ethanol. Then it is incubated at least 30 min at –20°C.
2. The precipitated nucleic acids are then centrifuged (12,000g, 15 min, 4°C), the
supernatant discarded, and the pellet dried at 40–50°C with open cover to remove
the surplus alcohol.
3. The DNA pellet can then be diluted in an adequate volume of ddH2O (to be suit-able for the cloning protocol take only 11.7 µL of ddH2O).
3.5.3 DNA Gel extraction
To elute DNA fragments from the agarose gels many kits are available from
several companies. One of the most convenient is the QIAquick Gel Extraction
kit (Qiagen, Hilden).
The band of interest can be cut out of the gel under a UV transilluminator
with a clean, sharp blade (wear glasses to protect your eyes). The fragment is
now transferred into a clean microfuge tube and the extraction can be performed
according to the protocol given.
3.8. Quantification by photometry
Absorption at 260 nm is measured in an appropriate photometer to determine
the concentration of nucleic acids. An absorption of 1 corresponds to 50 µg/mL
double-stranded DNA, 40  µg/mL single-stranded RNA, and 30  µg/mL
single-stranded oligonucleotides.
The purity of the probes is determined with the quotient of the OD (optical
density) at 260 nm:280 nm. This quotient should not be less than 1.8.
3.9. Cloning of PCR Fragments
3.9.1. Klenow Polymerase Treatment
The 5′:3′ polymerase activity and the 3′:5′ exonuclease activity of the Klenow
fragment of the polymerase I from E. coli is used to blunt the ends of the PCR
1. The DNA pellet from the ethanol purification is diluted in 11.7 µL of ddH2O. The
following reagents are then added:
1.5 µL of 10× Klenow-buffer
1.8 µL of Klenow enzyme (0.8 U/µg of DNA)
and the probe is incubated for 10 min at 37°C.
2. After that 1.2 µL of dNTPs (1.25 mM each nucleotide) are applied and a further
incubation step at 37°C for 30–35 min is performed.
3. Heating at 70°C for 10 min denatures the Klenow fragment.
108 Engel, Adibzadeh, and Pawelec
3.9.2. 5′-Phosphorylation of the Fragments
After the blunting of the fragments the 5′-ends have to be phosphorylated
before they can be cloned into a plasmid vector. The enzyme T4-polynucle-otide kinase (PNK) is employed for this purpose.
1. Add to the probe of 3.9.1:
2 µL of ATP (10 mM)
1 µL10 × PNK buffer
1 µL of PNK
(final volume: 20 µL)
An incubation step of 30 min at 37°C is performed.
2. At this stage the PNK can be inactivated for 10 min at 70°C and the probes can be
stored at –20°C.
3. Before cloning the probes have to be purified by agarose gel electrophoresis. The
fragments of interest can then be separated via the gel from additionally ampli-fied fragments and primer dimers from the reamplification reaction.
3.9.3. Preparation of the Plasmid Vector
1. The plasmid vector (e.g., pUC18, pUC19, Bluescript, or any other vector) is
digested with a blunt ending restriction enzyme that cuts the plasmid vector into
the multiple cloning site (MCS; i.e., SmaI) (see also Note 6).
The restriction conditions depend on the enzyme used and should be followed
according the product information. The volume should be between 20 and 50 µL
and 2 U of the enzyme per microgram DNA should be added. After the incuba-tion the enzyme has to be inactivated. It is mostly possible to heat inactivate the
restriction enzymes, but if this should not be the case (see the product informa-tion) an agarose gel purification or ethanol precipitation has to be done. The puri-fication of the vector from the restriction enzyme and the buffers is also necessary
prior to the following dephosphorylation reaction.
2. To avoid the religation of the vector in the ligation reaction, it has to be dephos-phorylated with an alkaline phosphatase. The dephosphorylation reaction is done
with 0.04 U alkaline phosphatase/µg DNA in 1× phosphatase buffer (provided by
the manufacturer) at 37°C for 60 min. After the incubation the vector has to be
purified in an agarose gel and eluted with a suitable method.
3.9.4. Ligation
For the ligation reaction there are many kits provided by several companies.
We have had good experience with the Rapid Ligation Kit (Boehringer, Mann-heim). This is a method for cloning blunt ended fragments into plasmid vec-tors. But of course any other ligation method, without using a kit, could be
successful (see also Notes 8 and 9).
A ligation procedure without using a kit requires the following reaction mix:
Differentially Expressed Genes 109
Dephosphorylated, restricted plasmid (50 ng)/phosphorylated, blunt ended frag-ment in a ratio 1:3 to 1:5
1× Ligase buffer
1 µL of T4-ligase (5 U/µL)
(total volume 20–40 µL)
The probe can then be incubated at 4°C overnight or at 16–20°C for 3–4 h.
The probes can be stored at –20°C.
3.9.5. Preparation of CaCl2 competent cells for transformation
The E. coli cells are treated following the CaCl2 procedure from Mandel
and Higa (13).
1. 50 mL of LB medium are inoculated with 5 mL of an overnight preculture of
the E. coli strain. The culture is then incubated at 37°C to an OD (600 nm) from
0.3 to 0.4.
2. The cells are centrifuged (2500g, 10 min, 4°C) and the pellet is resuspended in
100 mL of ice-cold 50 mM CaCl2 solution.
3. The cells are centrifuged again (only 1800g, 4°C).
4. The pellet is resuspended carefully in 20 mL of CaCl2 and incubated for 20 min
on ice.
5. The cells are centrifuged again (1800g, 4°C) and the pellet is then carefully resus-pended in 10 mL of ice-cold CaCl2 solution. In addition, glycerine is added to a
final concentration of 20%. The probes should be divided in aliquots of 200 µL in
microfuge tubes and then they can be stored at –80°C for several months.
3.9.6. Transformation
1. Prior to transformation, the ligation reaction mixture is filled with ddH2O to a
final volume of 100 µL.
2. This mixture is added to a 200-µL aliquot of not completely thawed competent
bacteria. The probe is then incubated 30 min on ice.
3. After that the bacteria are heat shocked for exactly 45 s at 42°C and 1 mL of 37°C
warm LB medium is added at once.
4. The bacteria are incubated for 30–40 min. (They should have the possibility to
divide once the plasmid incorporation is stabilized.)
5. Then aliquots of, e.g., 50, 100, and 200 µL are plated onto an LB agar plate with
appropriate antibiotics or other selection markers (e.g., β-galactosidase).
6. The plates are incubated at 37°C overnight and can be stored at 4°C for a few
days. Positive clones can be inoculated into LB medium with a suitable antibiotic
and after an overnight incubation at 37°C (shake) the plasmids can be isolated for
further control.
3.9.7. Plasmid Isolation by Alkaline Lysis
For plasmid isolation, many reaction kits are available that can result in
extremely pure plasmids. These are required, for example, for sequencing but
110 Engel, Adibzadeh, and Pawelec
for the control of plasmid transformation a simple protocol without using a kit,
as described here, is enough.
1. From 1 to 3 mL of an overnight bacteria culture, transformed with a plasmid
vector, are pelleted in a microfuge tube by centrifugation (full speed, 3 min).
2. The supernatant is discarded and the pellet is resuspended in 200  µL of cell
resuspension solution by pipetting.
3. Next, 200 µL of cell lysis solution are added and mixed by inverting. The solution
should clear almost immediately.
4. Then 200 µL of neutralization solution are added and the probes are again mixed
by inverting (not by pipetting).
5. Spin down at 12,000g for 10 min.
6. Transfer supernatant to new tube. Discard pellet.
7. Add 1/10 of 1 volume of 3 M sodium acetate, pH 7.0 and 2.5 volumes of ETOH.
8. The probes are centrifuged at full speed for 5 min and the supernatant is discarded.
9. The pellet should be washed in 300  µL of 70% ethanol and finally dried with
open cover at 40–50°C.
10. Then the pellet can be resuspended in 20–30 µL of ddH2O and stored at –20°C.
The ligation can be checked by a restriction digestion of the plasmid with
two enzymes that each cut at one end of the insertion site of the fragment in the
multiple cloning site of the plasmid vector. In a common agarose gel electro-phoresis the genuine positive plasmids can be detected.
3.10. Verification of the Differentially Amplified Fragments
in Northern Blot Analysis
3.10.1. RNA Gel Electrophoresis
with a Denaturating Formaldehyde Gel
1. To prepare a 1–1.5% formaldehyde gel add 1–1.5 g of agarose into 73 mL of
ddH2O. The agarose is dissolved by boiling and then cooled in a water bath to
50°C. The volume should be restored with ddH2O to 73 mL. Then 10 mL of 10×
MOPS buffer and 17 mL of formaldehyde (37%) are added and the solution is
mixed immediately and placed into a gel track. The fully polymerized gel can be
applied into a horizontal gel electrophoresis tray and covered with RNA running
buffer (1× MOPS buffer).
2. From 2 to 10 µg RNA is denatured 5 min at 65°C in the following mixture:
RNA in a final volume of 6 µL (diluted in RNase-free ddH2O)
12.5 µL of formamide (deionized)
2.5 µL of 10× MOPS
4 µL of formaldehyde (37%)
3. The probe is then chilled on ice and 1:10 sample buffer is added. The probes are
applied to the gel. The running conditions can be varied from 100 to 120 V for 3 h
or overnight at 20 V.
4. The RNA run can be documented after ethidium bromide staining as for DNA.
Differentially Expressed Genes 111
3.10.2. Northern Blotting
Here we describe a common method for Northern blotting that is per-formed with the ordinary capillary flow system. As RNA is electrophoresed
in the denaturing system the gel does not need any further denaturation or
The RNA is transferred from the gel onto a positively loaded nylon mem-brane in a chamber construction shown in Fig. 1.
The blotting setup is constructed in the following manner:
1. Set up the transfer support into the tray and place a wider glass plate onto the
support. Put two pieces of Whatman 3MM paper on the glass plate so that they
hang up into the tray. Fill the reservoir with transfer buffer (20× SSC) and wet
also the Whatman 3MM paper (avoid air bubbles).
2. Cut a piece of the nylon membrane of the same size as the gel (handle the filter at
the edges only!) and wet the filter for 5 min in the transfer buffer. Cut off the
lower right corner of the filter (so as not to lose orientation during the procedure).
3. Flood the two Whatman 3MM papers on the glass plate with transfer buffer and
place the gel onto the papers. Squeeze out air bubbles by rolling with an RNase-free pipet over the gel. Cut a piece of the size of the gel out of the middle of a
piece of common cling film and apply this around the gel. This will ensure that
the liquid flow from the reservoir is transferred through the gel.
4. Apply the nylon membrane carefully directly onto the gel and again squeeze out
air bubbles as described previously.
Fig. 1. Northern blotting setup.
112 Engel, Adibzadeh, and Pawelec
5. Wet two or three pieces of Whatman 3MM paper of the same size as the gel and
place them onto the membrane. Cut paper towels to the same size and put them
onto the Whatman papers to form a stack 7–8 cm high.
6. Apply a glass plate onto the top and a weight of 400–500 g onto it to ensure good
contact during the transfer.
7. The RNA is transferred overnight from the gel to the nylon membrane at room
8. After blotting, the RNA is fixed by incubating the membrane for 5 min onto a
Whatman paper that is soaked with 0.05 M NaOH (RNA site up). Before storing
the filter at 4°C in cling film it should be briefly washed with 2× SSC solution.
3.10.3. Probe Hybridization and Labeling
Here we describe a method which uses the standard megaprime protocol
from the Megaprime™ labeling kit (Amersham) (see also Notes 10 and 11).
1. From 2.5 to 25 ng of the DNA are dissolved in 10  µL of ddH2O. Twenty-five
nanograms of the template DNA are then applied into a microfuge tube and 5 µL
of the random primer are added.
2. The DNA is denatured at 95–100°C for 5 min in a boiling water bath.
3. The probe is collected by a short centrifugation step and then kept at room
4. To label the DNA template the following components are added:
Unlabeled dNTPs 4 µL each of dCTP, dTTP, dGTP; 2.5 mM
5 µL of reaction buffer
Radiolabeled dATP
5 µL of [32P]dATP (specific activity; 3000 Ci/mmol]
2 µL of DNA polymerase I (at –20°C until needed)
11 µL of ddH2O
5. The probe is mixed gently and incubated at 37°C for 10–30 min.
6. The reaction is stopped by heating the probe at 95°C for 5 min. The surplus
[32P]ATP can be removed on a Sephadex G50 column.
7. The probe can now be stored at –20°C for 2–3 d. Further storage should be
avoided because of the short half-life of 32P.
8. Prior to hybridization, the membrane is incubated for at least 15 min at 65°C or
overnight in a prehybridization box with prehybridization solution.
9. For the hybridization, the labeled DNA probe is denatured by heating to
95–100°C for 5 min, and then directly chilled on ice. The labeled DNA probe can
now be added into the prehybridization solution at a concentration of 1  × 106
cpm/mL and incubated at 55–65°C for at least 12 h (the higher the hybridization
temperature chosen, the higher the stringency of the hybridization).
10. After hybridization, the filters are washed by incubating for 5 min at 65°C.
This procedure is repeated if necessary and checked with a hand monitor for
radioactive detection (Geiger counter). The membrane is now wrapped in a nylon
filter (e.g., Saran Wrap) and the hybridization is detected by autoradiography.
Differentially Expressed Genes 113
4. Notes
1. Working with RNA demands some special precautions, as RNA is rapidly
degraded in the presence of the very stable enzyme RNase. The possibility of
contamination with RNase should be minimized. Therefore always wear gloves
when preparing or handling RNA probes. The laboratory equipment for RNA
procedures should be separated if possible or sterilized in an autoclave before
use. Equipment that cannot be autoclaved should be treated with 0.5 M NaOH or
70% ethanol. RNase can be inactivated with DEPC (CAUTION: carcinogen; use
a fume hood). DEPC should be added to every solution in a final concentration of
0.1% (but not to solutions that contain Tris) and then incubated overnight in a
fume hood with cover open, or if possible autoclaved for 20 min.
For RNA preparation, a method for isolating cytoplasmic RNA is useful, to
avoid amplification of incompletely spliced RNA in the reverse transcription.
Genomic DNA contamination is one of the major causes of false-positive
results. Although most kits guarantee preparation of DNA-free RNA, it would be
useful to check samples for possible DNA contaminations. This could be done by
an RT-PCR reaction with a primer to genes usually expressed that gives different
products with mRNA and gDNA (e.g., Ta ble 7).
In the RNA gel electrophoresis, the formamide that inhibits the RNases can be
reduced by half if the additional volume is needed.
2. It is useful to confirm the success of the reverse transcriptase reaction in a simple
common PCR reaction with primers to genes usually expressed in the material of
3. The use of too much cDNA in the amplification reaction, or in the reamplification,
can have a bad effect on the reaction. In the worst case, there may be no amplifi-cation of the DNA at all.
The various bands of the acrylamide gel should be cut out as small as possible,
to avoid contamination with neighboring bands.
4. Silver staining is a very sensitive method for oligonucleotide detection. To be
able to differentiate varying bands, the probe dilution in the acrylamide gel elec-trophoresis has to be optimal. Our experience has shown that 5–8 µL from the
amplification reaction are enough to detect clear bands.
The developing reaction in silver staining can happen very quickly. To avoid
the gel becoming too dark it is useful to prepare the stop/desilver solution before-Table 7
Primer Sequence of a β-actin gene that Gives Bands of Different Sizes
for cDNA and gDNA
Target Primer sequence (5′ → 3′) Product (bp)
β-Actin sense GGC GGC ACC ACC ATG TAC CCT 202 (312)
β-Actin antisense AGG GGC CGG ACT CGT CAT ACT
114 Engel, Adibzadeh, and Pawelec
hand, so that the gel can be transferred rapidly into the stop bath once the reaction
has progressed far enough.
5. Before cloning, 5–10  µL of the reamplified fragment should be checked in an
agarose gel and the DNA content should be measured. The other 40–45 µL can
then be treated with the chloroform phenol extraction protocol.
6. The enzyme content of the restriction probe should not be more than 1:10 of the
entire volume, because the enzymes are diluted in a very highly concentrated
glycerol solution that inhibits the enzyme activity.
7. Before running an agarose gel, incubate the isolated plasmid vector with 1 U of
RNase/20 µL of probe for 10 min at 37°C to remove RNA contamination.
8. Avoid too much pipetting after the ligase is added, because this enzyme is sheared
very easily and thereby loses its activity.
9. A satisfactory and very uncomplicated ligation kit is provided by Boehringer
Mannheim. The ligation protocol can be followed as described but the incubation
time can be extended to 30–45 min instead of the 5 min given in the manufac-turer’s directions for use.
10. The protocol that is described uses a laboratory equipped for radioactive meth-ods! It would be useful to prepare the radioactively labeled probes in microfuge
tubes that have a cover with a screw thread to minimize risk of aerosols upon
opening the tubes after heating.
The polymerase should be added last, because the reaction immediately starts
when the enzyme is applied. If one dNTP is missed, every amplification will stop
at the complementary nucleotide of the missing nucleotide.
11. The membrane should not be washed for too extended a period, because the
labeled probes would be washed away. After that the membrane can be hybrid-ized again with another probe. This process can be repeated up to 4–5 times.
1. Linskens, M. H., Feng, J., Andrews, W. H., Enlow, B. E., Saati, S. M., Tonkin, L. A.,
Funk, W. D., and Villeponteau, B. (1995) Cataloging altered gene expression in
young and senescent cells using enhanced differential display. Nucleic Acids Res.
25, 3244–3251.
2. Swisshelm, K., Ryan, K., Tsuchiya, K., and Sager, R. (1995) Enhanced expression
of an insulin growth factor-like binding protein (mac 25) in senescent human mam-mary epithelial cells and induced expression with retinoic acid. Proc. Natl. Acad.
Sci. USA 92, 4472–4476.
3. Salehi, M., Hodkins, M. A., Merry, B. J., and Goyns, M. H. (1996) Age-related
changes in gene expression in the brain revealed by differential display. Experientia
15, 888–891.
4. Wu, H. C. and Lee, E. H. (1997) Identification of a rat brain gene associated with
aging by PCR differential display method. J. Mol. Neurosci. 8, 13–18.
5. Liang, P. and Pardee, A. B. (1992) Differential display of eucaryotic messenger
RNA by means of the polymerase chain reaction. Science 257, 967–971.
Differentially Expressed Genes 115
6. Bauer, D., Müller, H., Reich, J., Riedel, H., Ahrenkiel, V., Warthoe, P., and Strauss,
M. (1993) Identification of differentially expressed mRNA species by an improved
display technique (DDRT-PCR) Nucleic Acids Res. 21, 4272–4280.
7. Liang, P., Bauer, D., Averboukh, L., Warthoe, P., Rohrwild, M., Müller, H., Strauss,
M., and Pardee, A. B. (1995) Analysis of altered gene expression by differential
display. Methods Enzymol. 254, 304–321.
8. Welsh, J., Chada, K., Datal, S. S., Cheng, R., Ralph, D., and McClelland, M. (1992)
Arbitrary primed PCR fingerprinting of RNA. Nucleic Acids Res. 20, 4965–4970.
9. Welsh, J., Rampino, N., McClelland, M., and Perucho, M. (1995) Nucleic acid
fingerprinting by PCR-based methods: applications to problems in aging and
mutagenesis. Mutat. Res. 338, 215–229.
10. McClelland, M. and Welsh, J. (1994) RNA fingerprinting by arbitrary primed PCR.
PCR Methods Appl. 4, 66–81.
11. Liang, P., Averboukh, L., and Pardee, A. B. (1994) Method of differential display.
Methods Mol. Genet. 5, 3–16.
12. Sambrook, F., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Labora-tory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor,
New York.
13. Mandel, M. and Higa, A. (1970) Calcium dependent bacteriophage DNA infection.
J. Mol. Biol. 53, 159–162.
14. Wei, Q., Xu, X., Cheng, L., Legerski, R. J., and Ali-Osman, F. (1995) Simultaneous
amplification of four DNA repair genes and β-actin in human lymphocytes by mul-tiplex reverse transcriptase PCR. Cancer Res. 55, 5025–5029.
Xenobiotic-Metabolizing Enzymes 119
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Xenobiotic-Metabolizing Enzyme Systems
and Aging
Christopher R. Barnett and Costas Ioannides
1. Introduction
The human body is continuously exposed to a wide array of structurally
diverse chemicals. Such exposure occurs even at the fetal stage as almost all
chemicals that are present in the mother’s blood can readily cross the placenta
and reach the fetus. Some of these chemicals are ingested voluntarily, for exam-ple, medicines and food additives, but the vast majority are taken involuntarily,
as environmental contaminants present in the air or in the occupational
environment. Undoubtedly, the most important source of such chemicals is the
diet, and many dietary constituents have been shown to induce many forms of
toxicity including cancer  (1). Exposure to chemicals is thus inevitable and
unavoidable. The body cannot exploit these chemicals either to generate energy
or transform them to building blocks and consequently its response is to rid
itself of their presence. This chapter discusses the role of drug-metabolizing
enzyme systems in this process and the effects of age. The measurement of
drug-metabolizing activities is of increasing importance in the safety evalua-tion of drugs in humans. This chapter describes the use of alkylphenoxazone
derivatives for investigating selected activities of drug-metabolizing enzymes.
Chemicals that reach the systemic circulation and are distributed throughout
the body are generally lipophilic, a property that allows them to traverse the
various cellular membranes. Such lipophilic chemicals are also difficult to
excrete through the kidney and bile. Consequently, to facilitate their elimina-tion, the body converts them to hydrophilic metabolites, which are much easier
to excrete. Furthermore, such metabolism terminates any biological activity
that may be manifested by these chemicals, as the products of metabolism are
biologically inactive, being unable to interact with the receptors for which the
120 Barnett and Ioannides
parent compounds often have high affinity. The metabolism of chemicals occurs
through an enzymic process that involves a number of enzyme systems present
in many tissues, the highest concentration being encountered in the liver. The
metabolism of chemicals is generally achieved in two phases. Phase I involves
primarily the incorporation of an atom of oxygen to the substrate, producing a
more hydrophilic, and in most cases biologically inactive, metabolite that can
now participate in Phase II metabolism. During Phase II metabolism, the newly
generated metabolite is conjugated with endogenous substrates, such as sulfate
and glucuronide, to form highly hydrophilic products that can now be very
readily eliminated.
1.1. Metabolic Activation of Chemicals
Although metabolism of chemicals is essentially a deactivation process,
with certain chemicals one or more metabolic pathways may lead to the gen-eration of reactive intermediates, a phenomenon known as metabolic activa-tion or bioactivation (Fig. 1). Because of their increased chemical reactivity,
Fig. 1. Metabolic activation of chemicals.
Xenobiotic-Metabolizing Enzymes 121
these intermediates interact covalently with vital cellular components, lead-ing to the manifestation of toxicity  (2,3). For example, most chemical car-cinogens are metabolized to reactive species that interact with DNA to induce
mutations that may lead to the formation of tumors. Generally, the activation
pathways are usually minor routes of metabolism and the body is well
equipped to deal with the limited amounts of reactive intermediates that are
produced. This defense mechanism proceeds through conjugation of the reac-tive intermediates with the endogenous tripeptide glutathione, the glutathione
conjugates being excreted in the urine and feces following additional pro-cessing that occurs principally in the kidney and intestine. It is therefore not
surprising that the cellular concentration of glutathione in the hepatocyte, the
major site of the bioactivation of chemicals, is high (about 10 mM). Depletion
of the tissue stores of glutathione, whether by chemicals or as a consequence
of poor nutrition, can potentiate markedly chemical toxicity. The toxicity of
the mild analgesic and antipyretic drug paracetamol (acetaminophen) is mark-edly exacerbated if the animals have been depleted of glutathione as a result
of inadequate nutrition (4).
It is evident that a chemical is subject to metabolism through a number of
pathways, most of which will result in deactivation. Certain routes of metabo-lism, however, will produce deleterious intermediates that are themselves sub-ject to deactivation through metabolism. Clearly, the amount of reactive
intermediates available for interaction with cellular components, and hence the
degree of toxicity, is largely dependent on the competing pathways of activa-tion and deactivation. If an animal species favors the activation pathways of a
chemical it will be susceptible to its toxicity whereas if deactivation is favored
it will be resistant. 2-Aceylaminofluorene is a carcinogen that undergoes
N-hydroxylation to generate the reactive, carcinogenic intermediates. The
guinea pig is unable to catalyze this reaction and consequently it is very resis-tant to the carcinogenicity of this chemical (5). Clearly, toxicity is not simply a
consequence of the intrinsic molecular structure and physicochemical proper-ties of the chemical, but is also largely dependent on the nature and level of the
enzymes present in the living organism at the time of exposure. Normally, the
activation pathway is a minor route of metabolism, but under certain circum-stances it may assume greater importance. Such a situation arises when the
deactivation pathways are saturated, as a result of intake of high doses of the
chemical or when the activation pathway is selectively induced, for example,
as a result of prior exposure to chemicals. Under such circumstances, the pro-duction of reactive intermediates is stimulated, overwhelming the deactivation
pathways, making an interaction with cellular constituents, and the ensuing
toxicity, more likely. Paracetamol is a very safe drug, the activation pathway
being a very minor metabolic route. In alcoholics, as a result of alcohol intake,
122 Barnett and Ioannides
this pathway is more active so that they may suffer hepatotoxicity even when
consuming therapeutic doses of the drug (6).
1.2. The Cytochrome P450-Dependent
Mixed-Function Oxidase System
Many enzyme systems participate in the metabolism of chemicals, both
Phase I and Phase II. In Phase I metabolism, undoubtedly the most important
are the cytochrome P450-dependent mixed-function oxidases, a ubiquitous
enzyme system found in almost every tissue, the highest concentration being
encountered in the liver (7). It is a very versatile enzyme system, capable of
metabolizing structurally very diverse chemicals of markedly different molecu-lar shape and size. There are not many chemicals that find their way in the
human body and are not metabolized, at least to a small extent, by the cyto-chrome P450 system and many are metabolized exclusively by this system. It
has evolved to deal with the increasing number of xenobiotics to which humans
are exposed. In addition to metabolizing xenobiotics, this enzyme system also
makes a major a contribution to the metabolism of endogenous substrates such
as steroid hormones, vitamins, fatty acids, and prostaglandins.
The cytochrome P450 system comprises an electron transport chain consisting
of the flavoprotein cytochrome P450 reductase and the hemoprotein cytochrome
P450, which acts as a terminal oxidase. It catalyzes the incorporation of one atom
of molecular oxygen to the substrate while the second atom forms water. Cyto-chromes P450 achieve their broad substrate specificity by existing as a superfam-ily of proteins, divided into families on the basis of their structural similarity.
Each family may be subdivided into subfamilies that may contain one or more
proteins. For example, the cytochrome P450 family one (CYP1) comprises two
subfamilies, namely A (CYP1A) and B (CYP1B). The CYP1A subfamily con-sists of two proteins (isoforms), CYP1A1 and CYP1A2. Only a single protein
belongs to the other subfamily (CYP1B1). Of the cytochrome P450 families, the
most important contributors to xenobiotic metabolism are CYP1–CYP3, and the
major characteristics of these are summarized in  Ta ble 1. The CYP1 family
appears to be the most important in the bioactivation of chemicals (8,9).
1.3. Regulation of Cytochromes P450
It has long been appreciated that humans differ markedly in the way they
respond to the same chemical exposure. Dramatic interindividual differences
have been noted in the blood levels of drugs following the intake of the same
dose, which could account for the fact that they experienced different pharma-cological effects, a number developing adverse effects commensurate with over-dosage. It has now been recognized that individuals may metabolize the same
drug at different rates, reflecting the activity of their drug-metabolizing enzymes,
Xenobiotic-Metabolizing Enzymes 123
particularly the cytochromes P450. Many factors are responsible for the marked
interindividual drug-metabolizing activity, including genetic background, nutri-tional status, presence of disease, and previous exposure to other xenobiotics.
Undoubtedly one of the factors that govern cytochrome P450 activity is
genetic makeup. Indeed, it was established some two decades ago that cyto-chrome P450 isoforms may be polymorphically expressed. This realization
followed observations that some persons, about 5–10% of Europeans, displayed
exaggerated responses after the intake of therapeutic doses of the antihyper-tensive drug debrisoquine. This drug is normally deactivated through
4-hydroxylation but the poor metabolizers are unable to carry out this pathway
because they lack a functioning CYP2D6, the cytochrome P450 enzyme
catalyzing this reaction (10).
The expression of cytochrome P450 activity is also regulated by the levels
of endogenous hormones as well by disease and especially previous exposure
to other chemicals, capable of inhibiting or inducing one or more cytochrome
P450 proteins. A number of studies established the importance of hormones
such as androgens, growth hormone, and thyroid hormone in the regulation of
cytochrome P450 enzyme (11). Changes in the levels and patterns of excretion
of hormones may result in selective modulation of cytochrome P450 proteins.
For example, hyposecretion of growth hormone has been implicated in the alter-ations in the hepatic profile of cytochromes P450 observed in animals with
insulin-dependent diabetes mellitus (12).
Exposure to environmental chemicals, as well as many drugs, can up-regu-late cytochrome P450 proteins in the liver and other tissues, so that chemicals
ingested after induction has occurred will be more extensively metabolized if
they rely on the induced enzymes for their metabolism. Human cytochrome
P450 proteins have been shown to be induced by alcohol; consumption of cru-Table 1
Principal Characteristics of the Xenobiotic-Metabolizing
Cytochrome P450 Enzymes
Typical drug Role in Inducibility
Family Subfamily substrate bioactivation by chemicals
CYP1 A Theophylline Very extensive Very high
B Theophylline Very extensive High
CYP2 A Propranolol Limited Modest
BWarfarin Limited High
CTolbutamide Poor Modest
D Debrisoquine Poor Not inducible
E Chlorzoxazone Extensive High
CYP3 A Erythromycin Limited High
124 Barnett and Ioannides
ciferous vegetables or of charcoal-broiled beef; smoking; and intake of drugs
such as omeprazole, rifampicin, and phenobarbitone. Cytochrome P450 pro-teins can also be down-regulated following exposure to exogenous chemicals
so that the metabolism of any subsequently ingested chemicals will be sup-pressed if they rely on the inhibited enzymes for their metabolism. This is very
much highlighted in the recently recognized interaction between normal grape-fruit juice and a number of drugs including dihydropyridine calcium channel
blockers such as felodipine, nisoldipine, and nifedipine as well as of other drugs
such as quinidine, midazolam, terfenadine, and cyclosporine (13). Simulta-neous consumption of grapefruit juice with these drugs resulted in higher
plasma levels than anticipated, leading to increased adverse effects. These drugs
are extensively metabolized in the intestine by CYP3A4, which is effectively
inhibited by grapefruit juice. Chemicals can have a differential effect on the
expression of individual cytochromes P450. For example, the dietary chemical
diallyl sulfide, a naturally occurring chemical in garlic, down-regulates
CYP2E1 but up-regulates CYP2B in the liver of rats (14).
Cytochrome P450 activity is also influenced by the presence of disease.
Again the effects are selective, in that only certain isoforms are influenced,
with some being depressed whereas others are stimulated. Hepatic disease such
as hepatocellular carcinoma has been shown to perturb the profile of cyto-chromes P450 in patients with cirrhosis and hepatocellular carcinoma (15). In
these situations the disease affects the liver itself, but hepatic cytochrome P450
levels may also be modulated in diseases where the liver is not the primary
target of disease such as insulin-dependent diabetes mellitus (12).
1.4. Drug Metabolism in the Aged
Although in animal studies drug-metabolizing activity has been reported to
diminish in the old, the limited studies conducted in humans do not appear to
support the view that age is an important determinant of drug metabolism activ-ity. Although a number of drugs are poorly eliminated in the elderly, this does
not necessarily reflect reduced metabolic, including cytochrome P450, activ-ity. They may be secondary to the normal physiological changes that accom-pany old age such as decreased renal capacity to excrete drugs and their
metabolites, reduced liver blood flow, decrease in liver mass, and changes in
plasma protein levels, one or all of which may account for the impaired drug
elimination. Both renal glomerular filtration and tubular function are altered in
the aged without any signs of kidney dysfunction. Blood flow in the old may be
as little as half of that of the adult, affecting particularly the elimination of
drugs having a high extraction ratio, which is defined as the difference in the
concentration of drug entering and coming out of the liver divided by the con-centration of drug entering the liver. Plasma levels of albumin, the major pro-
Xenobiotic-Metabolizing Enzymes 125
tein to which drugs bind, decrease in the aged, presumably the consequence of
reduced synthesis, leading to lower protein binding.
In animal studies, it was repeatedly shown that the metabolism of many
xenobiotics declines in old age, resulting in prolongation of the pharmacologi-cal effect of drugs (16,17). However, evidence for reduced capacity in the metab-olism of drugs through cytochrome P450 catalyzed pathways is lacking (18). It
is also feasible that the various cytochrome P450 proteins respond differently to
the onset of old age, but this remains to be investigated. In studies carried out in
male rats, aged between 1 wk and 2 yr, cytochrome P450 activity was investi-gated by determining the hydroxylation of testosterone at different positions
and was complemented by immunological determination of the apoprotein lev-els (19). It was evident that age-dependent changes differed among the cyto-chrome P450 enzymes studied. For example, hepatic levels of CYP2C11
disappeared in old age whereas CYP2A1 levels increased and those of CYP2E1
were unaffected.
The recent availability of in vivo probes displaying selectivity for specific
cytochrome P450 forms has made it possible to assess the effect of age on cyto-chrome P450 expression in humans. The N-demethylation of erythromycin,
a diagnostic probe for CYP3A4, the major cytochrome P450 enzyme in the
human liver, was determined by measuring the amount of carbon dioxide
exhaled. No difference was detectable between healthy aged volunteers
(age ranging between 70 and 88 yr) in comparison to younger adults (age rang-ing between 20 and 60), showing that the expression of CYP3A is not affected
by age in humans (20). Similarly, the expression of CYP2E1 was constant in
humans aged between 30 and 75 yr of age (21). In more recent studies, employ-ing as probes lignocaine (CYP3A4) and coumarin (CYP2A6), a decrease was
reported in the levels of these drugs with increasing age (22). In these studies
the authors compared healthy young volunteers (<25 yr) with healthy elderly
volunteers (>65 yr). Similarly, in recent extensive studies, the half-life of anti-pyrine, a drug whose metabolism involves a number of cytochrome P450 pro-teins, increased in the elderly whereas its clearance decreased (23). Clearly, the
effect of age on individual cytochrome P450 enzymes is far from being under-stood, and it is only now that such studies are being undertaken. It must be
emphasized that the old consume a disproportionate number of drugs com-pared with other subpopulations and an understanding of their ability to handle
drugs will lead to more effective treatment.
1.5. Measurement of Cytochrome P450 Activities
in Human Liver Using Alkoxyresorufins
In 1974, Burke and Mayer  (24) demonstrated that an alkoxyphenoxazone
derivative, ethoxyresorufin, could be metabolized by CYP1A1 with a high
126 Barnett and Ioannides
specificity. This specificity has been demonstrated for many animals species,
including humans. Furthermore, the use of methoxyresorufin and
pentoxyresorufin derivatives allow the measurement of CYP1A2 and CYP2B
proteins with high selectivity.
The metabolism of the alklyphenoxazone derivatives can be measured using
microsomal or whole cell protein (26). Both methods are comparable except
for the inclusion of dicoumarol in the whole cell protein method to prevent the
cytosolic reduction of resorufin to a nonfluorescent molecule by NAD(P)H
1.6. Method for the Measurement of Ethoxy-, Methoxy-, and
Pentoxyresorufin Dealkylation Using Human Liver Samples
This method can be used for measurement of CYP1A1 (ethoxyresorufin),
CYP1A2 (methoxyresorufin), and CYP2B (pentoxyresorufin) activities in
microsomal protein fractions or cell homogenates from primary hepatocytes or
cultured cells. It should be noted that ethyoxyresorufin may be deethylated to
some extent also by CYP1A2. In human liver, expression of CYP1A1 is very
low and ethoxyresorufin O-deethylase activity is largely attributable to
CYP1A2, although a small contribution from other subfamilies such as CYP2C
cannot be excluded.
2. Materials
2.1. Preparation of Hepatic Subcellular Fractions
1. Potter–Elvehjem glass–Teflon homogenizer (BDH, Poole, Dorset).
2. 1.15% (w/v) KCl, 4°C.
3. Refrigerated centrifuge capable of producing 9000g, ultracentrifuge.
2.2. Measurement of Cytochrome P450 Activities Using
For the direct measurement of alkoxyresorufin  O-dealkylase activity the
method of Burke and Mayer (24) can be used.
1. Pentoxy-, ethoxy- and methoxyresorufin as well as resorufin can be obtained from
Molecular Probes, Eugene, OR, USA. Alkoxyresorufins are dissolved in
dimethylformamide (Sigma Chemical, Dorset, UK) to provide stock concentra-tions of 0.53 mM ethoxyresorufin, 1 mM pentoxyresorufin, and 0.53 mM
methoxyresorufin. These solutions can be stored at –20°C until required and
should be maintained in the dark at all times. Resorufin is also dissolved in
dimethylformamide to produce a 0.1 mM stock solution. This fluorescent com-pound can be stored at –20°C in the dark until required.
2. Tris-HCl buffer (0.1 M, pH 7.8) prepared by dissolving 0.1 moles of Tris base
(Sigma Chemical, Dorset, UK) in 850 mL of distilled water. Using a calibrated
Xenobiotic-Metabolizing Enzymes 127
pH meter the pH of the buffer is adjusted to 7.8 using 3 M HCl. The buffer is
transferred to a 1-L volumetric flask and made up to the 1-L mark with distilled
water. The pH of the buffer solution is confirmed at pH 7.8 using a pH meter.
3. NADPH (50 mM, Sigma Chemical, Dorset, UK) is dissolved in 1% (w/v) sodium
hydrogen carbonate and kept at 4°C until required. This solution is made fresh
prior to performing the assays.
4. Dicoumarol (20 mM, Sigma Chemical, Dorset, UK) is prepared by dissolving
dicoumarol in 0.1 M Tris-HCl buffer, pH 7.8.
5. Spectrofluorometer with excitation wavelength of 510nm and emission wave-length of 586 nm with excitation and emission slit widths of 10nm and 2.5nm,
6. Positive controls. Samples of rodent liver microsomes that have high activity for
ethoxyresorufin, methoxyresorufin, or pentoxyresorufin can be obtained from
Xentox Limited, Northern Ireland, UK.
3. Methods
3.1. Preparation of Hepatic Subcellular Fractions
Microsomal fractions are prepared according to the method of Ioannides
and Parke (26).
1. Liver sample is weighed and and transferred to a glass beaker with volume capac-ity at least 5× that of the weight of the liver sample, for example, 10 g of liver in
a 50-mL beaker.
2. The sample is scissor-minced and transferred to the Potter–Elvehjem homog-enizer together with 3× the liver weight of 1.15% KCl (4°C).
3. Homogenize the sample using several up-and-down strokes of the homog-enizer.
4. The homogenate should be maintained at 4°C during the homogenization
process using an ice jacket (metal can filled with ice surrounding the glass
5. The homogenate is transferred to a measuring cylinder and made up to 4× the
initial sample weight with 1.15% (w/v) KCl, for example, 10 g of liver sample
made up to 40 mL of final homogenate volume with 1.15% (w/v) KCl. This is a
25% w/v liver homogenate.
6. The homogenate is transferred to centrifuge tubes and the tubes balanced for
centrifugation at 9000g for 20 min.
7. Following centrifugation at 9000g for 20 min in a refrigerated (4°C) centrifuge
the supernatant (S9) is decanted and may be stored at –70°C for up to 6 mo.
8. For microsomal preparation the S9 is transferred to ultracentrifuge tubes and bal-anced for ultracentrifugation at 105,000g for 60 min at 4°C.
9. The supernatant (cytosolic fraction) is discarded and the pellet resuspendend in a
volume of 1.15% w/v KCl equal to the volume of S9 initially placed into the
ultracentrifugation tube.
10. The microsomal suspension should be kept on ice and used the same day.
128 Barnett and Ioannides
3.2. Measurement of Alkoxyresorufin Metabolism
1. The reaction is carried out at 37°C. To a 3-mL fluorimetric cuvet (a 1.5-mL
microfluorimetric cuvet may be used with appropriate adjustment of volumes
given below) add the following reagents:
1.935 mL of 0.1 M Tris-HCl buffer, pH 7.8, prewarmed to 37°C
50 µL of microsomal suspension/cell homogenate*
3 µL of 0.53 mM ethoxyresorufin
5 µL of 0.53 methoxyresorufin
5 µL of 1 mM pentoxyresorufin
*If using cell homogenate (S9) then dicourmarol should be added to give a final
concentration of 10 µM (substitute 10 µL of Tris buffer for 10 µL of dicoumarol
stock solution).
2. The cuvet is introduced into the spectrofluorometer and a baseline recorded prior
to initiation of the reaction with 10 µL of NADPH solution.
3. The reaction is monitored continuously until a measurable gradient is obtained
and an initial rate of reaction can be determined.
4. Resorufin production from the alkoxyresorufin substrate can be calculated using
the standard resorufin solution.
5. A blank is prepared by replacing the microsomes or cell homogenate with 50 µL
of Tris-buffer.
6. Following the establishment of baseline fluorescence, at least three 10-µL samples
of the standard resorufin are introduced into the cuvet, noting the increase in
fluorescence after each addition.
3.3. Example Calculation
1. Ten microliters of 0.1 mM resorufin caused an increase of 15.5 fluorescence units.
2. Fifty microliters of sample A caused an increase of 1.2 fluorescence units per minute.
3. Therefore 1 mL of sample A would cause 1000/50 × 1.2 unit increase per minute
or 24 units per minute.
4. Ten microliters of 0.1 mM resorufin is equal to 1nmole of resorufin, therefore
1 nmol of resorufin will cause a 15.5 unit increase in fluorescence.
5. As such 1 mL of sample A produced 24/15.5 nmol resorufin per minute =
1.5 nmol/min/mL.
6. Having established the protein concentration in the microsomal suspension/cell
homogenate, the activity can be expressed as nmol/min/mg of protein.
Many spectrofluorometers will perform these calculations directly follow-ing the calibration step with the resorufin standard.
4. Notes
1. A baseline cannot be established as the fluorescence is increasing at a steady rate.
The alkoxyresorufin substrate may have become contaminated with NADPH or S9.
Make up fresh substrate and ensure that a separate pipet is used for each addition.
Xenobiotic-Metabolizing Enzymes 129
2. No reaction appears to be occurring. Check reaction system using positive con-trols. Check fluorimeter settings. Ensure that buffer is warmed to 37°C.
3. No reaction seems to be occurring. Increase amount of microsomal or S9 suspen-sion added and reassay.
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Assessing Antioxidant Status 133
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Assessing Age-Related Changes
in Antioxidant Status
The FRASC Assay for Total Antioxidant Power and Ascorbic Acid
Concentration in Biological Fluids
Iris F. F. Benzie and John J. Strain
1. Introduction
There is accumulating evidence that oxidative damage to protein, lipid, car-bohydrate and DNA is an important cause and/or effect of cellular and subcel-lular changes associated with disease, and is responsible for at least some of
the physiological, but ultimately fatal, changes that accompany aging (1–8).
Advancing age brings increasing risk of chronic degenerative disease includ-ing cancer, cardiovascular disease, cataracts, and dementia  (1–6,8). Immune
status declines, with consequent increased risk of infection and, owing to a
combination of physical and socioeconomic factors, nutritional status is often
poor in the elderly, increasing the likelihood of poor antioxidant status (9).
Improved antioxidant status helps minimize oxidative damage, and this may
delay or prevent pathological change (8–22). This suggests the possible utility
of antioxidant-based dietary strategies for lowering risk of chronic, age-related
disease (20–26). Vitamin C (ascorbic acid) and vitamin E (mainly  α-toco-pherol) are dietary derived antioxidants of major physiological importance
(25,27–31), but many other exogenous and endogenous antioxidants contrib-ute to the overall antioxidant status of the body (20,23,26,31,32). It is not yet
possible to say that benefit to health is attributable to specific antioxidants at
specific intake or plasma levels. Indeed it is likely that an optimal level of each
antioxidant is required for maintenance of optimal health, that is, that an opti-mal “total” antioxidant status is desirable (23,27,33,34). In addition, the vita-min C to vitamin E ratio may be important, with risk of oxidative stress related
134 Benzie and Strain
disease increasing at ratios below 1.0 (34). What constitutes optimal antioxi-dant status is not yet clear, however, and further study of the role of antioxidant
status, and of individual antioxidants and their interrelationships, in aging and
age-related disease is needed.
The ferric reducing (antioxidant) power (FRAP) assay,1 and its modified
version, the simultaneous ferric reducing (antioxidant) power and ascorbic acid
(FRASC) assay, is a technically simple, inexpensive, fast, sensitive, and robust
biochemical test useful for the assessment of antioxidant status of biological
fluids (35–37). FRASC can be performed using routinely available laboratory
equipment, and permits the direct measurement of biological fluids such as
blood plasma, cerebrospinal fluid, and urine. The antioxidant and ascorbic acid
content of these fluids, and of extracts of various drugs and dietary agents, can
be measured objectively and reproducibly using FRASC, allowing their poten-tial for improving the antioxidant status of the body to be assessed and com-pared (37–42). The assay is also analytically sensitive and precise enough to
assess postingestion response to dietary antioxidants. FRASC, therefore, offers
a practical analytical tool to help assess diet-, disease-, or age-related changes
in antioxidant status.
1.1. Rationale of the FRASC Assay for Total Antioxidant Power
and Ascorbic Acid Concentration in Biological Fluids (36,37)
In this assay, a ferric-tripyridyltriazine (FeIII–TPTZ) complex is reduced to
its ferrous form, which is blue colored and absorbs light of 593 nm. The ferric to
ferrous reaction is driven by the reductive action of electron donating antioxi-dants in the test sample, and the change in absorbance at 593 nm is directly
proportional to the combined, or “total,” reducing (antioxidant) power of these
antioxidants (35). In FRASC, ascorbic acid in the sample is selectively and spe-cifically destroyed by ascorbate oxidase. The change in absorbance in this case
is attributable to the remaining antioxidants, that is, the “total” less the contribu-tion of ascorbic acid. The difference in antioxidant power between two paired
samples, one treated with ascorbic oxidase and one untreated (and therefore still
containing ascorbic acid) is equal to the contribution of ascorbic acid in the
untreated sample (see Fig. 1). It is then a simple matter to calculate the molar
concentration of ascorbic acid in the test sample, and to obtain three indices of
antioxidant status: (1) the total antioxidant power, (2) the ascorbic acid concen-tration, and (3) the “non-ascorbic acid” antioxidant power of the sample.
2. Materials
1. 0.3 M Acetate buffer, pH 3.6; dissolve 3.1 g of sodium acetate trihydrate (Riedel-de Haen, Germany) in approx 500 mL of distilled and deionized water; add 16
1 U.S. patent pending.
Assessing Antioxidant Status 135
mL of glacial acetic acid (BDH Laboratory Supplies, England), and make up to a
final volume of 1 L with distilled and deionized water. This solution can be stored
at room temperature for up to 1 mo.
2. 0.01  M TPTZ (2,4,6 tripyridyl-s-triazine, Fluka Chemicals, Switzerland) in
0.04 M HCl (BDH). This solution can be stored at room temperature for up to
2 wk (see Note 1).
3. 0.02 M FeCl3
.6H2O (BDH). This solution can be stored at room temperature for
up to 2 wk.
4. 4000 U/L of ascorbate oxidase (EC (Sigma Chemical, St. Louis, MO,
USA) in distilled water. Aliquots of this solution should be stored at –70°C and
thawed when required.
5. To prepare working FRASC reagent, mix 20.0 mL of 0.3 M acetate buffer, pH
3.6, 2.0 mL of 0.01  M TPTZ solution in 0.04  M HCl; and 2.0 mL of 0.02  M
FeCl3·6H2O solution just before use.
Fig. 1. Measuring concept of FRASC. This figure shows the absorbance change
owing to FeIII reduction by antioxidants in the sample. Calculation of FRAP value is by
taking the 0–4-min change in absorbance at 593 nm for test sample (closed circles, 1)
and relating it to the 0–4-min absorbance change for the FeII standard (closed triangles,
2), with a reagent blank correction (open triangles, 3) for both. Calculation of ascorbic
acid results is by subtracting the 0–1-min absorbance reading of the ascorbate oxidase-treated test sample (open circles) from the matching water-treated sample (closed circles,
4); this signal is then related to that given by a standard solution of FeII (closed tri-angles) (or ascorbic acid, closed squares, 5) of appropriate concentration. (Reproduced
with permission from Benzie, I. F. F. and Strain, J. J. (1997) Redox Report 3, 233–238.)
136 Benzie and Strain
6. For calibration, standard solutions of FeSO4·7H2O (Riedel de Haen, Germany)
made up in water to a known concentration; for example, 100 µM, 500 µM, and
1000 µM are recommended (see Note 2). Reaction of FeII represents a one-elec-tron exchange reaction and is taken as unity, that is, the blank corrected signal
given by 1000 µM solution of FeII is equivalent to a ferric reducing/antioxidant
power (FRAP) value of 1000 µM. As ascorbic acid has a stoichiometric factor of
2.0 in this assay (35–37), reaction of ascorbic acid gives a change in absorbance
double that of an equivalent molar concentration of FeII , that is, a FeII standard of
100 µM is equivalent to 50 µM ascorbic acid, and a 100 µM solution of ascorbic
acid has a FRAP value of 200 µM.
Freshly prepared aqueous ascorbic acid solutions (ascorbic acid extra pure
crystals, Sigma Chemical, St. Louis, MO, USA) can also be used as calibrators
(36,37,42) (see Note 3). Refer to Note 4 for guidelines on quality control samples
and expected linearity and precision.
3. Methods
1. Samples: Serum, plasma, urine, saliva, tears, other biological fluids, and aqueous
and ethanolic extracts of drugs and foodstuffs can be used directly in FRASC.
However, as some antioxidants are unstable, samples should be kept chilled and
in the dark until testing, and should be tested with as little delay as possible.
Hemolyzed plasma or serum samples should be avoided. Heparinized plasma is
preferable to EDTA plasma and serum for FRAP and FRASC measurements, as
ascorbic acid is more stable in heparinized plasma (39).
2. To measure the total antioxidant power, as FRAP, and ascorbic acid in one test
(FRASC), ascorbic acid in one of a matching pair of sample aliquots is destroyed
by the addition of ascorbate oxidase. Ascorbic acid reacts very quickly with the
working reagent, and the 0–1-min reaction time window is used for calculation of
ascorbic acid results; the 0–4-min window is used for calculation of “total” anti-oxidant power (FRAP) results, that is, absorbance readings are taken at 0, 1, and
4 min after reagent/sample mixing (37°C incubation).
3. To prepare samples:
a. Add 40  µL of a 4000 U/L solution of ascorbic oxidase to 100  µL of test
b. Add 40 µL of distilled water to the paired 100-µL sample aliquot.
c. Calibrators and QC samples are treated similarly in pairs (see Note 5). This
predilution of samples, calibrators and QC samples can be performed in the
analyzer sample cups.
4. The paired ascorbate oxidase diluted (“+ao”) and water-diluted (“–ao”) samples
are then immediately loaded onto the analyzer for automated measurement (see
Note 6).
3.2. Data Collection and Calculation of Results
1. The FRASC assay can be performed using any type of automated analyzer that
permits blank corrected readings at 593 nm to be taken at selected intervals after
Assessing Antioxidant Status 137
sample-reagent mixing. In our laboratories the Cobas Fara centrifugal analyzer
(Roche Diagnostics Ltd., Basel, Switzerland) is used, and the user-defined test
program is presented in Ta ble 1.
2. The 0–4-min reaction time window is used for data capture for the FRAP value.
The absorbance change is translated into a FRAP value by relating the change of
absorbance at 593 nm of test sample to that of a standard solution of known
FRAP value, for example, 1000 µM FeII, as described in Eq. 1.
3. To obtain ascorbic acid results, the 0–1-min absorbance at 593 nm readings are
retrieved, and calculation of results is performed as described in Eqs. 2 and 3.
4. The nonascorbic acid antioxidant power (see Note 7) is calculated according
to Eq. 4.
5. Calculation of results is as follows:
Using the water-diluted samples, the FRAP (µM) value =
[FRAP]std (µM) × 0–4 minute A593 nm test sample / (1)
0–4 minute A593 nm standard
(see Note 6).
Table 1
Cobas Fara Test Program for FRASC Assay
Measurement code Abs
Reaction mode R1-I-S-A
Reagent blank reag/dil
Wavelength 593 nm
Temperature 37°C
R1 300 µL
M1 1.0 s
Sample volume 10 µL
Diluent name H2O
Volume 30 µL
First 0.5 s
Number 17
Interval 15 s
Reaction direction Increase
Number of steps 1
Calculation Endpoint
First M1
Last 17 (i.e., 0–4 min) for FRAP; reading 5
(i.e., 0–1 min reading) is retrieved
for calculation of ascorbic acid
Modified from Benzie, I. F. F. and Strain, J. J. (1997) Redox Report 3,
138 Benzie and Strain
Using the paired water (–ao) and ascorbate oxidase diluted (+ao) samples, the
ascorbic acid concentration is calculated as follows:
0–1 min ascorbic acid related change in A593 nm = (2)
(0–1 min A593 nm sample –ao) – (0–1 min A593 nm sample +ao)
ascorbic acid concentration (µM) = [ascorbic acid] std (µM) × (3)
0–1 min ascorbic acid related A593 nm of test sample /
0–1 min ascorbic acid related A593 nm of standard
(see Note 8).
nonascorbic acid antioxidant power = (4)
FRAP value (µM) – 2 × ascorbic acid concentration (µM)
Table 2 gives typical values obtained on fasting plasma samples from healthy
4. Notes
1. The TPTZ powder, as purchased, is normally white. However, in some bottles,
when opened, the contents have been found to be gray or yellow in color.
The reason for this coloration is not clear, but it does not appear to affect results.
The working FRASC reagent should be a pale yellow/orange color. Any visible
blue color indicates contamination by either ferrous iron or a reducing agent.
Visible blue color in the working reagent will give a high blank reading and
decrease sensitivity: do not use.
2. Do not attempt to use FeII standards >1500 µM, as there will be precipitation of
iron salts. Aqueous ferrous sulfate solutions of up to 1500 µM appear to be stable
for at least 1 mo at 4°C. These solutions should be clear and colorless.
3. It is important that ALL samples, calibrators, and QC samples be treated identi-cally and in matching ascorbate oxidase diluted and water diluted pairs. The same
absorbance reading should be obtained for the paired FeII solutions, that is, the
Table 2
FRAP Values and Ascorbic Acid Concentrations
(Mean; Median; SD; µmol/L), Using FRASC, of Fresh Fasting
EDTA Plasma from Healthy Subjects
All (n = 130) Men (n = 66) Women (n = 64)
Age (years) 43; 43; 16.4 42; 42; 16.3 43; 44; 16.6
FRAP 1018; 1004; 198 1086; 1077; 189 948; 927; 183
Ascorbic acid 51; 48; 17.9 49; 48; 13.8 52; 50; 21.3
Reproduced with permission from Benzie, I. F. F. and Strain, J. J. (1997) Redox Report 3,
Assessing Antioxidant Status 139
addition of ascorbate oxidase should not result in any difference in the absor-bance given as there is no ascorbic acid to destroy in these solutions.
4. Guidelines on quality control samples and expected performance:
For ease of use and reliability, aqueous ascorbic acid solutions at 100, 250,
500, and 1000 µM (equivalent to 200, 500, 1000, and 2000 µM FRAP, prepared
fresh daily) and aged QC serum freshly spiked with ascorbic acid are recom-mended as quality control samples. These should be run in parallel with test
samples to actively monitor the performance of the test and to ensure compara-bility with previous results.
Expected precision and sensitivity of the FRASC assay:
Precision is high in FRASC: typical within- and between- run CVs obtained in
our laboratories are, respectively, <1% and <3% at 900 and 1800 µM for FRAP
values. For ascorbic acid, typical within- and between-run CVs are <5% at 25,
50, 100, and 440 µM. Sensitivity is also high, and the limit of detection of the
FRAP assay is <2 µM reducing/antioxidant power.
Expected linearity:
The test as described will give a linear dose–response up to a FRAP value of
2000 µM. If samples with FRAP values of >2000 µM are to be tested, prior dilu-tion of the test sample in water is recommended. As stoichiometric factors are
constant in the FRASC assay (35), simple correction for the additional dilution is
the only extra calculation required to obtain the FRAP value of the sample.
Linearity of ascorbic acid dose–response of the test as described is up to at least
1000 µM.
5. The enzyme-linked destruction of ascorbic acid is very fast, even at room tem-perature, and additional incubation time with ascorbate oxidase is not needed.
6. The “nonascorbic acid antioxidant power” is the combined antioxidant power of
the other, similarly acting antioxidants such as uric acid and bilirubin (see refs.
35 and 37). There is also some contribution (approx 10% of the total) by the thiol
groups of plasma proteins.
7. This additional step can easily be added to the test program if desired, and a
direct printout of FRAP values in micromolarity obtained. Raw data are then
retrieved, with capture of the 0–1-min change in absorbance at 593 nm (i.e., read-ing number 5 on test as programmed) for calculation of the ascorbic acid concen-tration.
8. The value of the standards used for calculation of micromolar ascorbic acid and
for FRAP values must be expressed in micromolar ascorbic acid or FRAP value
“equivalents” as appropriate. Note that each molecule of ascorbic acid has the
ability to reduce two ferric ions, and therefore has a FRAP value of 2 µmol. Aque-ous ascorbic acid standards are stable for 24 h at 4°C. Ferrous sulfate solutions
are stable for at least 1 mo at 4°C.
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Comet Assay of DNA Damage 143
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Measurement of DNA Damage and Repair Capacity
as a Function of Age Using the Comet Assay
Peter H. Clingen, Jillian E. Lowe, and Michael H. L. Green
1. Introduction
A large number of studies indicate that DNA damage and mutation increase
with age in human cells and tissues (1). Age-related degenerative disorders in
which DNA damage has been invoked include heart disease and
neurodegenerative conditions such as Alzheimer’s disease, amyotrophic lateral
sclerosis, or Parkinson’s disease (2,3). Patients with deficiencies in DNA repair,
including xeroderma pigmentosum (XP) (4) and ataxia–telangiectasia (A–T)
(5) show characteristic patterns of neurodegeneration (as opposed to a failure
of normal development). The implication is that failure of repair can lead to
accumulation of damage and degenerative disease. XPs and A–Ts are hyper-sensitive to specific types of DNA damage, and the degenerative damage in
patients is tissue specific. DNA in every tissue, however, is under attack from a
range of endogenously formed mutagens, including reactive oxygen species,
nitric oxide, reactive metabolites, and breakdown products such as malon-dialdehyde. A series of repair enzymes recognize and remove these types of
DNA damage from the genome. Failure to repair DNA may cause the synthesis
of defective proteins, which will repair DNA less efficiently, and in turn lead to
propagation of further errors (6). Alternatively, oxidative damage to mitochon-drial proteins might cause less efficient processing of oxygen, release of higher
levels of reactive oxygen species and increased levels of background DNA
In this chapter, we describe protocols based on modifications of the alkali
Comet assay (single-cell gel electrophoresis) and describe how they may be
applied to the measurement of DNA damage and repair as a function of age.
The Comet assay is a simple, rapid, and inexpensive method for detecting DNA
144 Clingen, Lowe, and Green
strand breaks in individual eukaryotic cells. It has been used in vivo and in
vitro to assess DNA damage and repair induced by various physical factors
such as ultraviolet (UV) radiation  (7,8), ionizing radiation  (9,10), chemical
agents (11) as well as physiological factors such as diet (10), cytokines (12),
exercise (13), smoking, and aging  (14,15). A range of applications will be
addressed including:
• Detecting frank DNA strand breaks
• Investigating endogenous levels of damage
• Formation and repair of oxidative DNA damage using specific DNA repair
• Measuring intrinsic rates of cellular excision repair
Consideration is also given to experimental design and interpretation of data
so that meaningful results may be obtained.
1.1. Principle of Assay
The Comet assay is a flexible and sensitive procedure for measuring DNA
damage in a range of tissues and cell types. The methods described here have
been largely based on the protocols of Singh et al. (16) and Collins et al. (17).
Methods and applications of the assay have been extensively reviewed (18–22).
In the assay, a single-cell suspension of the cells or tissue under study are
embedded in low melting point agarose on a frosted microscope slide. The
slide is placed in a high salt lysis mixture that strips away cell membranes and
removes most proteins from chromatin to leave behind supercoiled DNA in
structures known as nucleoids (23). When placed in alkaline buffer the super-coiled DNA starts to unwind. During electrophoresis, DNA containing strand
breaks is pulled toward the anode to form a Comet tail while undamaged DNA
remains trapped within the nucleus. Comet tail length or tail moment are then
determined by image analysis. Within a defined dose range there appears to be
a linear relationship between the number of strand breaks and measures of
Comet length and/or tail moment. The assay specifically measures DNA strand
breaks and normally can detect less than one strand break per 107 basepairs.
These may represent:
• Direct single-stranded breaks produced by exogenous or endogenous mutagens
such as reactive oxygen species
• Alkali labile sites
• Breaks formed during the excision step of repair of damaged bases from cellular
Direct single-stranded breaks are detected using the standard form of the
assay. Although it has been postulated that these include alkali-labile sites, it is
Comet Assay of DNA Damage 145
controversial whether the conditions of the assay are sufficiently alkaline to
generate these sites. As a means to study overall levels of DNA excision repair
it is possible to incubate cells, damaged with genotoxic chemical or physical
agents, with inhibitors of DNA strand resynthesis such as aphidicolin, cytosine
arabinoside, or hydroxyurea. These inhibitors allow the accumulation of strand
breaks formed during DNA excision repair. Although this assay is nonspecific
in that it does not discriminate between the types of damage that are being
repaired it may be used to compare overall rates of excision repair. A more
exact approach is to use purified DNA repair enzymes. Cells embedded in aga-rose are treated with a damaging agent and lysed immediately or at various
times after treatment. Exposed nucleoids are then washed and incubated with a
repair enzyme that recognizes and cleaves DNA at the sites of specific DNA
lesions. Suitable enzymes include (1) T4 endonuclease V, which recognizes
ultraviolet radiation induced cyclobutane pyrimidine dimers (24); (2) endonu-clease III (Endo III), which recognizes oxidized pyrimidines  (17); (3)
formamidopyrimidine glycosylase (FPG), which recognizes oxidized purines
(25,26); and (4) uracil glycosylase, which recognizes uracil in DNA (27).
1.2. Practical Considerations
In theory, any cell type from any tissue is suitable for use in the Comet
assay provided that it can be made into a single-cell suspension without intro-ducing significant amounts of DNA damage. In practice and particularly for
age-related studies, the choice of cell type may be limited. Samples from a
large number of individuals should be available as should routine methods for
culture of the cell type. Age groups of donors should be wide enough apart to
maximize the opportunity for seeing any differences in the levels of DNA
damage or repair (e.g., age groups of 25–30, 45–50, and 65–75). Caution must
be taken when averaging levels of DNA damage or repair and sufficient
samples need to be analyzed to take into account interindividual variation.
Because both physical and physiological factors can affect levels of back-ground DNA damage  (7–15) these should be defined and controlled for as
carefully as possible. Details of medical history and lifestyle factors such as
recent exercise or smoking habits, alcohol consumption, and diet should be
obtained where possible.
The cells that lend themselves most readily to these studies are freshly iso-lated peripheral blood mononuclear cells, most of which are in a nondividing
(G0) state. Details of how to handle cultured human fibroblasts in the Comet
assay are also described. Although it would be possible to grow primary cul-tures of fibroblasts from small skin biopsies for a number of individuals of
different ages, the resources and time required to do so for a significant number
of donors would be considerable. Cultured fibroblasts would lend themselves
146 Clingen, Lowe, and Green
more readily to in vitro studies of aging, and to studies of genotypes that might
influence the aging process.
In addition to interindividual variation, intraexperimental variation should
be considered. It is essential to repeat experiments several times, to confirm
consistency of effects. A minimum of duplicate slides in three independent
experiments is recommended. This is particularly important in monitoring
populations for evidence of preexisting or endogenous DNA damage. Age-related
differences may be small and experiments may not have built-in controls for
damage arising from poor experimental technique. As a guide to what can be
achieved within a given study, one worker can readily perform and score one
experiment of eight sets of duplicate slides per day.
2. Materials
2.1. Cells and Tissues
We will describe the use of the assay with (1) freshly isolated human mono-nuclear cells, and (2) cultured normal human fibroblasts
2.2. Slides, Agarose, and Chemicals
1. Slides. Fully frosted (on one surface) glass microscope slides 76  × 26 mm
(Chance Propper, Smethwick, West Midlands, UK) (see Note 1). Coverslips,
22 × 22 mm, thickness no. 1.5 (Merck, Leicester, UK).
2. Agarose. Sigma type I (Sigma, Poole, Dorset, UK); NuSieve GTG low melting
point agarose (FMC BioProducts, Rockland, ME, USA).
3. Culture medium, as appropriate for each cell type. We incubate freshly iso-lated mononuclear cells in RPMI 1640 with phenol red (Life Sciences, Pais-ley, UK), supplemented with 10% pooled human AB serum, 200 IU/mL of
Penicillin, 200 µg/mL of streptomycin, 2 mM L-glutamine (Gibco-BRL, Pais-ley, UK), and 200 µg/mL of sodium pyruvate (Sigma, Poole, Dorset, UK). For
fibroblasts, we use minimum essential medium (MEM) containing 15% fetal
calf serum (FCS), 100 IU/mL of penicillin, 100 µg/mL of streptomycin, and
2mM L-glutamic acid. Histopaque is from Sigma (Poole, Dorset, UK), human
AB serum pools from Colindale Blood Transfusion Centre (London, UK) and
FCS from PAA Laboratories GmbH (Linz, Austria). Where indicated, RPMI
1640 without phenol red is used.
4. Dulbecco’s “A” phosphate-buffered saline (PBS, tablets from Oxoid,
Basingstoke, UK). Bacto trypsin is from Difco Laboratories (Detroit, MI, USA).
The contents of one vial are dissolved in 200 mL of PBS. Trypsin-EDTA contains
40% v/v trypsin solution and 0.4% EDTA in PBS. Other chemicals are from
Sigma (Poole, Dorset, UK).
5. All solutions are made up in double-deionized water (ddH2O).
Comet Assay of DNA Damage 147
6. Lysis mixture: 10 mM Tris base, 2.5 M NaCl, 200 mM NaOH, 100 mM EDTA-Na2, pH 10. Store at room temperature for up to 1 wk. One hour before use,
place at 4°C. Just before use add 1% v/v Triton X-100 and 10% v/v dimethyl
sulfoxide (DMSO).
7. Enzyme buffer. Use the recommended buffer for the enzyme. For T4 Endonu-clease V: 10 mM Tris-HCl, 10 mM EDTA-Na2, 75 mM NaCl, pH 8.0. For Endo
III and FPG: 40 mM N-[2-hydroxyethyl]piperazine-N’-[2-ethanesulfonic acid]
(HEPES), 100 mM KCl, 0.5 mM EDTA-Na2, 200 µg/mL of bovine serum albu-min (BSA), pH 8.0. Enzyme buffers may be made as 10 × stock solutions and
stored at –20°C. T4 Endo V was a gift from Dr. D. Yarosh, Applied Genetics (NY,
USA). Endo III and FPG were a gift from Dr. A. Collins (Rowett Institute, Aber-deen, UK). Commercial sources of the enzymes are available (Trevigen, Whitney,
Oxford, UK).
8. Electrophoresis buffer: 300 mM NaOH, 1 mM EDTA-Na2, made up on day of
use. The buffer must be at a defined temperature (we typically use 15°C).
9. Neutralization buffer: 400 mM Tris base, adjusted to pH 7.5 with concentrated
HCl. This can be stored at room temperature if filter sterilized.
10. Staining solution: 20 µg/mL of ethidium bromide in ddH2O. This can be stored
indefinitely at 4°C (see Note 2).
11. Two small water baths, one at 37°C and one at 45°C. Standard histology staining
troughs and racks (cover with black electrical tape to exclude light). Trays and
source of ice.
2.3. Electrophoresis
1. Gel electrophoresis boxes. Commercial or custom-made boxes can be used. We
use purpose built Perspex boxes, 24 × 28 × 7.5 cm, covered in black electrical
tape to exclude light. These hold 16–18 slides in two rows and use 1.5 L of elec-trophoresis buffer.
2. Power pack. Delivering at least 400 mA at 20 V.
3. Refrigerator. The lysis stage of the assay is carried out at 4°C. Solutions must be
prepared and electrophoresis must be carried out at a defined temperature. An air-conditioned room or cooled incubator are also advantageous for maintaining con-stant temperatures throughout the assay.
2.4. Microscopy
1. Fluorescence microscope, with an ethidium bromide filter (see Note 2). We use a
10× objective with a video camera for scoring. An integrating monochrome CCD
video camera is required for sensitivity.
2. Image analysis system. We currently use the Casys system (Synoptics, Cam-bridge, UK) or the Komet system (Kinetic Imaging, Liverpool, UK). Although
computerized software is desirable for objective scoring, it is possible to analyze
Comets without it (see Note 3).
148 Clingen, Lowe, and Green
3. Methods
3.1. Preparation of Cellular Material
Results in the Comet assay are critically dependent on gentle handling of the
test material. We have obtained acceptable results with the procedures outlined
in the following list. Where possible all procedures should be performed under
subdued lighting. Cultured cells are maintained at 37°C in a humidified atmo-sphere containing 5% CO2. Typically 1–2  × 104 cells are required per slide
when using 22  × 22 mm coverslips. Standard experiments of 16–18 slides
require a total of approx 2.5–5 × 105 cells.
1. Whole blood samples may be obtained using a Soft Touch (Boehringer, Lewes,
UK) or similar finger-pricking device and taken up using a Gilson-type dispos-able tip pipet (see Note 4). Blood (40 µL) is suspended in an Eppendorf tube with
160 µL of clear RPMI 1640 containing four heparin-coated beads (taken from the
barrel of a blood sampling syringe). This yields sufficient cells for eight slides
when diluted with 200 µL of 1.4% LMP agarose. Samples are held at room tem-perature and slides made as quickly as possible.
2. Mononuclear cell fractions (MNCs) from individual donors can be isolated from
larger blood samples (5–50 mL) (see Notes 5 and 6). Blood is diluted 1:1 with
RPMI 1640 and 10–15 mL carefully layered onto 10 mL Histopaque in centri-fuge tubes. These are centrifuged at 700g for 20 min at room temperature. The
MNCs are removed from the Histopaque/plasma interface with a sterile plastic
Pasteur pipet and approx 5 mL transferred into individual centrifuge tubes. These
are made up to 20 mL with RPMI 1640 and centrifuged for 10 min at 700g. The
pellets are resuspended with 25 mL of RPMI 1640 containing 10% human AB
serum and centrifuged for 10 min at 700g. The pellets are resuspended with 25 mL
of RPMI containing 10% human AB serum and the number of cells counted. The
cells are centrifuged for 10 min at 700g, resuspended at approx 3 × 106 cells/mL
in 10% DMSO/90% FCS and cryopreserved in 1-mL aliquots. Typically, 1 mL of
blood should yield approx 1 × 106 mononuclear cells. Prior to use for DNA
damage or repair experiments, cryopreserved mononuclear cells should be rapidly
thawed, washed with 5 mL of complete RPMI culture medium, centrifuged at
250g for 5 min, resuspended at 106 cells/mL in complete RPMI culture medium,
and incubated at 37°C overnight (see Note 7).
3. Primary human fibroblasts may be obtained from small skin biopsies and main-tained in tissue culture flasks in MEM culture medium (28). Cells are washed
with PBS and trypsinized with fresh trypsin-EDTA. It is essential to trypsinize
for a minimum period (typically less than 5 min at 37°C is sufficient). Two
volumes of MEM culture medium are added to inactivate the trypsin, and cells
are centrifuged at 250g for 5 min and resuspended in PBS at approx 106 cells/mL.
For repair kinetic experiments on fibroblasts lasting more than 8 h it is necessary
to preincubate cells for at least 72 h in MEM containing 0.5% FCS to inhibit cell
division (see Note 8). Typically, 3 × 105 cells per dish can be maintained in a
number of 5 cm diameter tissue culture grade Petri dishes.
Comet Assay of DNA Damage 149
4. Generation of single-cell suspensions from a variety of human and rodent tissues
has been reviewed (18).
3.2. Slide Preparation
1. Prepare 3 mL of 1% type I agarose and 2 mL of 1.4% NuSieve LMP agarose (see
Note 9), made up in clear RPMI 1640 or PBS (see Note 10). Microwave for the
minimum time necessary to melt the agarose.
2. Place the type I agarose in a 45°C water bath and the LMP agarose in a 37°C
water bath.
3. Use a pencil to label the slides for each treatment. Warm the slides to at least
40°C, on a metal tray. Add 80 µL of type I agarose while the slide is still warm.
Immediately lower a 22 × 22 mm coverslip over the agarose, avoiding incorpora-tion of air bubbles. When all the slides have been treated, place the tray over ice
and leave for at least 10 min (see Note 11).
4. Prepare the cell suspension for the second layer. For whole blood samples this is
described in Subheading 3.1.1. For single-cell suspensions, resuspend cells in
medium (37°C) at 1 × 106 cells/mL (for 16 slides 500 µL is required). Add an
equal volume of 1.4% LMP agarose (i.e., 500 µL) and mix gently. From the 1-mL
cell suspension place 45 µL (containing approx 2 × 104 cells) onto the 1% agarose
base layer of each slide and add a 22 × 22 mm coverslip. Slides do not need to be
prewarmed for this step (see Note 12). Allow to set over ice for at least 5 min.
3.3. Treatment of Material with a DNA-Damaging Agent
In a number of aging studies, no specific DNA-damaging treatment is
applied, and the assay is used to detect levels of preexisting damage in blood
and other tissues. Special consideration must be given to the design of such
studies, if they are to have a chance of yielding useful data (see Note 13).
A wide variety of potential DNA damaging agents have been tested using
the Comet assay (18). Note 14 gives experimental conditions for a number of
common DNA damaging agents used in the Comet assay (H2O2, nitric oxide
donors, xanthine/xanthine oxidase, ionizing radiation, and UV-C radiation).
Depending on the nature of the experiment, treatment may be at any tempera-ture between 0 and 37°C. Treatment on ice or at 4°C is a convenient tempera-ture to minimize cellular DNA repair. For all treatments, preliminary time
course and dose–response experiments are required to establish the optimum
reaction conditions and to ensure Comet measurements are in the linear range.
1. Single-cell suspensions or monolayers of cultured cells may be incubated with
the damaging agent added directly to the growth medium or appropriate buffer.
This is suitable if extended treatment times are required. The disadvantage of this
method is that it takes up to 20 min for the slides to be prepared, during which
repair or further strand breakage can occur, before the slides can be placed in
lysis mixture.
150 Clingen, Lowe, and Green
2. Cells may be treated after they have been embedded in the agarose on the slide.
Slides without coverslips may be submerged in buffer containing the test agent.
Alternatively, 100 µL of buffer with the damaging agent may be overlaid on the
agarose, left for 10–15 s to permeate, and a coverslip added. We calculate the
concentration of damaging agent, assuming a total volume of 225 µL taking into
account the volume of the agarose layers (100 µL + 45 µL + 80 µL). Caution must
be taken to ensure that the agarose does not dry out. All incubations should be
performed in a humidified atmosphere (we typically use a black box with slides
suspended on racks over moistened tissue) or sufficient buffer applied through-out the treatment. The advantage of treating cells on the slides is that it is possible
to obtain time courses for rapid processes, such as repair of ionizing radiation-induced or oxidative DNA damage.
3.4. Standard assay for single strand breaks
This method is suitable for investigating the induction and repair of single
strand breaks following treatment with a DNA damaging agent, or for detect-ing endogenous levels of strand breakage (see Note 15).
Damaging agents such as ionizing radiation, H2O2, xanthine/xanthine
oxidase, and nitric oxide donors are capable of producing DNA strand breaks
directly. Initial levels of damage are detected immediately after treatment.
Repair is typically rapid and completed within 15–60 min. For these reasons
cells embedded in agarose should be treated on the slide with the damaging
agent suspended in suitable medium or buffer.
A typical assay of 16 slides will consist of 8 different treatments (including
controls) performed in duplicate.
1. For detecting initial levels of damage, after treatment, coverslips are removed
and one set of control and treated slides lysed immediately by gently lowering a
staining rack into a trough containing 150 mL of lysis mixture (4°C). For lysis,
slides are held for at least 1 h at 4°C. Although slides may tolerate lysis at 4°C
overnight we would recommend a minimum of 1 h and a maximum of 6 h. This
facilitates rapid repair studies, as cells incubated for 0–4 h may be lysed and
analyzed in a single experiment.
2. For repair, slides are washed 3× by gently lowering a staining rack into a trough
containing 150 mL of medium at 4°C (without damaging agent). Slides are over
laid with 100 mL of medium, a coverslip added, and incubated for 0–4 h as
required in a dark, humidified atmosphere at 37°C.
3. At the specified time, coverslips are removed from duplicate slides containing
cells that have undergone repair and the slides placed in the same lysis mixture as
described in  Subheading 3.4.1. After the last slides have been placed in lysis
incubate at 4°C for a further 1 h.
4. Slides are drained by carefully blotting their sides on tissue and placed on the gel
shelf in the electrophoresis box so that the agarose layer is positioned toward the
Comet Assay of DNA Damage 151
anode (we use two rows of eight slides). Slides should be placed close together
and all spaces filled with blanks to prevent any movement.
5. A volume of electrophoresis buffer at the correct temperature (see Note 16) is
gently added to just cover the slides (see Note 17). The slides are incubated for a
fixed period for unwinding (we use typically 40 min; see Note 16).
6. Electrophoresis. An electric current is applied (we use 20 V/24 min).
7. Remove some electrophoresis buffer by suction. Gently remove the slides and
place them on a staining tray. Gently rinse with neutralizing buffer, stand for
5 min, drain, and repeat twice.
8. Place 35 µL of ethidium bromide solution on the surface of the agarose of each
slide and add a coverslip.
9. Score slides as soon as possible. For storage, place slides at 4°C in the dark over
moist tissue in a closed box (see Note 18).
3.5. Comet Assay for Detecting Endogenous Levels
of Oxidative DNA Damage
Freshly isolated mononuclear cells exhibit little or no single strand breakage
in the standard Comet assay as described in Subheading 3.4. However, low
steady-state levels of oxidized purines and pyrimidine bases can be detected
using Endo III and FPG repair enzymes.
A typical assay will consist of samples from four donors. Each sample must
be incubated both with and without enzyme, giving two sets of duplicate slides
for each donor.
1. Slides are prepared as outlined in Subheading 3.2. and lysed immediately.
2. After lysis the slides are drained by carefully blotting their sides on tissue and
washed by placing in a fresh staining trough containing 150 mL of enzyme buffer
at 4°C for 5 min. Replace the buffer and repeat twice.
3. Slides are drained and placed on a tray. Add 50 µL of diluted enzyme (see Note 19)
to the surface of the agarose and add a coverslip to spread the enzyme solution
uniformly. Each set of duplicate slides treated with enzyme must have as a control,
duplicate slides treated with buffer minus enzyme. Incubate for 1 h at 37°C in a
dark humidified atmosphere.
4. Place the slides in the gel box and add electrophoresis buffer to stop enzyme
activity. For detection of enzyme-sensitive sites, we normally omit the unwind-ing step and apply electrophoresis immediately. However, for detecting low lev-els of endogenous oxidative damage it may be advantageous to retain an
unwinding step to increase the sensitivity of the assay.
5. Electrophoresis and subsequent steps are as described in Subheading 3.4.6.
3.6. Repair of Oxidative Base Damage
The rate of repair of oxidative base damage can also be determined using
either Endo III or FPG. In this protocol sufficient controls are required to take
into account levels of endogenous damage (untreated cells with enzyme) and
152 Clingen, Lowe, and Green
the formation of single-stranded breaks (treated cells without enzyme). Conse-quently, untreated and damaged cells incubated with or without enzyme are
required for each time point. The yield of enzyme sensitive sites (ESS) is cal-culated from increase in Comet length as follows:
ESS = [(treated cells with enzyme) – (treated cells without enzyme)] – (1)
[(untreated cells with enzyme) – (untreated cells without enzyme)].
A typical assay will consist of 16 slides from a single donor, duplicate con-trol slides incubated with and without enzyme, and three sets of duplicate slides
treated with the same concentration or dose of damaging agent and incubated
with and without enzyme for defined times.
1. Prepare slides as outlined in Subheading 3.2.
2. Treat with DNA damaging agent on the slide over ice to minimize cellular repair
of damage.
3. To establish initial levels of damage immediately place duplicate control and
treated slides into lysis mixture.
4. For repair, wash remaining slides 3× with 150 mL of medium (without damaging
agent), and add 100 µL of medium and a coverslip to the agarose layer.
5. Incubate and at defined time intervals place in same lysis mixture as described in
Subheading 3.6.3.
6. After the last slides have been placed in lysis, incubate at 4°C for a further 1 h.
7. Subsequent steps are as described in Subheading 3.5.2.
3.7. Measuring Rates of Overall Excision Repair
This assay makes use of the cell’s own DNA repair capacity to reveal DNA
damage. Rates of excision repair can be established by incubating cells with
inhibitors of DNA resynthesis either immediately or at later times after treat-ment with a DNA damaging agent. It is particularly suitable for studies using
agents such UV-C radiation which, at the doses used in the Comet assay, do not
produce detectable levels of direct strand breaks.
A typical assay will consist of 16 slides from a single donor, duplicate con-trol slides, and seven sets of duplicate slides treated with the same concentra-tion or dose of damaging agent and incubated at defined time periods for repair.
1. Prepare slides as outlined in Subheading 3.2.
2. Remove coverslip and UV-irradiate cells embedded in agarose (see Note 12).
3. For initial rate of excision repair overlay duplicate control and treated slides with
100 µL of medium containing 100 µM cytosine arabinoside and 10 mM hydrox-yurea. Incubate for 1 h in a humidified atmosphere at 37°C and transfer to lysis
mixture at 4°C (see Note 20).
4. To follow the progress of repair overlay remaining slides with 100 µL of medium
without cytosine arabinoside and hydroxyurea and incubate at 37°C for various
times up to 4 h.
Comet Assay of DNA Damage 153
5. As required, carefully drain duplicate slides and overlay agarose with 100  µL
containing 100 µM cytosine arabinoside and 10 mM hydroxyurea. Incubate for a
further 1 h in a humidified atmosphere at 37°C.
6. After 1 h, transfer slides treated with inhibitor to the same lysis mixture as
described in Subheading 3.7.3.
7. After the last slides have been placed in lysis mixture incubate at 4°C for a
further 1 h.
8. Subsequent steps are as described in Subheading 3.4.4.
3.8. Scoring
1. Remove the slides from the storage box, wipe dry and allow to warm to room
temperature to avoid condensation.
2. Identify fields containing Comets under the microscope and score, either using an
image analysis system, or assigning Comets to arbitrary categories (see Note 21).
3.9. Interpretation and Analysis
1. There is extensive discussion over the most appropriate parameter to score in the
Comet assay, whether tail length is adequate, or whether tail moment or a related
function is more appropriate. In our experience, and in most cases, the parameter
chosen makes almost no difference.
2. There has also been debate on the statistical distribution of tail moment and other
parameters. It is not difficult to show that Comet parameters are not normally
distributed, but statistical analysis should be based on the consistency of repeat
determinations and experiments, where distribution of individual Comet mea-surements will be far less critical.
3. Nevertheless, variation in levels of damage between Comets on the same slide
may be a useful indicator of the nature of the damaging event and it is desirable to
keep a record of this information.
4. Results are typically expressed as mean Comet length (or tail moment) or increase
or decrease in this parameter over untreated controls. These are arbitrary units of
DNA damage or repair. It is possible, with caution, to calibrate the number of
strand breaks against a known damaging agent, such as ionizing radiation
5. In our view the important issues are (1) design of a study with an adequate num-ber of subjects or observations; (2) repetition of experiments; (3) preparation and
disaggregation of the cells; and (4) consistency in scoring procedures.
4. Notes
1. A procedure for using standard clear slides and precoating them with agarose is
available (29).
2. Other DNA stains can be used, for instance DAPI (17) or YOYO-1 (30). Ethidium
bromide or propidium iodide are bright and show good stability.
3. Other systems include Perceptive Instruments (Little Yeldham, Essex, UK). It is
possible to obtain reasonably good quantitation without image analysis, by plac-ing nuclei into categories according to the extent of damage (31).
154 Clingen, Lowe, and Green
4. The predominant nucleated cells in whole blood are likely to be polymorpho-nuclear leukocytes, especially neutrophils. We have found that these give infor-mative results in the Comet assay and have not found undue difficulties arising
from the presence of red blood cells in the preparation (10).
5. The predominant cells in the mononuclear fraction are T cells (about 70%), with
about 10% B cells and the remainder mainly monocytes.
6. This procedure can be scaled down using Eppendorf tubes to provide MNCs from
50–100 µL blood samples obtained from finger pricks.
7. After cryopreservation, mononuclear cells should be incubated at least overnight
but not longer than 48 h before use.
8. It is necessary to limit division; otherwise cell growth will dilute out unrepaired
damage and give the appearance of repair. A similar difficulty would arise with
activated lymphocytes.
9. Although other concentrations of agarose may be used, those specified are firm
enough to withstand multiple application and removal of coverslips.
10. If cells are suspended in LMP agarose that was made up in their normal culture
medium, they can be treated and incubated for at least several hours. Agarose made
up in PBS is satisfactory if cells are to be used immediately, but damage accumu-lates if the cells are incubated on the slide. It is possible to make up both agarose
layers with medium.
11. A common difficulty is detachment of the bottom agarose layer from the slide.
The most important precaution to avoid this is to ensure that the agarose is added
to a warm slide, so that it spreads completely before it sets. It is also necessary to
be sure to leave the slides long enough over ice for the agarose to set. A final
remedy is to increase the concentration of agarose in the bottom layer (we now
use 1% and 0.7% agarose concentrations, whereas we originally used 0.5% in
both layers).
12. In warm humid weather we have found a problem with damage in controls that we
attribute to condensation on the surface of the agarose. We normally obtain accept-able results by minimizing the time that the coverslip is removed from the slide.
13. In monitoring human populations for levels of background damage, results may
influenced by such factors as the time of day that the sample is taken, or what the
subject has recently eaten (10), exercise (13), smoking (14), and medication. Such
factors must be defined as carefully as possible. It is strongly recommended that
any study should include taking repeat samples from each subject.
14. Example treatments: 0.25–10 Gy of ionizing radiation, cells irradiated on slides
without coverslips; 0.1–2.5 J/m2 UV-C (254 nm) radiation, coverslips need to be
removed and care must be taken to ensure that there are no photosensitisers or
UV-absorbent compounds in the medium (e.g., phenol red); 10–300 mM H2O2,
added in solution with a coverslip on ice for 0–10 min; 50 µM–0.1 U/mL xan-thine–xanthine oxidase, added as a solution with a coverslip and incubated at
37°C for 1 h. 1 mM S-Nitrosoglutathione is added as a solution with a coverslip
to cells embedded in agarose or added to medium of cultured cells and incubated
for 30 min to 24 h.
Comet Assay of DNA Damage 155
15. In addition to single-stranded breaks, the alkaline Comet assay may also detect
alkali-labile sites. The nature of these sites has not been well defined and there is
some controversy whether the conditions used are sufficiently alkaline for their
16. We use electrophoresis buffer at 15°C, but any temperature between 4°C and
room temperature is feasible, provided that the same temperature is used in each
experiment. Again, another “unwinding time” can be chosen, provided that the
same time is used throughout. We find that 40 min gives greatest sensitivity, with-out undue damage in controls. The time should be shortened if it is necessary to
make the assay more robust.
17. Make sure that the shelf of the gel box is completely horizontal, otherwise migra-tion of DNA will be affected by the position of the slide within the gel box.
18. Slides can also be dried and stored indefinitely. Omit the staining step. Remove
the coverslip and leave on the bench overnight. To score the slides, add 100 µL of
PBS and a coverslip. Leave for 2 h at room temperature. Remove the coverslip,
add 35 µL ethidium bromide staining solution, replace the coverslip, and score.
19. Optimum concentration of each enzyme will need to be determined in trials to
ensure that saturating conditions are applied.
20. Addition of inhibitor is essential to see excision breaks with normal human fibro-blasts following treatments such as ultraviolet irradiation. We have not found it
necessary to see strand breakage by reactive oxygen species in human islets of
Langerhans. Freshly isolated human lymphocytes have extremely low deoxyri-bonucleotide pools and show delayed strand rejoining without the need to add
any inhibitor. In this case, breaks represent transient repair intermediates (32).
21. To avoid bias, it is essential to set out in advance rules for accepting or rejecting
Comets for scoring. Our strategy is to accept unless there are specific grounds for
rejection. We reject Comets close to the edge of the slide, Comets grossly out of
focus, and overlapping Comets that cannot readily be separated.
The work on which these protocols have been based was funded in part by
the EC (ENV4-CT95-0174).
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its influence on genotoxicity, in Basic Science in Toxicology (Volans, G. N. [J. S.],
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Comet Assay of DNA Damage 157
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single-strand breaks in human-lymphocytes by low-doses of γ-rays. Int. J. Radiat.
Biol. 68, 563–569.
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Immunochemical Assay for DNA Damage 159
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Measurement of DNA Damage and Repair in Human
White Blood Cells by an Immunochemical Assay
Govert P. van der Schans
1. Introduction
DNA is the most important target molecule for cell killing or the induction
of cellular damage by chemical or physical agents. Cell killing depends on the
type and amount of damage induced. Exposure to physical or chemical agents
can induce a large variety of lesions in DNA: single- and double-stranded
breaks, as well as damage to the bases and sugar residues (not leading to a
break). It is important to be able to quantify the different types of DNA damage
to obtain information about their persistence. The latter may attribute to our
understanding of how the various lesions are involved in cell death and/or muta-tion induction.
Several methods are currently used to quantify single-stranded breaks in
cellular DNA. The immunochemical assay, introduced by us (1,2) for the deter-mination of single-stranded breaks, is based on the application of monoclonal
antibodies directed against single-stranded DNA. In fact it is a combination of
the so-called unwinding assay with an immunochemical quantification of the
single-stranded DNA formed upon unwinding. The principle of the method is
that DNA starts to unwind—under strictly controlled alkaline conditions—at
each (double- and single-) strand break (and at each lesion converted into such
breaks in alkaline medium), resulting in a long stretch of single-stranded DNA
(ssDNA) to which the antibody can bind. The amount of single-stranded DNA
is a measure for the amount of breaks. With this method the amount of damage
induced by ionizing radiation in DNA in cells of human blood can be detected
This chapter represents Part of a European patent request, No. 93201672. 8 (10 June 1993)
entitled: “Method for detecting single-strand breaks in DNA,” by G. P. van der Schans.
160 van der Schans
within 1 h, after doses as low as 0.2 Gy. The precoating of microtiter plates
with anti-ssDNA antibody enables the detection of ssDNA fragments directly
in alkali-treated blood samples. Prelabeling and isolation of the nucleated cells
from the blood is not necessary. Only a few microliters of blood is required and
the assay can be applied simultaneously on a large number of samples.
The method is also applicable to other cell types that can be obtained in
suspension and can be applied in a large variety of studies on induction and
repair of damage, including that on aging (3).
A special application is that on sperm cells (4). Because in these cells the
DNA is rather tightly packed the alkaline treatment conditions have to be modi-fied to release the DNA. Owing to the absence of any repair in these cells an
accumulation of (oxidative) damage occurs, resulting in a background level of
single-stranded breaks corresponding to that induced by 50–200 Gy γ-rays.
2. Materials
2.1. Preparation of DNA Samples (Somatic Cells)
1. Blood collected in evacuated glass tubes containing EDTA.
2. RPMI 1640 culture medium containing 10% fetal calf serum.
3. Cultured mammalian cells.
4. MQ-water: Demineralized water purified further by filtration through a Milli-Q
filter (class 1 according to ISO 3696).
5. 1 M NaOH solution: Dissolve 3.93 g of NaOH (in tablet form) in (by weighing)
98 g of MQ-water (or proportionally other amounts) in a 100-mL medium flask.
Close with two tissues of parafilm covered with a loosely tightened screwcap.
Can be stored at room temperature for 2 mo.
6. 1.3 M NaCl solution: Dissolve 38.0 g of NaCl in MQ-water and fill up to 500 mL.
Can be stored at room temperature for 2 mo.
7. Alkali solution: Add 2.00 mL of 1 M NaOH or 2.50 mL of 1 M NaOH to 103 g of
a 1.3 M NaCl solution (calculated pH 12.3 or 12.4, respectively). Prepare on the
day of use.
8. 0.25 M NaH2PO4 solution: Dissolve 8.63 g of NaH2PO4·H2O in MQ-water and
fill up to 250 mL.
9. Polystyrene cluster tubes, Costar, cat. no. 4408.
10. Polystyrene tubes with cap, 12.4/75 mm, Greiner, no. 120161.
11. 1.5-mL Eppendorf tubes.
12. Pipet tips (white opaque), 1–200 µL, Costar, no. 4862 (see Note 2).
13. Pipet tips (blue), 100–1000 µL, Eppendorf, no. 0030 015.002.
14. Pipets, P100, P200 and P1000, Gilson.
15. 12-Channel pipet, 25–200 µL; Micronic (Macap) cat. no. 200–12.
16. Sonicator/cell disruptor, Ultrasonics, W370 with microtip.
17. Humidified incubator of 37°C.
18. Reaction tube mixers, Vibrofix, IKA, model VF1.
Immunochemical Assay for DNA Damage 161
2.2. Preparation of DNA Samples (Spermatozoa)
1. Spermatozoa, diluted ejaculates delivered in straws containing 5–25 × 106 sperm
cells in 200-µL aliquots, fresh or frozen in liquid nitrogen.
2. 10× Phosphate-buffered saline (PBS) solution: 81.8 g of NaCl, 14.4 g of
Na2HPO4·2H2O, 2.0 g of KH2PO4, and 1.9 g of KCl in 1 L MQ-water. Check the pH;
it should be between 7.0 and 7.4. Can be stored at room temperature for 2 mo.
3. PBS solution: Add to a 10-L polythene vial, with valve, 9 L of MQ-water and 1 L
of 10× PBS, and mix thoroughly.
4. Solution A: 732 µL of 10% Triton X-100 in MQ-water, 879 µL of dithiothreitol of
200 mg/mL MQ-water (freshly prepared), 2.77 mL of 1 M NaOH, and 65.4 mL
of 2 M urea (freshly prepared).
5. 0.25 M NaH2PO4 solution: Dissolve 8.63 g of NaH2PO4·H2O in MQ-water and
fill up to 250 mL.
6. 1.5-mL Eppendorf tubes.
7. Polystyrene round-bottom tubes, 6.5 mL, Greiner, no. 151101.
8. Pipet tips (white opaque), 1–200 µL, Costar, no. 4862 (see Note 2).
9. Pipet tips (blue), 100–1000 µL, Eppendorf, no. 0030 015.002.
10. Pipets, P100, P200, and P1000, Gilson.
11. Sonicator/cell disruptor, Ultrasonics, W370 with microtip.
12. Humidified incubator of 37°C.
2.3. Coating of the Microtiter Plates
1. Tween-20 (polyoxyethylenesorbitan): Sigma cat. no. P-1379.
2. 10× PBS solution: 81.8 g of NaCl, 14.4 g of Na2HPO4.2H2O, 2.0 g of KH2PO4,
and 1.9 g of KCl in 1 L of MQ-water. Check the pH, which should be between 7.0
and 7.4. Can be stored at room temperature for 2 mo.
3. PBS solution: Add to a 10-L polyethylene vial, with valve, 9 L of MQ-water and
1 L of 10× PBS, and mix thoroughly.
4. Na-PBS: Buffered NaCl solution (NPBI bv, Emmer-Compascuum, The Nether-lands, 8.2 g of NaCl, 1.9 g of Na2HPO4‚2H2O, and 0.3 g of NaH2PO4·2H2O per
liter, pH 7.4, 500 mL, sterile).
5. D1B monoclonal antibodies: Dilute D1B monoclonal antibodies prepared and
purified as described elsewhere (1,2), to a concentration of 10 µg/mL in Na-PBS
(see Note 10). Prepare this solution no sooner than 1 h before use.
6. PT: Add to a 25-L polyethylene flask, with valve, 18 L of MQ-water, 2 L of 10×
PBS, and 10 g of Tween-20, and mix thoroughly.
7. Fetal calf serum (FCS): Gibco-BRL, cat. no. 10106–110, stored at –20°C in
500-mL flasks.
8. Heat-inactivated FCS (hiFCS): Thaw a 500-mL flask with FCS in a 37°C water
bath. Divide under sterile conditions (in a laminar flow cabinet) in 30-mL flasks
and/or 50 10-mL polystyrene tubes with cap. Incubate all flasks and tubes for
0.5 h in a 56°C water bath. Place in refrigerator. Can be stored for 2 mo (2–8°C).
9. PT + 5% hiFCS: Dilute hiFCS with PT in a ratio of 1 mL of hiFCS and 19 mL PT.
Prepare this solution no sooner than 1 h before use.
162 van der Schans
10. High-binding polystyrene microtiter plates, Costar, 96-well, cat. no. 3590/9018.
11. Pipet tips (white opaque), 1–200 µL, Costar, no. 4862.
12. Pipet tips (blue), 100–1000 µL, Eppendorf, no. 0030 015.002.
13. Pipets, P100, P200, and P1000, Gilson.
14. 12-Channel pipet, 25–200 µL; Micronic (Macap) cat. no. 200–12.
15. 96-Well dispenser, Transtar 96, adjustable volume, cat. no. 7605 and Transtar
elevator cat. no. 7606, Costar.
16. Plate washer, Skanwasher 300, Skatron.
17. 12-Channel handplate washer, with tube connected to a polypropylene flask
with PT.
18. Plate vibrators, Titertek (Flow) and IKA (model MTS4).
2.4. Sandwich Enzyme-Linked Immunosorbent Assay (ELISA)
1. 1 M NaOH solution: Dissolve 3.93 g of NaOH (in tablet form) in (by weighing)
98 g MQ-water (or proportionally other amounts) in a 100-mL medium flask.
Close with two tissues of parafilm covered with a loosely tightened screwcap.
Can be stored at room temperature for 2 mo.
2. 1.3 M NaCl solution: Dissolve 38.0 g of NaCl in MQ-water and fill up to 500 mL.
Can be stored at room temperature for 2 mo.
3. Alkali-solution: Add 2.00 mL of 1 M NaOH to 103 g of a 1.3 M NaCl solution
(calculated pH 12.3).
4. Tween-20 (polyoxyethylenesorbitan): Sigma cat. no. P-1379.
5. 10× PBS solution: 81.8 g of NaCl, 14.4 g of Na2HPO4.2H2O, 2.0 g of KH2PO4,
and 1.9 g of KCl in 1 L of MQ-water. Check the pH, which should be between 7.0
and 7.4. Can be stored at room temperature for 2 mo.
6. PBS solution: Add to a 10-L polyethylene vial, with valve, 9 L of MQ-water and
1 L of 10× PBS, and mix thoroughly.
7. PT: Add to a 25-L polyethylene flask, with valve, 18 L of MQ-water, 2 L of 10×
PBS, and 10 g of Tween-20, and mix thoroughly.
8. FCS: Gibco-BRL, cat. no. 10106–110, stored at –20°C in 500-mL flasks.
9. hiFCS: Thaw a 500-mL flask with FCS in a 37°C water bath. Divide under sterile
conditions (in a laminar flow cabinet) in 30-mL flasks and/or 50 10-mL polysty-rene tubes with cap. Incubate all flasks and tubes for 0.5 h in a 56°C water bath.
Place in refrigerator. Can be stored for 2 mo (2–8°C).
10. Sodium dodecyl sulfate (SDS): Sigma, cat. no. L-4509.
11. 10% SDS solution: Dissolve 10 g of SDS in 100 mL of MQ-water. Can be stored
at room temperature for 2 mo.
12. PT + 0.05% SDS + 5% hiFCS: Make up solution of PT, hiFCS, and 10% SDS in
the ratio of 1 mL of hiFCS, 19 mL of PT, and 100 µL of 10% SDS. Prepare this
solution no sooner than 1 h before use.
13. D1B-AP conjugate: Dilute D1B-AP conjugate, prepared as described elsewhere
(2), to the optimal prescribed dilution (1000–40,000×; see Notes 4 and 10) in PT
+ 0.05% SDS + 5% hiFCS. Prepare this solution no sooner than 1 h before use.
Immunochemical Assay for DNA Damage 163
14. 0.1  M MgCl2 solution: Dissolve 2.03 g of MgCl2·6H2O in 100 mL of
15. Reaction buffer: Dissolve 5.26 g of diethylamine (DEA) in 5 L of MQ-water and
add 50 mL of 0.1 M MgCl2, adjust pH to 9.8, and divide over ten 500-mL flasks.
Can be stored for 6 mo in the cold room. Before use, raise the temperature to
room temperature. Check the pH (at room temperature) and, if needed, adjust the
pH to 9.8. After use place the flask back in the refrigerator.
16. 4-methylumbelliferyl phosphate (MUP) dilithium salt·3H2O: Boehringer, cat. no.
405 663, check expiration date. Mol wt: 322.1. Store in the dark at –20°C.
17. MUP stock solution: Dissolve the whole contents of a flask of MUP (approx 250 mg
net weight by back weighing) in reaction buffer, pH 9.8, in  clean glassware  at a
concentration of 20 mM (6.44 mg/mL), and divide into 1-mL portions in 1.5-mL
Eppendorf tubes (with cap) and store at –20°C. Can be stored for 6 mo (in the dark).
18. MUP solution: Thaw the required amount of MUP stock solution and dilute 1:100
in reaction buffer, pH 9.8. The remaining MUP stock solution can be placed back
in the freezer, marked with ink pen. Prepare this solution no sooner than 2 h
before use.
19. Pipet tips (white opaque), 1–200 µL, Costar, no. 4862 (see Note 2).
20. Pipet tips (blue), 100–1000 µL, Eppendorf, no. 0030 015.002.
21. Pipets, P100, P200, and P1000, Gilson.
22. 12-Channel pipet, 25–200 µL; Micronic (Macap) cat. no. 200-12.
23. 96-Well dispenser, Transtar 96, adjustable volume, cat. no. 7605 and Transtar
elevator cat. no. 7606, Costar.
24. Plate washer, Skanwasher 300, Skatron.
25. 12-Channel handplate washer, with tube connected to a polypropylene flask
with PT.
26. Sonicator/cell disruptor, Ultrasonics, W370 with microtip.
27. Humidified incubator of 37°C.
28. Fluorescence microtiterplate reader, Cytofluor II (Perseptive Biosystems,
Framingham, MA, USA).
29. Plate vibrators, Titertek (Flow) and IKA (model MTS4).
30. Reaction tube mixers, Vibrofix, IKA, model VF1.
3. Methods
3.1. Preparation of DNA Samples (Somatic Cells)
1. Cells (e.g., human total WBC) in 1 mL of blood are irradiated with 0, 1, 2 or 5 Gy
60Co-rays at 0°C. (Note: Blood may contain infectious particles; wear appropri-ate protective devices.)
2. At yellow light, at 20 + 1°C (see Note 1): 200 µL of alkaline solution of pH 12.3
or 12.4 is added quickly, but avoiding shaking afterwards, to 30 µL of 10× diluted
blood (diluted in RPMI 1640 + 10% FCS, or in PBS) on the bottom in a cluster
tube (in 96-wells tray;  see Note 3). The pH chosen depends on the extent of
unwinding that is required.
164 van der Schans
3. Neutralize after 6 min with 35 µL of 250 mM NaH2PO4, added to the bottom of
the tube.
4. Immediately after neutralization the solution is sonicated at 20°C (set at 2.5,
duration 1 s).
5. The samples are used immediately for the ELISA, or stored at 4°C (approx 1 wk)
or frozen (–20°C). Standing in cluster tubes at 4°C results in slow disappearance
of ssDNA from the solution (adsorption to the wall?). Therefore, at later use,
warm first to room temperature and sonicate before dilution.
6. The DNA samples (blood, other body cells, cultured cells) as such can be used in
the sandwich ELISA. When samples are to be analyzed on the same day then
store in the refrigerator (at 4°C) up to approx 1 h before use; otherwise, store
samples at –20°C.
3.2. Preparation of DNA Samples (Spermatozoa)
1. Straws with frozen sperm are thawed at 37°C and sperm are pressed out in a
1.5-mL Eppendorf tube.
2. Immediately thereafter the cell suspension is mixed by pipetting and 20-µL
aliquots are pipeted in triplicate into 1.5-mL Eppendorf tubes and placed on ice.
3. Within 2.5 h these samples are diluted 40× with PBS at 20°C.
4. In yellow light, at 20 ± 1°C, immediately after mixing by pipetting, 60-µL aliquots
of the diluted samples are transferred in duplicate to 6.5-mL polystyrene round-bottom tubes.
5. 415 µL of solution A is added by running it along the wall of the tube and after
7 min (yellow light, 20°C) neutralized with 150 µL of 250 mM NaH2PO4, imme-diately followed by a brief sonication.
6. The samples are used immediately in the ELISA, or stored at 4°C (approx 1 wk).
Standing in cluster tubes at 4°C results in slow disappearance of ssDNA from the
solution (adsorption to the wall?). Therefore, at later use, warm first to room
temperature and sonicate before dilution.
3.3. Coating of the Microtiter Plates
1. Add, to the 96-wells dispenser, 100 µL of the D1B-solution (10 µg of D1B/mL of
Na-PBS) to a high-binding polystyrene microtiter plate.
2. Vibrate the plate for 10 min at room temperature (maximal 2 × 8 plates/vibrator).
3. Wash the plate 3× with PT buffer with a plate washer (as indicated in the “Wash
Program Sheet”) or with a hand plate washer.
4. Add with the 96-well dispenser 100 µL PT + 5% hiFCS per well.
5. Vibrate the plate for 10 min at room temperature (maximal 2 × 8 plates/vibrator).
6. Wash the plate once with PT buffer with the hand plate washer.
7. Stack the plates in groups of 10. Pack in aluminum foil and store at –20°C. (Write
date on aluminum foil.) Coated plates can be stored 2 mo.
8. Before use place plates beside each other open on the table to equilibrate to room
temperature (this equilibration process normally takes 10–60 min).
Immunochemical Assay for DNA Damage 165
3.4. Sandwich ELISA
1. Make 100% ssDNA samples from an aliquot of the DNA samples to be assayed:
Add, at room temperature, 25 µL of DNA sample to a cluster tube, and add
200 µL of a solution of 1.3 M NaCl + 0.02 M NaOH, pH 12.3. Shake for 1 s on a
reaction tube mixer or sonicate (set at 2.5, duration 1 s), and neutralize with
25 µL of 0.25 M NaH2PO4. In the case of 100% ssDNA samples of sperm, these
are always sonicated before neutralization.
2. Add to the first row of a D1B-coated “high-binding” polystyrene plate (with a
12-channel pipet) 120 µLPT and in all other rows 70 µL.
3. Place the samples in rows of 12, including the 100% ssDNA samples, according
to the scheme prepared in advance.
4. Add of each sample (with a 12-channel pipet) 20 µL in a well of the first row of the
plate. In the case of sperm DNA samples add the 20-µL aliquots with a single pipet.
5. Make 1:1 serial dilutions until the last row.
6. Add to wells 5–8 of the last row of each plate 140 pg of single-stranded DNA
(intended to correct for plate-to-plate variations).
7. Incubate at room temperature under continuous vibration for at least 5 min (see
Note 7, 2 × 8 plates/vibrator).
8. Wash 3× with PT buffer (hand wash apparatus or plate washer).
9. Add 100 µL of D1B-AP solution/well (with dispenser, use filter plate up to 200×,
or with 12-channel pipet; see Note 5).
10. Incubate again at room temperature under continuous vibration for at least 5 min
(up to 2 × 8 plates/vibrator).
11. Wash 3× with PT buffer (hand wash apparatus or plate washer). The last wash
should be extensive and done using the hand wash apparatus (see Notes 8 and 9).
12. Add 100 µL of MUP solution/well (with dispenser or with 12-channel pipet).
13. Incubate for 1 h at 37°C in a humidified incubator (cover on plate, do not stack plates).
14. Read fluorescence in all wells (Cytofluore II, gain 45) and transfer data to a PC
with lotus or excel spreadsheet (see Note 6).
3.5. Calculation of ssDNA Percentage
Provided limited amounts of DNA are used a linear relationship is obtained
between the input amount of DNA and the level of fluorescence, both for com-pletely and for partially single-stranded DNA. To calculate the fraction of
single-strandedness in a particular sample, the ratio of the fluorescence of the
DNA dilutions to that of the corresponding completely single-stranded DNA
dilutions, both corrected for background fluorescence (fluorescence in wells
without DNA), is divided by a factor of 10 (because of the predilution of the
completely single-stranded DNA sample) according to the following formula:
(flsample – flbackground)
% single-strandedness = —————————————— × 100%
10 × (fl100% ss – flbackground)
166 van der Schans
The calculations are carried out only for those dilutions for which a linear
relationship is observed between the amount of DNA in the wells and the level
of fluorescence. In daily practice, this holds for fluorescence values <1500. On
the other hand values below 500 fluorescence units become less accurate.
Therefore, the percentage of single-strandedness calculated for the highest
amounts of DNA in the well with a fluorescence <1500 and for the lowest
amounts of DNA in the well with a fluorescence >500. The average of the two
data is considered as the most realistic one.
If none of the fluorescence values in a serial dilution falls within the required
range then the computer program takes the value of the second dilution in the
case of low fluorescence values and that of the sixth dilution in the case of high
fluorescence values.
4. Notes
1. The use of an air-conditioned room at 20°C with yellow light is recommended for
the alkaline treatment of the samples.
2. Use of pipet tips is recommended because of high precision in pipetting slightly
viscous solutions.
3. If the use of cluster tubes causes difficulties (bad mixing), then, as an alternative,
polystyrene round-bottom tubes (volume 6.5 mL) should be used. In that case
200 µL alkaline solution is added on top along the wall, to each tube separately).
4. The optimal dilution of a new batch of D1B-AP conjugate is assessed by compar-ing the optimal dilution with that of an old batch. The optimal dilution is the one
that results in an almost equal fluorescence as obtained with the optimal dilution
of the old batch. The optimal dilution is a compromise between an optimal fluo-rescence with respect to the range of the microtiter plate reader and the costs of
the D1B-AP conjugate.
5. Owing to the fact that only a small part of the ssDNA fragment binds to the walls
of the immobilized anti-ssDNA antibodies, enough antibody binding sites remain
available to bind the same antibodies conjugated with alkaline phosphatase.
6. The computer program to be used depends on the purpose of the experiment.
When samples are tested with a variable number of cells then it is preferred to
have the 100% ssDNA samples on the same plate. The computer program directly
calculates the % ssDNA of these samples.
When the 100% ssDNA samples are not present on the same plate, then the
corresponding computer program uses a fixed value for the 100% ssDNA
samples, corrected only for possible plate-to-plate variations. The resulting per-centages of ssDNA should be corrected for the real fluorescence values of the
100% ssDNA samples.
7. The time of incubation to adsorb the ssDNA to the D1B antibody is at least 5 min.
The incubation period can be extended up to half an hour, but longer periods
should not be used.
Immunochemical Assay for DNA Damage 167
8. Sometimes the use of the plate washer, and/or the 96-well dispenser presents
problems owing to (1) contamination of the washer with high amounts of
D1B-AP and (2) contamination or leakage of the dispenser. In that case, the solu-tions should be added manually, with the 12-channel pipet, and the plates should
be washed with the hand wash apparatus. The latter is preferred, at least for the
last wash step before adding the reaction buffer.
9. Historically, before adding substrate in reaction buffer, the plates were washed
once with 0.1 M diethanolamine, pH 9.8. Later on this step was omitted, as higher
fluorescence values without substantial increase of the background fluorescence
were obtained.
10. Both the required antibody, D1B, as well as its conjugate, D1B-AP, are made
available on request to the author at a cost necessary to cover production, devel-opment, and shipment.
1. Van der Schans, G. P., Van Loon, A. A. W. M., Groenendijk, R. H., and Baan, R. A.
(1989) Detection of DNA damage in cells exposed to ionizing radiation by use of
anti-single-stranded-DNA monoclonal antibody. Int. J. Radiat. Biol. 55, 747–760.
2. Timmerman, A. J., Mars-Groenendijk, R. H., Van der Schans, G. P., and Baan, R. A.
(1995) A modified immunochemical assay for the detection of DNA damage in
human white blood cells. Mutat. Res. 334, 347–356.
3. King, C. M., Bristow-Craig, H. E., Gillespie, E. S., and Barnett, Y. A. (1995) In vivo
antioxidant status, DNA damage, mutation and DNA repair capacity in cultured
lymphocytes from healthy 75- to 80-year-old humans. Mutat. Res. 377, 137–147.
4. Van der Schans, G. P. (1993) Method for detecting single-strand breaks in DNA.
European patent request, no. 93201672.8, 10 June 1993.
8-oxoguanine Levels in Nuclear DNA 171
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Measurement of 8-Oxo-deoxyguanosine in
Lymphocytes, Cultured Cells, and Tissue Samples
by HPLC with Electrochemical Detection
Sharon G. Wood, Catherine M. Gedik, Nicholas J. Vaughan,
and Andrew R. Collins
1. Introduction
8-Oxoguanine is one of the most studied base oxidation products found in
DNA. It has potential biological significance, because if present in DNA that is
replicating, it can lead to incorporation of adenine rather than cytosine in the
daughter strand. Thus it is considered as a premutagenic lesion. It occurs as a
result of attack by reactive oxygen species released during the inflammatory
response, and in small but significant amounts during normal respiration. The
hydroxyl (·OH) radical (arising from H2O2 by the transition metal ion-catalyzed
Fenton reaction within the nucleus) is most likely responsible for the formation
of 8-oxoguanine. Analytical methods—gas chromatography with mass spectro-metric detection (GC–MS) and high-performance liquid chromatography (HPLC)
—were developed for quantitation of oxidized bases produced in experimental
studies of radiation and chemical damage to DNA, and these methods were
naturally also applied to the measurement of background levels of oxidized bases
in cellular DNA (1). With GC–MS, very high levels of 8-oxoguanine have been
reported, typically between 10 and 100 for every 105 normal guanines. It has
recently been recognized that spurious oxidation of DNA readily occurs during
isolation and hydrolysis of DNA, and derivatization of the bases for analysis.
HPLC, normally applied to measurement of the nucleoside, 8-oxo-deoxyguano-sine (8-oxo-dG), has generally given values below those obtained with GC–MS;
but with HPLC, too, oxidation artefacts have been identified. Currently much
effort is going into reconciling the different approaches to the measurement of
172 Wood et al.
oxidative DNA damage (GC–MS, HPLC, and various repair endonuclease-based
DNA breakage assays), with some success. Most researchers in the field agree
that a realistic figure for the background level of 8-oxo-dG in normal human
cells is around 1 per 105 dGs, or even less.
One of the most popular hypotheses of aging is that oxidative damage to
biomolecules accumulates, leading ultimately to dysfunction of proteins, mem-branes, and DNA replication and repair machinery. 8-Oxo-dG in DNA seems a
reasonable and convenient biomarker for the process of aging. Here we describe
the method that we have developed for the analysis of 8-oxo-dG by HPLC with
electrochemical detection (HPLC–ECD) in cultured cells, lymphocytes, and
tissues (though in human studies, the availability of tissue samples is obviously
very limited). We have introduced modifications to limit oxidation in vitro—
inclusion of various antioxidants and chelators during the preparation, storage,
and hydrolysis of DNA, lowering the temperature and time of incubation with
proteinase K, optimization of sample preparation for storage (2). The proce-dure described here gives the lowest levels of 8-oxo-dG in samples of rat liver.
2. Materials
1. Homogenization buffer: 10 mM Tris-HCl, 0.4  M NaCl, 5 mM deferoxamine
mesylate (DF; Sigma, cat. no. D9533). Prepare in HPLC grade H2O (Rathburn
Chemicals, Walkerburn, Scotland). Adjust to pH 8.0 with dilute HCl. Make up to
1 L with water and freeze at –20°C as aliquots; cover containers in aluminum foil
to protect from light (DF is unstable and light-sensitive, see Note 1). Just before
use, thaw and add Triton X-100 (Sigma, cat. no. T9284) to 0.5%. Triton X-100 is
viscous; to measure volume accurately, warm the Triton and use a plastic pipettor
tip with the end cut off. Mix the solution very thoroughly.
2. 40 mM Tris-HCl buffer: Prepare in HPLC grade H2O, adjusting to pH 8.5 with
dilute HCl. Store as aliquots at –20°C.
3. Ribonuclease buffer: 10 mM Tris-HCl, 0.4 M NaCl; prepare in HPLC grade H2O,
adjusting to pH 8.0 with dilute HCl. Store at –20°C.
4. RNase T1 (ICN, cat. no. 101079) from Aspergillus oryzae: Powder, stored at –20°C.
Add ribonuclease buffer to powder to give 103 U/mL. Aliquot into micro-centrifuge tubes, put in 80°C water bath for 15 min (to destroy any contaminating
DNases), and allow to cool slowly to room temperature. Store aliquots at –20°C
(see Note 2).
5. RNase IIIA (Sigma, cat. no. R5125) from bovine pancreas: Powder, stored at –20°C.
Prepare solution of 1 mg/mL in ribonuclease buffer. Aliquot into microcentrifuge
tubes, put in 80°C water bath for 15 min, and allow to cool slowly to room tem-perature. Store at –20°C (see Note 2).
6. Proteinase K (Boehringer Mannheim, cat. no. 1373 196) from  Tritirachium
album: Supplied as solution ready for use. Store at 4°C.
7. Denley Spiramix 5 roller mixer.
8. Quartz Suprasil 150-µL microcuvet (Hellma, cat. no. 105.201-QS).
8-oxoguanine Levels in Nuclear DNA 173
9. DNase 1 (Boehringer Mannheim, cat. no. 104132) from bovine pancreas: Store
at –20°C. Supplied as a solid; dissolve 20,000 U in 4 mL of 40 mM Tris-HCl, pH
8.5 (i.e., 5 U/µL) and store as aliquots at –20°C (see Note 2).
10. Alkaline phosphatase from calf intestine (Boehringer Mannheim, cat. no.
1097075). Supplied at about 20 U/µL (varies between batches) in triethanola-mine buffer. This stock should be kept at 4°C. Dilute to 0.25 U/µL with 40 mM
Tris-HCl, pH 8.5, and store aliquots at –20°C. Do not use the dephosphoryla-tion buffer supplied because it contains EDTA which would remove the Mg
2+ ions required for DNA hydrolysis (see Note 2).
11. Phosphodiesterase II from calf spleen (Boehringer Mannheim, cat. no. 108251),
supplied as 2 mg/mL suspension in ammonium sulfate solution. This stock should
be stored at 4°C. Its activity is about 2 U/mg, that is, 4 U/mL; however, there is a
20% loss of activity in 6 mo. Dilute to 0.25 U/mL with 40 mM Tris-HCl, pH 8.5,
and store aliquots at –20°C (see Note 2).
12. Phosphodiesterase I from  Crotalus durissus  (Boehringer Mannheim, cat. no.
108260), supplied in 50% glycerol, 1 mg in 0.5 mL, approx 1.5 U/mg or 3 U/mL.
Store at 4°C. For use, dilute 3× in 40 mM Tris-HCl, pH 8.5 (dispense from the
stock solution using sterile conditions).
13. Syringe filters with 0.2 mm pore size (Whatman, cat. no. 6777-0402).
14. HPLC hardware (isocratic system): We use Gilson 306 pump 10 washed self cen-tering (wsc) pump head; Gilson 805 manometric module; Gynkotek GINA 50
cooling autosampler at 5°C.
15. HPLC columns: Guard column, Supelco pellicular 5 µm, 2 cm; analytical col-umn, Capital Analytical ODS Apex 3 µm, 4.6 × 150 mm.
16. HPLC mobile phase: 50 mM Potassium phosphate, 8% methanol, pH 5.5. Mix
96 mL of 1 M KH2PO4 (HiPerSolv grade for HPLC; BDH, cat. no. 153184 U)
and 4 mL of 1 M K2HPO4 (Aristar grade; BDH, cat. no. 45233) with HPLC grade
H2O to 1.84 L. Fine adjust pH with dilute orthophosphoric acid and add 160 mL
of HPLC grade methanol to 2 L final volume (Rathburn Chemicals). Filter
through a 0.45 mm nylon filter under vacuum, to remove any particulate material
and to degas. Store at 4°C. Replace weekly to avoid bacterial growth.
17. Gilson Holochrome UV detector for measuring dG.
18. ESA Coulochem II electrochemical coulometric detector with a 5021 condition-ing cell and a high sensitivity 5011 analytical cell. The measurement of the low
levels of background 8-oxo-dG requires the use of coulometric detection as
opposed to the less sensitive amperometric method.
19. Data collection and analysis is by Gynkotek Chromeleon software.
Note: Mention of brand names does not imply endorsement of these prod-ucts in preference to other similar materials.
3. Methods
Total DNA isolated from eukaryotic cells includes mitochondrial DNA.
Although this is a minor component compared with the nuclear DNA, it may
174 Wood et al.
have a much higher content of oxidized bases (since the mitochondria are the
site of production of reactive oxygen species), and so might lead to a signifi-cant overestimation of nuclear DNA damage. The best course, therefore, is to
isolate nuclei first. This, in addition, avoids the possibility of DNA oxidation
arising from cytoplasmic contamination with peroxisomes, or with free iron or
copper ions, that can catalyze the Fenton reaction. Liver cytoplasm is particu-larly likely to provide an oxidative environment.
3.1. Isolation of DNA
1a. Isolation of nuclei from cultured cells: Suspend 20 × 106 cells in 3 mL of homog-enization buffer with Triton X-100 in a 15-mL centrifuge tube, leave for 5 min on
ice, and centrifuge 10 min at 1200g, 4°C. Wash with 5 mL of Triton-free homog-enization buffer, centrifuge for 5 min at 1200g, 4°C, and disperse the pellet well
in 1 mL of Triton-free buffer. Measure volume and make up to 4.7 mL with buffer.
1b. Isolation of nuclei from human lymphocytes: Suspend 18  × 106 lymphocytes
(expected yield from 30 mL of blood, isolated by standard density gradient sedi-mentation method) in 5 mL of homogenization buffer with Triton X-100 and
continue as described in step 1a above.
1c. Isolation of nuclei from tissue samples (e.g., liver): Weigh out approx 150 mg of
liver (fresh, or frozen under liquid N2) as quickly as possible to avoid exposure to
air. Add to 4 mL of ice-cold homogenization buffer (with Triton X-100). Transfer
to a glass homogenizer tube (Potter–Elvehjen) on ice, and break up the tissue
with about six strokes of the homogenizer over 1 min. Start at slow speed, turn up
to full speed, and slow down before removing pestle from tube. Place homoge-nate in a 15-mL tube and centrifuge for 10 min at 1200g, 4°C. Discard superna-tant, agitate pellet by shaking, add 5 mL of Triton-free homogenization buffer to
wash the pellet, and centrifuge for 5 min at 1200g, 4°C. Discard supernatant, and
resuspend the pellet well in 1 mL of Triton-free homogenization buffer by shak-ing. Measure volume and make up to 4.7 mL with buffer.
2. Add 10% sodium dodecyl sulfate (SDS) in HPLC grade H2O to the nuclear suspen-sion to give a final concentration of 0.6%. The pellet must be well dispersed at
this stage; otherwise SDS will not lyse all the nuclei. Gently invert tube several
times to mix. Incubate for 10 min at 37°C. Add 200 µL of RNase IIIA and 10 µL of
RNase T1. Gently invert 20× to mix, and with a wide-opening pipet aspirate up and
down twice. Incubate for 30 min at 37°C. Add 1 mg of proteinase K (usually about
70 µL but varies from batch to batch). Mix by aspirating gently twice up and down
with a pipet. Incubate for 30 min at 37°C. Cool to room temperature.
3. Transfer the reaction mix to a stoppered glass tube (i.e., resistant to chloroform).
Add an equal volume of chloroform/isoamyl alcohol (24:1). Shake vigorously
for about 15 s to mix. Centrifuge in a glass centrifuge tube for 10 min at 2400g,
room temperature, with no brake. Collect the upper aqueous layer, taking care
not to disturb the cloudy interface between the two phases. If the interface is
solid then carefully pour off the top layer; however, if interface is not com-
8-oxoguanine Levels in Nuclear DNA 175
pacted then it is best to slowly pipet off the upper phase. The condition of the
interface varies with the type of cells used.
4. Repeat step 3 with the aqueous layer.
5. Transfer aqueous layer to a 15-mL centrifuge tube and measure volume. Add
Y mL of 6 M NaCl where Y = 0.311 × measured volume. Vortex-mix immediately
for 10 sec. Centrifuge for 10 min at 2000g, room temperature.
6. Carefully decant supernatant into a 50-mL tube, discarding the pellet. Cool the
supernatant on ice for 15 min. Add 2 vol of ethanol (at –20°C). Invert gently to mix.
Leave on ice for 10 min to aid precipitation of DNA. At this stage some of the DNA
may be clear and gelatinous, occupying a substantial volume at the bottom of the
tube. The rest of the DNA may appear white and floating in the solution.
7. Remove as much ethanol as possible. Wash with 20 mL of ice-cold 70% ethanol
3×, removing the ethanol by aspiration. The DNA will become more compact and
white in color after the first wash.
8. Pick up the DNA pellet on a plastic pipettor tip and transfer to a microcentrifuge
tube. Remove as much ethanol as possible with the pipettor tip. Dry DNA under
a stream of N2; this will take about 5 min. If several samples are processed
together, it is useful to have a manifold outlet for the N2 supply.
9. Add 800 µL of 40 mM Tris-HCl, pH 8.5. Pass N2 over the sample for 1 min and
then seal the top with Nesco Film (Para Film). Slowly turn on a roller mixer at
4°C overnight in the dark. Then leave for 2 h at 37°C to ensure that the DNA is
completely dissolved. Dilute a 10 µL sample to 150 µL and read optical absor-bance at 260 nm and 280 nm in a microcuvet; the ratio of absorbance 260/280 nm
is a measure of purity of DNA and should be around 1.8–1.9 (pure DNA is
1.9–2.0). A more extensive purification would give even cleaner DNA but at the
risk of further oxidation. Absorbance at 260 nm is used to calculate the approxi-mate yield of DNA; 1 AU is equivalent to 50  µg/mL of double strand DNA.
Generally 150–250 µg of DNA are obtained by our purification procedure.
10. Store the DNA solution at –80°C under N2 gas until required.
3.2. Hydrolysis of DNA
Our method is based on that of Richter et al. (3) with the exception that we
find 10× lower amount of phosphodiesterase I to be just as effective. DNA is
hydrolyzed to its constituent nucleosides.
1. The volume containing 75 µg of DNA is calculated for the solution prepared in
Subheading 3.1., step 9 above. Allowing for enzymes and MgCl2, calculate the
volume of buffer required for a final volume of 500 µL and place this amount in a
microcentrifuge tube on ice. Add the following:
a. 3 µL of DNase (5 U/µL)
b. 3 µL of alkaline phosphatase (0.25 U/µL)
c. 3 µL of phosphodiesterase II (0.25 U/mL)
d. 3.8 µL of phosphodiesterase I (1 U/mL)
e. 10 µL of 0.5 M MgCl2
176 Wood et al.
Add the DNA sample and vortex-mix briefly. Incubate for 2 h at 37°C.
2. Filter through a 0.2  µm syringe-filter. Dispense the filtrate into three vials
(120 µL each). In a fourth vial, place 108 µL of filtrate and 12 µL of standard
10 nM 8-oxo-dG and mix.
3.3. HPLC Analysis
A schematic diagram of the HPLC system is shown in Fig. 1.
1. Isocratic system; mobile phase 50 mM potassium phosphate, 8% methanol,
pH 5.5; flow rate 0.5 mL/min; injection volume 100 µL.
Electrochemical detection; 5021 conditioning cell set at 100 mV, 5011 ana-lytical channel 1 set at 150 mV, 5 s filter time, 5 nA full scale (range), 0.1 V
output, 0% offset. 8-Oxo-dG is detected by channel 2 which is set at 400 mV, 5 s
filter time, 1 nA full scale (range), 0.1 V output, 0% offset.
UV detection of dG at 254 nm, with sensitivity set at 0.2 AU.
2. Detection of the very low levels of 8-oxo-dG in normal samples (Fig. 2) requires
certain precautions to reduce background noise to a minimum.
• Prepare new mobile phase buffer weekly.
•Wash out the reservoir weekly with methanol.
•When not in use, purge the mobile phase with helium.
3. Standards: standards of dG and 8-oxo-dG are run separately. This is important
because dG inevitably contains a low level of 8-oxo-dG. When making up stan-dard solutions, do not rely on weighing, or on the information on the label.
Calculate the concentration from the extinction coefficients.  At 1 mM, dG
Fig. 1. A schematic diagram of the HPLC system.
8-oxoguanine Levels in Nuclear DNA 177
and 8-oxo-dG have an absorbance of 13AU (254 nm) and 12.3AU (245 nm),
respectively. Standard curves are run on each day, with dG at 50, 100, and 150 mM
and 8-oxo-dG at 0.25, 0.5, 1, and 2 nM.
4. Do not leave samples for longer than 12 h at 5°C in the autosampler; there is
some evidence of oxidation after this time.
5. Check for carryover of 8-oxo-dG by inserting a 0.25 nM 8-oxo-dG standard after
each set of quadruplicate samples. Check that the peak area is constant through-out the run.
6. The concentration of dG is estimated from the UV peak; 8-oxo-dG from the elec-trochemical signal at 400 mV. Results are usually expressed as the number of
residues of 8-oxo-dG per 105 dG.
4. Notes
1. As DF is light sensitive, cover tubes with foil, keep lid on water bath, etc. Also
DF is unstable at room temperature and 4°C, so the solid and buffers containing
DF should be stored at –20°C. The buffers should be freshly thawed before use
and surplus discarded.
2. Do not attempt to economize by refreezing working solutions of enzymes and
saving them for use at a later date. Be ruthless and discard unused solutions.
We are grateful for financial support from the Ministry of Agriculture, Fish-eries and Food and the Scottish Office Agriculture, Environment and Fisheries
Fig. 2. ECD and UV traces for rat liver nuclear DNA with a typically low back-ground level of 8-oxo-dG.
178 Wood et al.
1. Collins, A., Cadet, J., Epe, B., and Gedik, C. (1997) Problems in the measurement
of 8-oxoguanine in human DNA. Carcinogenesis 18, 1833–1386.
2. Gedik, C. M., Wood, S. G., and Collins, A. R. (1998) Measuring oxidative damage
to DNA; HPLC and the comet assay compared. Free Radic. Res., 29, 609–615.
3. Richter, C., Park, J.-W., and Ames, B. N. (1988) Normal oxidative damage to
mitochondrial and nuclear DNA is extensive.  Proc. Natl. Acad. Sci. USA 85,
Mutation and the Aging Process 179
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Mutation and the Aging Process
Mutant Frequency at the HPRT Gene Locus
as a Function of Age in Humans
Yvonne A. Barnett and Christopher R. Barnett
1. Introduction
Aging is a complex, biological process that is contributed to by intrinsic
(genetic) and extrinsic (nutrition, infectious agents, xenobiotic exposure, etc.)
factors (1). Several decades ago it was first proposed that instabilities in the
organization and expression of the genetic material was likely to be involved in
the aging process (reviewed in ref. 2). Indeed, since that time much experimen-tal evidence has been published that details increases in DNA damage (3–8)
and mutation (8,9–15) in various cells and tissues with age in humans.
Organisms are continuously exposed to a variety of extrinsic biological,
chemical, and physical factors that may alter the structure and therefore have
the potential to modify in vivo the function of a wide range of biomolecules,
including DNA (16–20). If modifications to the structure of DNA are not rec-ognized and removed/repaired, then mutations may result. Mutations in essen-tial genes, in association with the age-related alterations in proteins and lipids,
may result in the degradation of structural elements within the cells, tissues,
and organs of the body, leading to a decline in biological function and eventu-ally to disease and death (1,20,21). There has been a large numerical growth in
the number of older people around the world, due in large part to improve-ments in environmental conditions such as nutrition, housing, sanitation, and
medical and social services. In industrialized countries with low fertility levels
this has resulted in large gains in median population ages. In the 1900s the
maximum life expectancy was around 47 yr, but now this has risen to a mean of
approx 75 yr (22). Further, due in large part to improvements in environmental
180 Barnett and Barnett
conditions such as nutrition, housing, sanitation, and medical and social ser-vices, attempts at increasing the average life expectancy and the quality of life
in the elderly can be achieved only by slowing down the molecular processes
underlying aging. To facilitate such intervention, the factors that cause aging at
the cellular level must be identified and understood, including presumably those
agents that lead to DNA damage and increase the likelihood of mutation.
The detection of in vivo somatic mutations has enhanced our knowledge of
the causes and mechanisms of mutagenesis in somatic cells (23). The ease of
collection of peripheral blood cells together with the presence of intact phase I
and phase II xenobiotic metabolizing enzyme systems within these cells has
led to the development of a number of bioassays for the detection and
quantitation of mutant frequency including the determination of hemoglobin
variants or glycophorin A variants in erythrocytes and the hypoxanthine
phosphoribosyl transferase (HPRT) clonal assay in lymphocytes (these and others
are reviewed in ref. 24).
HPRT is encoded by a gene that spans 44 kb containing 9 exons and is
located on the long arm of the X chromosome (Xq26). It is a constitutively
expressed, nonessential enzyme that functions in the purine salvage pathway to
convert hypoxanthine and guanine to their respective 5′ monophosphate nucleo-sides. In addition, HPRT can utilize a number of base analogs such as 6-thiogua-nine (6–TG) to produce the corresponding ribonucleoside monophosphates.
Cells harboring mutation in the HPRT gene are able to survive in the presence
of 6-TG, whereas wild-type cells will accumulate the highly cytotoxic 6-TG-monophosphate and die. This differential sensitivity to the cytotoxic potential
of 6-TG between HPRT+ and HPRT– cells forms the basis of assays that deter-mine the frequency of HPRT– cells (6-TG resistant) within a mixed HPRT+/
HPRT– cell population.
The most frequently used method to quantitate background mutant frequency
at the HPRT gene locus is a clonal assay. Essentially, lymphocytes are incu-bated in the absence (non-selective conditions) or presence (selective condi-tions) of 6-TG in 96-well microtiter plates for approx 14 d, after which time
wells are scored according to the presence (positive) or absence (negative) of
clones. The zero form of the Poisson distribution (P0) is then used to determine
the cloning efficiency (CE) in the absence and presence of the selection agent,
and the mutant frequency (MF/106 cells) calculated. Although the clonal assay
can be demanding of time, finances, and resources it has the advantage that the
clones can be expanded for determination of mutant status and the mutational
spectrum analyzed (25,26). The remainder of this chapter provides a thorough
description of the clonal assay to quantitate HPRT mutant frequency within
cultured peripheral blood derived human lymphocytes.
Other methods can be used to detect HPRT– lymphocytes. An autoradio-grahic technique, based on the ability of lymphocytes to incorporate [3H]thy-
Mutation and the Aging Process 181
midine into their DNA, following a short period of stimulation with phytohe-magglutinin (PHA), in the presence or absence of the selective agent 6-TG, can
be used to determine frequency of 6-TG-resistant T lymphocytes in samples of
peripheral blood (27). Recently a variation of the autoradiographic assay has
been reported, an immunohistochemical method that enables variant frequency
to be calculated  (28). Using this technique lymphocytes are cultured in the
presence of 5′-bromodeoxyuridine (BrdU), in the presence or absence of the
selective agent 6-TG. The frequency of viable cells in selective and nonselec-tive conditions can then be determined by fluorescence microscopy following
labeling of the cells with a suitably tagged anti-BrdU antibody. The variant
frequency is then calculated, although it is found to be higher than mutant fre-quency determined using the clonal assay, because of the detection of a high
frequency of phenocopies in this assay (due to the detection of cycling HPRT–
lymphocytes). Both of these nonclonal techniques have the advantage of being
relatively simple, rapid, and inexpensive. However, they may overestimate
mutant frequency and have the ability to verify phenotype, genotype, or clonal
2. Materials
1. Lymphocyte separation medium (LSM, ICN Flow,Wycombe, Buckingham,
2. 25-mL Sterile universal containers (Sterilin, Stone, Staffordshire, England).
3. Graduated, sterile tissue culture pipets (Sterilin, Stone, Staffordshire, England).
4. Autoclaved glass Pasteur pipets.
5. RPMI 1640 — Dutch Modification (Imperial Laboratories, Smeaton Rd., West
Portway, Andover, Hampshire, England).
6. Foetal calf serum (FCS; Gibco-BRL, Life Technologies, Renfrew Rd., Paisley,
Scotland), heat-inactivated (56°C/30 min).
7. Hybridoma medium, serum-free medium (HL-1, BioWhittaker UK, Ashville
Way, Wokingham, Berkshire, England).
8. Phytohemagglutinin, HA15 (Murex Diagnostics, Murex Biothech Ltd., Centra
Rd., Dartford, England). Reconstitute each bottle of freeze-dried PHA by adding
5 mL of sterile distilled water using a sterile disposable hypodermic syringe.
Wipe cap with alcohol, then pierce the center of the rubber plug, holding the
syringe in a vertical position.Transfer the reconstituted PHA to a sterile universal
and store at 4°C for a maximum of 1 mo.
9. 6-TG (available as 2-amino-6-mercaptopurine, Sigma). Dissolve 3.34 mg of 6-TG
in 0.5% sodium carbonate (0.5g/100 mL). Cover the container with tin foil (6-TG
is photosensitive) and store at 4°C for a maximum of 1 mo.
10. Human recombinant interleukin-2 (rIL-2), (Chiron UK, Salamander Quay West,
Hareford, Middlesex). rIL-2 is delivered from the supplier in a vial containing
1.2 mg of freeze-dried powder. Reconstitute the contents of the vial with 1.2 mL
of sterile distilled water to give 1 mg/mL containing 1.8 × 107 U/mL of rIL-2.
Stock rIL-2 should be stored frozen at a concentration of 1 × 105 U/mL (add 1.2 mL
182 Barnett and Barnett
of reconstituted rIL-2 to 214.8 mL of RPMI + 10% FCS), aliquot into 1.5-mL
volumes in sterile Eppendorf tubes, and store at –20°C until required. Each batch
of rIL-2 must be tested to establish the most suitable concentration for use in the
cloning assay (see Note 1).
11. Feeder cells, RJK (Epstein–Barr virus [EBV]-transformed B-lymphoblastoid)
cells that carry complete HPRT deletion are used as feeder cells in the cloning
assay. RJK cells should be cultured in RPMI 1640 (Gibco-BRL), 10% heat-inac-tivated FCS, 100 U/mL of penicillin, 100 mg/mL streptomycin, and 0.2 mg/mL
of sodium pyruvate. Irradiate (40 Gy) exponentially growing RJK cells prior to
their use in a cloning assay.
12. Plastic reagent reservoirs for multipipets (Sigma, Fancy Rd., Poole, Dorset,
13. 96-well, flat-bottomed microtiter plates (Nunc, Rosklide, Denmark).
14. Pyruvic acid — sodium salt (Sigma, Fancy Rd., Poole, Dorset, England).
15. Penicillin/streptomycin (1000 U/mL and 1000 mg/mL) (Gibco-BRL, Life Tech-nologies, Renfrew Rd., Paisley, Scotland).
16. L-Glutamine (200 mM) (Gibco-BRL, Life Technologies, Renfrew Rd., Paisley,
17. RPMI 1640 with  L-glutamine (Gibco-BRL, Life Technologies, Renfrew Rd.,
Paisley, Scotland).
18. Lithium-Heparin Vacutainers, 10 mL (Becton-Dickinson, Cowely, Oxfordshire,
19. 3-mL Cryotubes (Nunc, Roskilde, Denmark).
20. Lymphocyte culture medium (LCM) should be prepared in advance and stored at
4°C: 435.9 mL of RPMI 1640, Dutch modification; 50.0 mL of 10% heat-inacti-vated FCS; 100 U/mL of penicillin, 5.0 mL of 100 mg/mL of streptomycin; and
9.1 mL of 0.2 mg/mL of sodium pyruvate.
21. 4+ should be made up in advance and stored in 9 mL aliquots in sterile containers
at –20°C until required: 50.0 mL of 20 mM L-glutamine; 100 mL of  100 U/mL
penicillin and 100 µg/mL streptomycin; 50.0 mL of 20 mg/mL sodium pyruvate.
22. Cloning medium should be made up just before use: 46.0 mL RPMI (Dutch
modification); 10.0 mL heat-inactivated FCS; 20.0 mL hybridoma medium;
1.0 mL PHA; 2.0 mL rIL-2; 8.0 mL  4+; 1  × 107 feeder cells resuspended in a
volume of 3.0 mL.
3. Methods
3.1. Isolation of Lymphocytes from Peripheral Blood Samples
1. Peripheral blood samples should be collected into lithium–heparin-coated blood
tubes (see Note 2).
2. Following collection, gently dilute whole blood 1:1 with RPMI 1640 (Gibco-BRL)
at room temperature. Carefully layer 15 mL of diluted blood onto 10 mL of LSM
at room temperature. Centrifuge at 400g for 30 min at room temperature  (switch
Mutation and the Aging Process 183
brake off on centrifuge). Following centrifugation gently aspirate (see Note 3) the
upper layer using a sterile glass Pasteur pipet, to within 0.5 cm of the opaque
interface containing the mononuclear cells (MNCs). Discard the aspirate.
3. Gently aspirate the opaque layer containing the MNCs and transfer to a sterile
universal containing 15 mL of RPMI 1640 (Gibco-BRL). Mix MNC suspension
gently by aspiration and then centrifuge at 250g for 10 min. Remove supernatant
by aspiration and discard.
4. Pool each MNC pellet (for each subject blood sample) and wash 1× with RPMI
1640 (Gibco-BRL) at room temperature. Remove an aliquot, add trypan blue,
and count viable cells using a hemocytometer. Recovery is generally in the region
of 2.5 million MNC/mL of whole blood.
5. Resuspend MNCs in LCM at a concentration of 1 × 106 cells/mL and incubate over-night at 37°C, in a 5% CO2/air, humidified atmosphere. Monocytes will adhere to the
surface of the flask while lymphocytes remain in suspension in the culture medium.
3.2. Cryopreservation of MNCs
It may not always be possible to perform the cloning assay on the same day
as the blood is collected. In such circumstances the MNC fraction can be
cryopreserved until required.
1. Prepare freeze down medium (FDM): 90% FCS (not heat inactivated) and 10%
2. When the MNC fractions from one blood sample have been prepared (Subhead-ing 3.2., step 4) add FDM to achieve a final concentration of 3 × 106 MNCs/mL.
3. Transfer the resultant cell suspension into 3-mL cryotubes and then place the
tubes into a polystyrene box. Put the box in a freezer at –70°C overnight, then
transfer to liquid nitrogen for long-term storage.
3.3. Thawing of Previously Cryopreserved Lymphocytes
If cryopreserved, lymphocytes have to be gently thawed 24 h prior to cloning.
1. Thaw the required number of vials rapidly in a 37°C water bath.
2. Transfer the contents of the vials to a universal and dilute dropwise, slowly with
cold RPMI 1640 (Dutch Modification) to a volume of 20 mL. Agitate the univer-sal container during addition of the RPMI.
3. Centrifuge at 440g for 10 min. Remove supernatant and resuspend in warm RPMI
1640 containing 5% FCS (heat-inactivated).
4. Centrifuge at 440g for 10 min.
5. Discard supernatant and resuspend pellet in warm culture medium at a concen-tration of 1 × 106 cells/mL.
6. Incubate overnight at 37°C in a 5% CO2/air, humidified atmosphere. After overnight
incubation count lymphocytes using a hemocytometer. The number of lymphocytes
per milliliter is usually less than 1 × 106. A successful thaw will yield 7–9 × 105 cells/mL.
A count of less than 4 × 105 cells/mL should not be used for cloning.
184 Barnett and Barnett
7. Dilute lymphocyte suspension to 2 × 105 cells/mL using warm culture medium
(RPMI 1640 (Dutch modification), 10% heat-inactivated FCS, 100 U/mL of peni-cillin, 100 mg/mL of streptomycin and 0.2 mg/mL of sodium pyruvate.
3.4. Determination of Cloning Efficiency
in Non-Selective Medium
1. Take 1 mL of lymphocytes at 2 × 105 cells/mL and carry out the serial dilutions in
duplicate (A–E) (see Table 1).
2. Add 10 mL of cloning medium to 10 mL of E in a sterile universal container.
Keep universal in 37°C water bath until ready to plate out.
3. Decant cloning mixture into a sterile plastic reagent reservoir (see Note 4).
4. Using an eight-channel multipipet and sterile tips, transfer 200 µL of the mixture
into each well of a 96-well microtiter plate. Set up two such 96-well plates.
Clearly code each plate.
3.5. Determination of Cloning Efficiency in Selective Medium
1. For each selective plate to be made up (5–8 in total should be set up) mix together
in a universal container 10 mL of lymphocytes at 2  × 105 cells/mL, 9 mL of
cloning medium, and 1 mL of 6-TG.
2. Decant cloning mixture from the universal container into the reagent reservoir.
3. Using an eight-channel multipipet and sterile tips, transfer 200  µL of cloning
mixture into each well of a 96-well microtiter plate. Clearly code each plate.
3.6. Incubation and Scoring of Cloning Efficiency Plates
1. Stack all nonselective and selective plates for each blood sample and tape together
(see Note 5). Incubate plates on a sloped shelf at 37°C in a 5% CO2/air humidi-fied atmosphere for 14–15 d.
2. At 1–2 d before the end of the plate incubation period, reverse the slope of the
plates by turning through 180°. The live cells present in each well should move
toward the top of the well and form a crescent.
Table 1
Preparation of a 30 Lymphocyte/mL Suspension
Volume (mL) of
Serial lymphocyte suspension Volume of Resultant
dilution from 2 × 105 cells/mL warm (37°C) concentration of
step initial suspension culture medium lymphocytes/mL
A 1.0 1.0 1 × 105
B 0.5 of suspension A 4.5 1 × 104
C 0.5 of suspension B 4.5 1 × 103
D 0.5 of suspension C 4.5 1 × 102
E 3.75 of suspension D 8.75 30
Mutation and the Aging Process 185
3. To score the plates, place each plate on the stage of an inverted microscope and
commence scoring from top left hand well down the remaining wells. Move
across and score upwards. Exclude all wells in which evaporation has taken place.
4. A well should be scored positive if more than 50 viable lymphocytes are present
(see Note 6).
3.7. Calculation of MF/106 cells
Using the figures obtained (number of positive or number of negative wells
and total number of wells counted per plate) from the scoring of the CE plates
in selective and nonselective conditions the following equations are used to
calculate MF/106 cells.
CE in nonselective plates or selective plates = –ln (P0) CE plate / no. of cells per well
P0 = no. of negative wells / total no. of wells
MF/106 cells = CE in selective plates / CE in nonselective plates
4. Notes
1. For each new batch of IL-2 purchased it is necessary to establish the optimal
amount of IL-2 to support lymphocyte culture. To do this, a number of clon-ing efficiency plates (non-selective conditions) are set up using a range of
IL-2 concentrations, for example, 50, 100, 150, 200, 250, 300, 350, 400, and
450 U/mL. The concentration at which optimum growth conditions (highest
cloning efficiencies in nonselective plates) are achieved is then chosen for
use in all subsequent cloning assays until that batch of IL-2 runs out.
2. Following collection, blood tubes, if they require transportation, should be
protected from sudden temperature changes and vibration, as these factors
contribute to low lymphocyte cloning efficiencies.
3. An aspiration pump creates a vacuum to remove all unwanted layers from
the separation process. A vacuum pump is attached to a Buchner flask, cre-ating a vacuum that extracts all unwanted materials through a glass Pasteur
4. The same reagent reservoir can be used per sample for nonselective and selective
plates provided that the nonselective plates are plated first, so as not be to con-taminated with 6-TG.
5. To reduce evaporation from the 96-well plates, place the plates inside a plas-tic storage container with small holes to permit gaseous exchange.
6. Viable cells are pleomorphic with regular borders and bright, nongranular
centers. They form tightly packed, regular shaped clumps. Dead feeder cells
and lymphocytes (various sizes with irregular borders and dark granular cen-tres) also form clumps but these clumps are loosely packed.
186 Barnett and Barnett
Dr. Yvonne Barnett is grateful for the support of Dr. Jane Cole, MRC Cell Muta-tion Unit, University of Sussex for help with the establishment of the cloning assay
within the Cancer and Aging Research Laboratories at the University of Ulster. The
authors acknowledge the assistance of Ms. Caroline Warnock in the prepara-tion of this manuscript.
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HPRT Mutational Spectrum 189
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Somatic Mutations and Aging
Methods for Molecular Analysis of HPRT Mutations
Sai-Mei Hou
1. Introduction
The increasing information on the specific DNA sequence alterations that
occur in mutated genes of human somatic cells has allowed the establishment
of mutational spectra. Endogenous and exogenous exposures as well as indi-vidual susceptibility factors seem to contribute to the complexity of mutational
spectra. The X-linked HPRT (hypoxanthine phosphoribosyl transferase) gene
in human T lymphocytes has been considered as a suitable target for studying
somatic in vivo mutations  (1), including those related to aging. The mutant
frequency in adults is 10-fold higher than that in newborns and the proportion
of point mutations is also considerably higher in adults (90% vs 20%), possibly
as results of accumulation of point mutations induced later in life and dilution
of the V(D)J-recombinase-mediated spontaneous deletions observed in new-borns (2). A significant age-related increase of HPRT mutant frequency (1–3%/yr)
has been reported in most studies, with a more rapid increase in smokers com-pared to nonsmokers (reviewed in ref. 1).
Knowledge of the whole HPRT gene sequence (3) and the polymerase chain
reaction (PCR) techniques have made the molecular characterizations of all
types of mutations possible. More than 2000 mutants have been characterized
and included in the HPRT mutation database (4). The wide spectrum of muta-tions in this reporter gene resembles very much that in the p53 tumor suppres-sor gene, sharing many common features, such as heterogeneity, fingerprints,
and strand specificity of carcinogenic agents. It would be of great interest if
mutations related to aging-specific damages could be recovered from the HPRT
mutational spectrum in human populations.
190 Hou
The HPRT mutational spectrum consists mainly of point mutations (mis-sense, nonsense, frameshift), splice mutations, and larger deletions. Multiplex
PCR (MP-PCR, 5) has replaced the Southern blotting technique used earlier
for detection of mutants with larger genomic arrangements  (6). The reverse
transcription PCR (RT-PCR) and subsequent sequencing enables the identifi-cation of point mutations in the coding region such as base substitutions and
small deletions/insertions. By this technique, splice errors (exon skippings)
and larger genomic alterations can also be recognized as aberrant mRNA/cDNA
products, but can be truly distinguished only by the genomic PCR and sequenc-ing which enables the identification of the splicing mutations (mainly in the
splice sequences at the exon/intron boundaries) and the intronic deletion break
The primary strategy for mutational screening has been to first distinguish
large genomic alterations from point mutations by MP-PCR  (7). The point
mutations are then further subdivided into splicing and coding errors by RT-PCR.
A partial gene deletion is defined by an abnormal MP-PCR pattern and a shorter
cDNA band, and a total gene deletion by an absence of HPRT DNA from both
MP-PCR and RT-PCR. A normal MP-PCR but an abnormal cDNA product
(shorter or longer) is classified as a “splice error.” Finally, a “coding error” is
characterized by normal products from both MP-PCR and RT-PCR.
However, larger deletions in normal individuals detectable by multiplex PCR
seem to be much less frequent  (7) than previously reported from Southern
analysis (6). This could be related partly to mutations affecting intronic restric-tion sites, and partly to limitations of PCR in amplifying larger deletions
involving several exons and larger insertions (> 1 kb). Mutations affecting
primer annealing would result in an overestimate of deletion frequency, but
seemed to be a rare event. The multiplex PCR requires optimal priming condi-tions for exon fragments, both within, across, and together. The reactions often
need to be repeated or verified by exon-specific PCR, not only for the “nega-tive” exons, but probably also for the “positive” exons.
Furthermore, genomic alterations can be detected only in cells from male
donors owing to the masking effect of the inactive X-chromosome in females,
although PCR across common breakpoints may be applied to screen for specific
types of larger rearrangements regardless of gender, such as the V(D)J-recombinase-mediated deletions of exons 2 and 3 (2). A screening strategy
starting with cDNA analysis followed by confirmation at the genomic level
should thus be considered as the most convenient way of screening, as it mini-mizes the total number of PCR reactions and facilitates the spectrum establish-ment of point mutations in both males and females (Fig. 1).
The methods described below (published in refs. 7–9) are modified from the
original techniques developed for molecular analysis of unexpanded mutant
HPRT Mutational Spectrum 191
clones without RNA and DNA preparation, as they directly utilize lysates pre-pared from small numbers of cells for the cDNA synthesis  (10) and the
MP-PCR (11).
In the RT-PCR, cDNA was synthesized by reverse transcription of HPRT
mRNA and amplified by a two-step (nested) PCR amplification. In the MP-PCR,
all exons of the gene can be amplified at the same time in one reaction  (5).
Each fragment is of different size, and loss of fragment or change in its size can
easily be detected after electrophoresis. For easier optimization, however, the
Coding error
Base substitution
Small deletion
Small insertion
Exon skipping
(Splice error)
Exonic breakpoints
(Genomic rearrangement)
Splicing mutation
(at splice site)
Larger rearrangement
(intronic breakpoints)
Genomic PCR
and sequencing
and sequencing
T-cell cloning
6-thioguanine selection
Fig. 1. An optimal strategy for screening and identification of HPRT mutations.
192 Hou
amplifications are carried out in two separate reactions. The short fragments
corresponding to exons 1, 2, 4, and 6 are amplified in one reaction, and the long
fragments containing exons 3, 5, 7 + 8, and 9 in a second reaction.
2. Materials
2.1. Equipments and Enzymes
1. PCR: All PCR reactions are performed in DNA-Engine™ (MJ Research, PTC-200)
or GeneAmp 2400® (Perkin Elmer, Foster City, CA, USA) thermal cyclers. All
enzymes and dNTPs originate from Promega (Madison, WI, USA) or Pharmacia
Biotech (Uppsala, Sweden).
2. Sequencing: Magnetic beads and concentrator used for strand separation are
purchased from Dynal (Oslo, Norway). Sequencing kit and equipment are
obtained from Applied Biosystems (Foster City, CA, USA).
2.2. RT-PCR Primers and Buffers
1. RT-PCR primers (20 µM stocks, f = forward, r = reverse):
RT1f; (–50) 5′-ACC GGC TTC CTC CTC CTG AG-3′ (–31)
RT2r; (721) 5′-GAT AAT TTT ACT GGC GAT GT-3′ (702)
RT3f; (–19) 5′-TAC GCC GGA CGG ATC CGT T-3′ (–1) (+/– biotinylation)
RT4r; (697) 5′-AGG ACT CCA GAT GTT TCC AA-3′ (678) (–/+ biotinylation)
2. cDNA internal sequencing primers (Cy–5 labeled, 20 mM stocks):
RT5f: (124) 5′-ATT ATG GAC AGG ACT GAA-3′(141)
RT6f: (166) 5′-GAG ATG GGA GGC CAT CAC AT-3′(185)
RT7r: (302) 5′-CTG ATA AAA TCT ACA GTC AT-3′(283)
RT8r: (373) 5′-AAG TTG AGA GAT CTT CTC CAC-3′(353)
3. cDNA cocktail: 50 mM Tris-HCl, pH 8.5; 75 mM KCl; 3 mM MgCl2; 2.5% Nonidet
P-40 (NP-40); 10 mM dithiothreitol (DTT); 500 µM of each dNTP; 1.6 µM of primer
RT2r; 1 U/µL of RNase inhibitor; and 2.5 U/µL of M-MLV reverse transcriptase.
4. 10× cDNA PCR buffer: 150 mM Tris-HCl, pH 8.5; 600 mM KCl; and 15 mM
2.3. MP-PCR Primers and Buffers
1. 10× buffer T (sterile filtered): 670 mM Tris-HCl, pH 8.8, 67 mM MgCl2, 166 mM
(NH4)2SO4, 50 mM β-mercaptoethanol, and 68 mM EDTA.
2. Lysis buffer (master mix, 150 µL/sample): 15 µL of 10× buffer T, 7 µL NP-40 (10%),
7 µL of Tween-20 (10%), 15 µL of proteinase K (1 mg/mL) and 106 µL of H20.
3. 2× MP-PCR buffer: 134 mM Tris-HCl, pH 8.8, 13.4 mM MgCl2, 33.2 mM
(NH4)2SO4, 10 mM β-mercaptoethanol, and 13.6 µM EDTA.
4. 10× primer mix (for primer sequences see Ta ble 1):
a. Long fragments (exons 3, 5, 7–8, and 9, forward and reverse): 1.6 µM MP3,
2.4 µM MP5, 2.4 µM MP78, and 2 µM MP9.
b. Short fragments (exons 1, 2, 4, and 6, forward and reverse): 5 µM MP1, 1 µM
MP2, 1 µM MP4, and 1.6 µM MP6.
HPRT Mutational Spectrum 193
3. Methods
3.1. RT-PCR
1. cDNA synthesis: Collect 6000 cells (preferably fresh) in an Eppendorf tube
with 1 mL of ice-cold phosphate-buffered saline (PBS). Centrifuge 5 min at
13,000 rpm (4°C). Resuspend the cells in 20 µL of cDNA cocktail. Incubate for
1 h at 37°C (run a control tube with cDNA cocktail only).
2. cDNA PCR: Use 5 µL of the cDNA reaction in a PCR (50 µL) with 0.2 µM of
primer RT1f and 0.1 µM of primer RT2r, 200 µM of each dNTP, PCR buffer, 10%
dimethyl sulfoxide (DMSO), and 1 U of Ta q polymerase. After an initial denatur-ation for 4 min at 94°C, run 30 cycles of 94°C (30 s), 50°C (30 s), and 72°C
(1 min), followed by a 7-min polymerization at 72°C.
3. Nested PCR: Use 2 µL of the PCR reaction as template in a nested PCR (50 µL)
with 0.4  µM of both RT3f primer and RT4r primer (one of them biotinylated)
under the same conditions as described previously.
Table 1
HPRT Primers Used in the Multiplex PCR (5)
Exona Primer sequencesb
Long fragments
3MP3F: (16252) 5′-CCT TAT GAA ACT TGA GGG CAA AGG-3′ (16275)
(1059) MP3R: (17310) 5′-TGT GAC ACA GGC AGA CTG TGG ATC-3′ (17287)
5MP5F: (31442) 5′-CAG GCT TCC AAA TCC CAG CAG ATG-3′ (31465)
(708) MP5R: (32149) 5′-GGG AAC CAC ATT TTG AGA ACC ACT-3′ (32126)
7+8 MP78F: (38667) 5′-GAT CGC TAG AGC CCA AGA AGT CAA G-3′ (38691)
(1533) MP78R: (40199) 5′-TAT GAG GTG CTG GAA GGA GAA AAC-3′ (40176)
9MP9F: (40443) 5′-GAG GCA GAA GTC CCA TGG ATG TGT-3′ (40466)
(1278) MP9R: (41720) 5′-CCG CCC AAA GGG AAC TGA TAG TC-3′ (41698)
Short fragments
1MP1F: (1205) 5′-TGG GAC GTC TGG TCC AAG GAT TCA-3′ (1228)
(626) MP1R: (1831) 5′-CCG AAC CCG GGA AAC TGG CCG CCC-3′ (1808)
2MP2F: (14678) 5′-TGG GAT TAC ACG TGT GAA CCA ACC-3′ (14701)
(572) MP2R: (15249) 5′-GAC TCT GGC TAG AGT TCC TTC TTC-3′ (15226)
4MP4F: (27765) 5′-TAG CTA GCT AAC TTC TCA AAT CTT CTA G-3′ (27792)
(334) MP4R: (28098) 5′-ATT AAC CTA GAC TGC TTC CAA GGG-3′ (28075)
6MP6F: (34850) 5′-GAC AGT ATT GCA GTT ATA CAT GGG G-3′ (34874)
(441) MP6R: (35290) 5′-CCA AAA TCC TCT GCC ATG CTA TTC-3′ (35267)
aNumber within parentheses indicates the length of exon fragment amplified.
bGenomic positions for the first and last base indicated within parentheses.
194 Hou
4. Reading: Load 10 µL of the reaction on a 3.75% polyacrylamide gel. Save the
remaining sample for magnetic bead (Dynal) separation and single-stranded DNA
3.2. MP-PCR
1. MP-lysate: Pellet 30,000 cells and wash with PBS. Resuspend the cells, add 150 µL
of lysis buffer, and mix carefully. Incubate 56°C for at least 1 h. Heat inactivate the
proteinase K at 96°C for 10 min. Store at –20°C (–80°C for longer storage).
2. MP-PCR: Mix the following for a 50-µL PCR: 25 µL 2× MP-PCR buffer (pre-heated at 37°C for 10 min), 5 µL of DMSO, 3 µL of dNTP mix (25 mM), 5 µL of
10× primer mix, and 3.7 µL of H2O. Overlay the reaction with mineral oil and run
a hot start (80–94°C, 5 min) in a thermocycler. Add 4 U of Ta q polymerase and
8 µL of the template. Heat up the reaction to 94°C (4.5 min) and run 33 cycles of
94°C (30 s), 61°C (2 min), and 68°C (2 min), followed by a 7 min prolonged
polymerization (68°C).
3. Reading: Load 25 µL of the PCR product on a 1.4% agarose gel.
3.3. Exon-Specific PCR
For identification of splicing mutations, exon-specific primers closer to the
splice sites are used at different annealing temperatures (Table 2). Run a hot
start and 30 PCR cycles (30 s each segment) in the cDNA PCR buffer but with
5 µL of MP-lysate, 0.5  µM of each primer, 200  µM of each dNTP, 10% of
DMSO, and 1 U of Ta q polymerase.
3.4. Direct Sequencing
1. Strand separation: Immobilize biotinylated PCR product (45 µL) on streptavidin-coated magnetic beads. Separate DNA strands in alkali (1  M NaOH, 0.075%
Tween-20) with the help of a magnetic concentrator. Precipitate nonbiotinylated
DNA strand with 1/10 volume of 3 M sodium acetate, pH 5.2, and 2 vol of ice-cold 95% ethanol. Wash twice with 70% ethanol. Vacuum dry and dissolve in
2. Sequencing reactions: Use the biotinylated or nonbiotinylated strand, 0.2 µM of a
Cy–5 labeled sequencing primer, and PRISM Sequenase® Terminator single-stranded DNA sequencing kit according to the manufacturer’s instructions.
3. Run the reactions on a 373A or 377A Automated Sequencer.
4. Notes
4.1. RT-PCR
The main problem in the RT-PCR has been failure in obtaining PCR prod-ucts from all MP-PCR-positive mutants, especially when using frozen pellets
after long-term storage. This may be explained partly by difficulties in han-dling small frozen cell pellets, and partly by limited quality and quantity of
templates in the beginning. The latter may partly be related to growth arrested
HPRT Mutational Spectrum 195
clones, which may be comparable to the unsuccessfully expanded clones. The
mutation itself, such as nonsense, frameshift, deletion, and splice mutation,
may also result in low copy number of mRNA (12).
Make at least two pellets for both RT- and MP-PCR, and save the remaining
of the clone as “rest pellet.” Store all pellets at –80°C. Use pellets (preferably
fresh) from nonselected (wild-type) clones or previously successfully amplified
lysates as positive controls. The possibility of using another gene (expressed at
a comparable level) as an internal control for cDNA synthesis/PCR may also
be considered. For RT-PCR-negative mutants, the “rest pellets” can be
subjected to isolation of RNAs using commercially available kits. When
necessary, the individual exons may be amplified and screened for mutation by
SSCP (single-strand conformation polymorphism) before sequencing of the
mutated exon.
Table 2
HPRT Exon-Specific Primers Used for Genomic PCR and Sequencing (8)
Exon Primer sequence
1 E1Fb: (1599) 5′-GCG CCT CCG CCT CCT CCT CTG-3′ (1619)
(65°C)a E1Rc: (1801) 5′-CCG CCC GAG CCC GCA CTG-3′ (1784)
2 E2Fc: (14678) 5′-TGG GAT TAC ACG TGT GAA CCA ACC-3′ (14701)
(61°C) E2Rb: (15249) 5′-GAC TCT GGC TAG AGT TCC TTC TTC-3′ (15226)
3 E3Fc: (16519) 5′-TCC TGA TTT TAT TTC TGT AG-3′ (16538)
(55°C) E3Rb: (16973 5′-ATA TCC TCC AAG GTG ACT AG-3′ (169549)
4 E4Fc: (27765) 5′-TAG CTA GCT AAC TTC TCA AAT CTT CTA G-3′ (27792)
(57°C) E4Rb: (28098) 5′-ATT AAC CTA GAC TGC TTC CAA GGG-3′ (28075)
5 E5Fc: (31440) 5′-TAC AGG CTT CCA AAT CCC AG-3′ (31459)
(57°C) E5Rb: (31707) 5′-GCT TAC CTT TAG GAT GGT GC-3′ (31688)
6 E6Fc: (34728) 5′-CCT GCA CCT ACA AAA TCC AG-3′ (34347)
(57°C) E6Rb: (35781) 5′-TCT GCC ATG CTA TTC AGG AC-3′ (35762)
7+8 E78Fb: (39768) 5′-CCC TGT AGT CTC TCT GTA TG-3′ (39887)
(50°C) E78Rc: (40200) 5′-TTA TGA GGT GCT GGA AGG AG-3′ (40181)
9 E9Fc: (41353) 5′-AAC CCT GAC AAC TAA TAG TG-3′ (41372)
(57°C) E9Rb: (41551) 5′-AGG ACT CCA GAT GTT TCC AA-3′ (41532)
aAnnealing temperature within parentheses.
bAfter primer name indicates that the primer is biotinylated.
cAfter primer name indicates that the primer is Cy-5 labeled.
196 Hou
4.2. MP-PCR
Use preferably DNA or lysates from nonselected (wild-type) clones as posi-tive controls, and DNA from the RJK853 cells with a total HPRT-gene deletion
(13) as a negative control. To confirm partial deletions, amplify single HPRT
exons using conditions for MP- or exon-specific PCR (Tables 1 and 2), but
with a primer concentration of 0.5 µM, 200 µM of each dNTP and 1 U of Ta q
To distinguish methodological failure from total deletion, include a primer
pair for another gene as an internal DNA/PCR control in an exon-specific PCR.
The intronless coding region of the N-acetyltransferase 2 gene can be ampli-fied by using forward primer (1)5′-GTC ACA CGA GGA AAT CAA ATG-3′(21)
and reverse primer (1002)5′-GAG AGG ATA TCT GAT AGC AC-3′(1022) at an
annealing temperature of 58°C (14).
4.3. Siblings and Spectrum
For establishing a more reliable spectrum, the clonality should be checked
for clones with identical mutations from same individual, especially when the
individual in vivo mutant frequency is extremely high. The method for analysis
of T-cell receptor (TCR) γ-gene rearrangement have been described by de Boer
et al.  (15). Clonal MP-lysate can be used in a two-step PCR with primers
(annealing at 60 and 63°C) originally described by Bourguin et al. (16):
Outer forward: 5′-GAA GCT TCT AGC TTT CCT GTC TC-3′ (Var)
Outer reverse: 5′-CGT CGA CAA CAA GTG TTG TTC CAC-3′ (J)
Inner forward: 5′-CTC GAG TGC GCT GCC TAC AGA GAG G-3′ (Var)
Inner reverse: 5′-GGA TCC ACT GCC AAA GAG TTT CTT-3′ (J)
The nested PCR product is subjected to an analysis of restriction fragment
length polymorphisms (RFLPs) with restriction digestions (BstO1 and RsaI)
followed by electrophoresis in a 15% polyacrylamide gel (17).
Finally, to avoid artifacts in comparing mutational spectra between groups
of individuals with different ages and/or exposures, the number of clones col-lected from each subject for molecular analysis should be kept as even as pos-sible. A large collection of mutations identified from many different subjects
would probably be more suitable than that from few individuals.
1. Cole, J., and Skopek, T. (1994) Somatic mutant frequency, mutation rates and muta-tional spectra in the human population in vivo. Mutat. Res. 304, 33–106.
2. Fuscoe, J. C., Zimmerman, L. J., Lippert, M. J., Nicklas, J. A., O’Neill, J. P., and
Albertini, R. J. (1991) V(D)J recombinase-like activity mediates hprt gene deletion
in human fetal T-lymphocytes. Cancer Res. 51, 6001–6005.
HPRT Mutational Spectrum 197
3. Edwards, A., Voss, H., Rice, P., Civitello, A., Stegeman, J., Schwager, C., Zim-merman, J., Erfle, H., Caskey, C. T., and Ansorge, W. (1990) Automated DNA
sequencing of the human hprt locus. Genomics 6, 593–608.
4. Cariello, N. F. (1994) Software for the analysis of mutations at the human  hprt
gene. Mutat. Res. 312, 173–185 (database updated 1996)
5. Gibbs, R. A., Nguyen, P. N., Edwards, A., Civitello, A. B., and Caskey, C. T. (1990)
Multiplex DNA deletion detection and exon sequencing of the hypoxanthine
phosphoribosyltransferase gene in Lesch-Nyhan families. Genomics 7, 235–244.
6. Albertini, R. J., O’Neill, J. P., Nicklas, J. A., Heintz, N. H., and Kelleher, P. C.
(1985) Alterations of the hprt gene in human in vivo-derived 6-thioguanine-resis-tant T lymphocytes. Nature 316, 369–271.
7. Österholm, A.-M., Fält, S., Lambert, B., and Hou, S.-M. (1995) Classification of
mutations at the human hprt-locus in T-lymphocytes of bus maintenance workers
by multiplex-PCR and reverse transcriptase-PCR analysis.  Carcinogenesis 16,
8. Österholm, A.-M. and Hou, S.-M. (1998) Splicing mutations at the hprt locus in
human T-lymphocytes in vivo. Environ. Mol. Mutagenesis 32, 25–32.
9. Podlutsky, A., Österholm, A.-M., Hou, S.-M., Hofmaier, A., and Lambert, B. (1997)
Spectrum of point mutations in the coding region of the hypoxantine phosphoribosyl
transferase (hprt) gene in human T-lymphocytes  in vivo. Carcinogenesis  19,
10. Yang, J. L, Maher, V. M., and McCormick, J. J. (1989) Amplification and direct
nucleotide sequencing of cDNA from the lysate of low numbers of diploid human
cells. Gene 83, 347–354.
11. Fuscoe, J. C., Zimmerman, L. J., Harrington-Brock, K., and Moore, M. M. (1992)
Large deletions are tolerated at the hprt locus of in vivo derived human T-lympho-cytes. Mutat. Res. 283, 255–262.
12. Steen, A.-M., Sahlén, S., Hou, S.-M., and Lambert, B. (1993) Hprt-activities and
RNA phenotypes in 6-thioguanine resistant human T-lymphocytes. Mutat. Res. 286,
13. Yang, T. P., Patel, P. I., Chinault, A. C., Stout, J. T., Jackson, L. G., Hildebrand, B.
M., and Caskey, C. T. (1984) Molecular evidence for new mutation at the hprt locus
in Lesch–Nyhan patients. Nature 310, 412–414.
14. Hou, S.-M., Lambert, B., and Hemminki, K. (1995) Relationship between hprt
mutant frequency, aromatic DNA adducts and genotypes for GSTM1 and NAT2 in
bus maintenance workers. Carcinogenesis 16, 1913–1917.
15. de Boer, J. G., Curry, J. D., and Glickman, B. W. (1993) A fast and simple method
to determine the clonal relationship among human T-cell lymphocytes. Mutat. Res.
288, 173–180.
16. Bourguin, A., Tung, R., Galili, N., and Sklar, J. (1990) Rapid, nonradioactive detec-tion of clonal T-cell receptor gene rearrangements in lymphoid neoplasms. Proc.
Natl. Acad. Sci. USA 87, 8536–8540.
17. Bastlova, T. and Podlutsky, A. (1996) Molecular analysis of styrene oxide-induced
hprt mutation in human T-lymphocytes. Mutagenesis 11, 581–591.
Susceptibility of LDL to Oxidation 199
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Assessment of Susceptibility
of Low-Density Lipoprotein to Oxidation
Jane McEneny and Ian S. Young
1. Introduction
It is now recognized that oxidation of low-density lipoprotein (LDL) is a key
event in the development of atherosclerosis (1). In vivo, oxidation is believed to
occur primarily in the arterial wall. In early atherosclerotic lesions oxidation
may be initiated by enzymes, including myeloperoxidase and 15-lipoxygenase,
or by reactive nitrogen species, while in more advanced lesions transition met-als including copper play a role (2). Oxidized LDL has a number of atherogenic
properties (3). It is taken up via the macrophage scavenger receptor and pro-motes the formation of foam cells. It is chemotatic for circulating monocytes
and inhibits the migration of tissue macrophages out of the arterial wall. In addi-tion, oxidized LDL promotes platelet aggregation, is directly toxic to cells in the
arterial wall, promotes the synthesis of a range of cytokines and growth factors,
and inhibits nitric oxide mediated arterial dilation.
The assessment of the susceptibility of LDL to oxidation in vitro has become
a widely used technique since it was introduced by Esterbauer (4). The assay is
based on the assumption that LDL, which is easily oxidized in vitro, will also be
readily oxidized in vivo. Following isolation, the oxidation of LDL is monitored
by continuously measuring absorbance at 234 nm, which reflects the formation
of conjugated dienes. The period of time prior to an increase in absorbance is
referred to as the lag time, and is the period during which endogenous antioxi-dants within the LDL particle protect it from oxidation. A prolonged lag time
implies increased resistance to oxidation. There then follows a period of rapid
increase in absorbance referred to as the propagation phase, during which a free
radical mediated chain reaction results in peroxidation of polyunsaturated fatty
acids within the LDL particle. When all polyunsaturated fatty acids have become
200 McEneny and Young
oxidized, there is a second plateau period, which may be followed by a slight
decrease in absorption due to decomposition of lipid hydroperoxides.
Because this technique involves isolation of LDL from aqueous phase chain
breaking antioxidants such as urate and ascorbate, the susceptibility of LDL to
oxidation is influenced mainly by its composition and size (5). Increased anti-oxidant content, particularly of α-tocopherol and ubiquinol, favors increased
resistance to oxidation. The fatty acid composition is also important, with an
increased percentage content of monounsaturated fatty acids protecting against
oxidation and an increased content of polyunsaturated fatty acids decreasing
resistance to oxidation. An increase in the cholesteryl ester to cholesterol ratio
also results in increased susceptibility to oxidation, as does a preponderance of
small dense LDL particles.
Increased susceptibility of LDL to oxidation is a feature of many of the
degenerative diseases associated with aging. In healthy elderly men lag times
are reduced in comparison with younger subjects (6). Increased LDL oxidation
is also a feature of established coronary artery disease, peripheral vascular dis-ease, diabetes mellitus, and stroke (7–10). Oxidation of LDL may contribute to
impaired vascular endothelial reactivity in old age (11), and hence contribute
to the development of both myocardial ischemia and cerebrovascular disease.
Various interventions have been shown to improve the resistance of LDL to
oxidation, including antioxidant supplements and change to a diet rich in
monounsaturated fatty acids (12,13). However, as yet there are no prospective
studies showing that a reduced lag time is an independent risk factor for the
development of vascular disease.
The assay as originally described by Esterbauer involved a prolonged ultra-centrifugation step (up to 24 h) to isolate LDL from EDTA plasma, followed by
dialysis to remove potassium bromide, EDTA, and small chain breaking anti-oxidants such as urate and ascorbate. In total, the procedure required approx
48 h, and artifactual oxidation and loss of endogenous antioxidants from LDL
was therefore a problem (14). The modified procedure as described here has a
number of significant advantages. First, a rapid ultracentrifugation protocol using
a bench top ultracentrifuge allows the isolation of LDL in 1 h. Second, LDL is
purified by size-exclusion chromatography on a Sephadex column, obviating
the need for prolonged dialysis. Third, the measurement of the lag time prior to
the onset of oxidation and the calculation of rate of propagation has been auto-mated, removing any possibility of subjective bias in determining results.
2. Materials
Metal ion contamination in buffers and solutions used in the extraction and
purification of LDL can initiate oxidation prematurely. To minimize their pres-ence all solutions are made using Millipore quality water.
Susceptibility of LDL to Oxidation 201
1. Fasting peripheral venous blood is taken into 10-mL glass tubes containing
lithium heparin (50 kU/L). Plasma is isolated by centrifugation (1500g for
10 min) and stored in 1-mL aliquots at –70°C until required. These samples are
stable for up to 6 mo (see Note 1).
2. Polyallomer belltop centrifuge tubes (3 mL; 13  × 32 mm; Beckman cat. no.
3. Potassium bromide; 0.4451 g (Sigma cat. no. P5912) is placed directly into the
ultracentrifuge tube. This is performed by means of a small homemade funnel
made from a small sheet of flexible plastic, containing an opening small enough
to fit into the neck of the ultracentrifuge tube.
4. Sodium chloride solution (d = 1.006g/mL: 0.196 molal — made by adding 11.42 g to
1 L deionized water). This solution is stored at 4°C and is stable for up to 1 mo.
5. Phosphate-buffered saline (PBS), pH 7.4, 0.01 M. This solution is stored at 4°C
and is stable for up to 1 mo.
6. Peristaltic pump for overlaying NaCl solution onto density adjusted plasma. We
use a Gilson Miniplus 2 (Gilson Medical Electronics, France, cat. no. 50482) at a
setting of 400 for this procedure. The pump is fitted with narrow-bore Auto
Analyser tubing (Bran and Luebbe, Germany, cat. no. 116-0549-08) or similar,
connected to a 21 gauge needle (21G) attached using parafilm.
7. Beckman tabletop ultracentrifuge (TL100), together with a Beckman fixed-angle
rotor 30° (TL100.3). The advantage of using a benchtop ultracentrifuge of this type is
that it allows rapid isolation of LDL, greatly shortening the overall length of the
procedure and allowing the isolation to be carried out in the absence of EDTA.
8. Beckman Tube Topper Sealer, which includes tube topper, stand, seal cap, seal
guide, heat sink, and removal tool (Beckman cat. no. 348137) together with tube
spacers (Beckman cat. no. 355937). The latter are required to prevent distortion
or movement of the ultracentrifuge tubes within the rotor.
9. 21G needle, 2-mL syringe (clean needle and syringe for each LDL sample to be
isolated, plus two extra needles per ultracentrifugation run).
10. 2-mL O-ring tubes for collection of crude LDL (Sarstedt cat. no. 72.694), 10-mL
tubes for collection of desalted PD10-treated LDL (Sarstedt cat. no. 57.469), and
4-mL tubes for protein estimation and standard curve (Sarstedt cat. no. 55.478).
11. PD10 columns containing Sephadex G25 (Pharmacia, Milton Keynes, UK). These
columns are disposable. However, they can be reused many times by ensuring
they are thoroughly washed with at least 25 mL of PBS immediately prior to
reuse. Columns are prepared and stored at 4°C.
12. Bovine serum albumin (BSA) solution, 25 µg/mL (Sigma cat. no. A2153) (see
Note 2).
13. Bio-Rad dye reagent, 300  µL/sample (Bio-Rad 500-006, Bio-Rad Hemel,
Hempstead, UK), stored at 4°C.
14. Disposable semimicro cuvets, 1 mL (Sarstedt, cat. no. 67.746).
15. Semimicro quartz cuvets (cells), 1 mL (Starna, Romford, Essex). The number of
quartz cells required depends on the capacity of the spectrophotometer; in our
case we require 6 cells.
202 McEneny and Young
16. Disposable cups, 20 mL (Sarstedt, cat. no. 73.1056).
17. Nusonics ultrasonic cleaner (Quayl Dental, Worthing, Sussex).
18. Copper chloride solution (40  µmol/L, BDH cat. no. 10088). This solution is
stored at 4°C and is made in sufficient quantities for use in a series of experi-ments (see Note 3).
19. Saline solution (0.9% NaCl: BDH cat. no. 10241AP).
20. Decon-90 (4% solution; Decon Laboratories, E. Sussex).
21. HCl (0.5 M made from 37%, Janssen cat. no. 12.463.47).
22. Thermostatically controlled spectrophotometer (37°C, Hitachi U-2000-1, cat. no.
HIT/121-0032), containing automatic six-cell positioner (Hitachi cat. no. HIT/
121-0304) linked to a PC with software package for automatic handling of data
(Hitachi, Enzyme Kinetic Data System).
23. Microsoft Excel software package for data computation utilizing a specially writ-ten macroprogram (15) (a copy of this macro is available from the authors).
3. Methods
Separation of the lipoprotein species is achieved by flotation nonequilib-rium ultracentrifugation. The method described is very rapid and requires
less than 2 h total preparation time (including ultracentrifugation) prior to
1. To a Beckman 3-mL ultracentrifuge tube containing 0.4451 g of KBr is added
0.9 mL of heparinized plasma. This plasma is added from a pipet connected to
fine-bore tubing (approx 2.5 cm in length and a diameter small enough to enter
the neck of the ultracentrifuge tube). The ultracentrifugation tube is temporarily
sealed with parafilm, allowing gentle inversion to dissolve the salt. This solution
has a resultant density of 1.30 g/mL.
2. To this density-adjusted plasma, NaCl (d = 1.006 g/mL) is added, using the tub-ing and 21G needle connected to the peristaltic pump. The needle requires shap-ing to enable it to rest on the inside of the ultracentrifuge tube. Allow the solution
to gently run down the side onto the top of the density adjusted plasma (ensure no
mixing or distortion of the layers occurs during this procedure).
3. The ultracentrifugation tubes are sealed using the Beckman Tube Topper Sealer.
They are gently placed in the Beckman rotor together with the Beckman spacers.
Ultracentrifugation is performed using the following settings: 4°C, 100,000 rpm
(541,000g) for 1 h. An acceleration and deceleration setting of 6 is chosen; total
ultracentrifugation run time is 1 h and 6 min. The acceleration and deceleration
parameters prevent disturbance of the density gradient created during ultracen-trifugation.
Note: During ultracentrifugation the following steps are performed.
a. Preparation of columns: one column per sample is prepared (at 4°C) by wash-ing through with 25 mL of PBS prior to use. These columns are designed to
retain a reservoir of buffer within the gel bed. They do not run dry and may be
left unattended for short periods of time.
Susceptibility of LDL to Oxidation 203
b. Preparation of quartz cells (see Note 4).
c. Preparation of protein standard curve (see Note 2).
4. On completion of ultracentrifugation, the ultracentrifuge tubes are gently
removed from the rotor and placed in the rack holder. The LDL band is visible in
all cases, and in most is a distinct orange band located approximately one third
distance from the top of the tube. It is distinctly separate from other lipoprotein
species, that is, very-low-density lipoprotein (VLDL) and high-density lipopro-tein (HDL) (the former being found at the top of the tube and the latter below the
level of LDL) (see Fig. 1). The sample within the ultracentrifuge tube is contained
within a vacuum that has to be broken to remove the sample without distortion of
the layers. This is achieved by first inserting a needle (21G) at least twice at the
top of the tube, leaving it in place on the second puncture; this allows the entry of
air into the tube. At the position of the LDL layer puncture the side of the tube
with a second needle to remove a plug of polyallomer that would otherwise
prevent the removal of sample. The LDL layer is then removed through this open-ing by a third needle attached to a 2-mL syringe in a volume of 0.9 mL (ensure
the needle remains within the LDL band). A new needle and syringe is used for
the collection of each LDL sample. (Removal of the LDL by aspiration mini-mizes contamination with other lipoproteins and plasma proteins which can occur
if the sample is collected by downward fractionation). The LDL sample is placed
in a 2-mL O-ring tube and placed on ice for immediate use. This sample equates
to “crude” LDL.
Fig. 1. Positions of VLDL, LDL, and HDL after ultracentrifugation.
204 McEneny and Young
5. Size-exclusion chromatography for desalting crude LDL using the PD10 column.
Crude LDL is contaminated with KBr. The sample also contains small molecules
such as urate that can alter its susceptibility to oxidation. Desalting is performed
as follows: 0.5 mL of crude LDL is added to the prewashed PD10 columns; this is
allowed to enter the gel bed. Two milliliters of PBS is then added and also allowed
to enter the gel bed. Both these eluants are discarded. The LDL is then obtained
by adding a second 2-mL aliquot of PBS and this eluant is collected into a 10-mL
tube. The LDL sample is now termed “desalted” (PD10) LDL and is placed on
ice until the sample is ready to be oxidized after protein determination. (After
collection of the LDL sample start preparing the columns for reuse by washing
with PBS before sealing).
Note: Subjecting LDL to column chromatography renders it less stable. Crude
LDL can be stored at 4°C for up to 2.5 h with little deterioration of lag time.
However, PD10-treated LDL must be used as rapidly as possible. It is therefore
important to prepare all stages post PD10 treatment in advance while ultracen-trifugation is being performed (e.g., quartz cells and protein assay).
6. Protein determination and standardization of PD10 LDL: LDL protein is read
against the prepared BSA standard curve. The dilution of LDL is taken into
account by multiplying the figure obtained by a factor of 15, as 100 µL of PD10
LDL is added to 1100 µL of water plus 300 µL of Bio-Rad dye reagent, giving a
final volume of 1500 µL and thus a dilution of 15. The LDL is then standardized
to 50 µg/mL of protein with PBS, taking into account the volume of copper solu-tion added. The final volume of each cuvet is 1000 µL.
The volume of LDL (Y) and PBS (Z) required can be obtained using the
following formula:
Y µL of LDL = 50/X µg/mL LDL × 1000
Z = 1000 µL– (Y µL of LDL + 50 µL Cu 2+)
where X is the concentration of protein determined from the standard curve × 15,
Y is the volume of LDL required in the cell to give a final concentration of
50 µg/mL when the volume is made up to 1000 µL, and Z is the volume of PBS
required to dilute the LDL to the required concentration (see Note 5).
Dilution of the LDL sample is performed in the quartz cuvet. The copper chlo-ride solution is added after LDL and PBS. (The action of the pipet is used to mix
the copper solution with the LDL/PBS mixture by expelling and drawing the
copper solution back and forth several times. In our experience this procedure is
more effective than cell inversion). The samples are then placed into the spectro-photometer and data collection initiated.
7. Oxidation of the sample is followed in the thermostatically controlled spectro-photometer. Change in absorbance is followed at 234 nm and recorded every
2 min. The change in absorbance is recorded on a Nimbus PC-386 utilizing the
Enzyme Kinetic Data System software package. This automated system removes
the need for lengthy manual recording.
8. Oxidation profile: A typical profile is shown in Fig. 2A, and has two distinct phases:
Susceptibility of LDL to Oxidation 205
a. Initial or lag phase: This is a slope with a slow increase in absorbance. The
duration of this phase is indicative of the inherent resistance to oxidation of
the LDL sample being examined.
b. Propagation phase: The lag phase is followed by a steep increase in absor-bance due to the rapid formation of conjugated dienes. This occurs only when
all the endogenous antioxidants within the LDL molecule and their protective
Fig. 2. Typical curve obtained by monitoring oxidation of LDL with copper at 234 nm.
206 McEneny and Young
effect have been utilized. When the propagation phase is complete a plateau
of maximum absorbance is reached.
9. Data analysis:  See Fig. 2B. Determination of lag time using the mathematical
program removes subjective error which may occur if determined manually. This
is calculated using a specially written macroprogram after the results obtained on
the PC are converted into ASCII file format and imported into the spreadsheet
program Excel. The lag time is calculated as the intercept between the line of
maximum slope of the propagation phase and the baseline where absorbance was
at time = zero (see Note 6).
4. Notes
1. Heparinized plasma is used in preference to serum purely for logistic reasons, as
serum collection lengthens total preparation time. However, this technique may
be applied equally well to serum samples with no detectable difference in lag
time when compared to heparinized plasma.
2. Protein standards and protein estimation: Stock BSA (25 µg/mL) is prepared by
placing 12.5 mg of BSA into a 500-mL beaker. Add approx 300 mL of deionized
water and gently stir (care is required as vigorous mixing can cause BSA to foam
and make it difficult to bring to the correct volume). Add this solution to a 500-mL
volumetric flask and rinse the beaker with remaining water to bring to correct
volume. Aliquot 7 mL of this solution into 10-mL tubes, cap, and freeze at –20°C.
This solution is stable for up to 6 mo.
Working standard solutions (0, 2.5, 10, 15, 20 µg/mL) are prepared from the
stock BSA. These together with the samples are diluted with distilled water to a
final volume of 1.2 mL and prepared in duplicate in the 4-mL tubes. Three
hundred microliters of Bio-Rad dye reagent is then added, making a final volume
1.5 mL. The standards and samples are then gently inverted and absorbance at
595 nm recorded within 5–60 min. The PD10 LDL is made up to 1.2 mL using
0.1 mL of sample and 1.1 mL of distilled water. If the protein concentration of
crude LDL is required use only 10 µL and dilute to 1.2 mL with water (dilution
factor for crude LDL is 150).
3. Copper chloride solution: The working solution of copper chloride is 40 µM. This
solution is made by serial dilution of a 33.2 mM stock solution.
a. Solution A: 33.2 mM — 0.567 g of CuCl2·2H2O made in a 100-mL volumet-ric flask using deionized water.
b. Solution B: 332 µM —1 mL of solution A is added to a 100-mL volumetric
flask and the volume is made up using saline solution.
c. Solution C: 40 µM working solution — 6.024 mL of solution B is added to a
50-mL volumetric flask and the volume made up using either PBS or saline
solution (addition of PBS in the earlier solutions causes precipitation).
4. Preparation of quartz cells: New or used cells must undergo the following proce-dure before use.
Each cell is rinsed at least 10× with distilled water. They are then placed into a
separate disposable cup containing sufficient 4% Decon–90 to cover them. The
Susceptibility of LDL to Oxidation 207
cups are placed in the sonic bath and sonicated for 5 min. The cells are then
inverted (not the cups), while trying to retain as much solution as possible within
the cells. They are then sonicated again for 5 min, removed, rinsed with distilled
water (10×), and placed in 0.5 M HCl for at least 1 h prior to use/reuse.
Before use in the oxidation experiment the acid-soaked cells must be thor-oughly cleaned. The cells are rinsed with distilled water (10×), placed into clean
cups containing water, and sonicated for 5 min, before again inverting and soni-cating for a further 5 min. The cells are then allowed to drain on absorbent paper
and are now ready for use.
The reason for this rigorous cleaning regime is because oxidation of LDL pro-duces lipoperoxides which may adhere to the side of the cells. These seed the
oxidation reaction and would artefactually increase the rate of oxidation in the
sample being examined. In our hands this rigorous cleaning procedure is essen-tial if reproducible results are to be obtained.
5. A worked example of the formula is shown here. If the concentration of LDL
protein is 157 mg/mL, determined from the standard curve X dilution factor, the
volumes of LDL (Y) and PBS (Z) are as follows:
Y = 50/157 × 1000 = 318 µL of LDL
As the LDL is made up to a final volume of 1000 µL the volume of PBS required,
taking into account the 50 µL of copper to be added, is
Z = 1000 – (318 µL of LDL + 50 µL of Cu) = 632 µL of PBS
Therefore to the cell is added 318 µL of LDL + 50 µL of Cu + 632 µL of PBS. The
final volume is 1000 µL.
6. The lag time is determined by taking the point of maximum slope over an interval
of 20 min and fitting a line by the least squares method. The point of maximum
slope was found by repeated computation of the average slope for 11 consecutive
points moving along 1 point at a time, enabling deviations due to random noise
from the spectrophotometer to be removed.
1. Witztum, J. L. (1993) Susceptibility of low-density lipoprotein to oxidative modifica-tion. Am. J. Med. 94, 347–356.
2. Heinecke, J. W. (1997) Mechanisms of oxidative damage of low density lipoprotein
in human atherosclerosis. Curr. Opin. Lipidol. 8, 268–274.
3. Westhuyzen, J. (1997) The oxidation hypothesis of atherosclerosis: an update. Ann.
Clin. Lab. Sci. 27, 1–10.
4. Esterbauer, H., Striegl, G., Puhl, H., and Rotheneder, M. (1989) Continuous moni-toring of  in vitro  oxidation of human low-density lipoprotein.  Free Radic. Res.
Commun. 6(1), 67–75.
5. Esterbauer, H., Gebicki, J., Puhl, H., and Jurgens, G. (1992) The role of lipid-peroxidation and antioxidants in oxidative modification of LDL. Free Radic. Biol.
Med. 13(4), 341–390.
208 McEneny and Young
6. Napoli, C., Abete, P., Corso, G., Malorni, A., Postiglione, A., Ambrosio, G.,
Cacciatore, F., Rengo, F., and Palumbo, G. (1997) Increased low-density lipopro-tein peroxidation in elderly men. Coron. Art. Dis. 8, 129–136.
7. Karmansky, I., Shnaider, H., Palant, A., and Greuner, N. (1996) Plasma lipid oxida-tion and susceptibility of low density lipoproteins to oxidation in male patients with
stable coronary artery disease. Clin. Biochem. 29, 573–579.
8. Iribarren, C., Folsom, A. R., Jacobs, D. R., Gross, M. D., Belcher, J. D., Eckfeldt,
J. H. (1997) Association of serum vitamin levels, LDL susceptibility to oxidation,
and autoantibodies against MDA-LDL with carotid atherosclerosis — a case-con-trol study. Arterioscler. Thromb. Vasc. Biol. 17, 1171–1177.
9. Yoshida, H., Ishikawa, T., and Nakamura, H. (1997) Vitamin E lipid peroxide ratio
and susceptibility of LDL to oxidative modification in non-insulin-dependent dia-betes mellitus. Arterioscler. Thromb. Vasc. Biol. 17, 1438–1446.
10. Palinski, W. and Horkko, S. (1997) Biological consequences of oxidation in athero-sclerosis and other pathologies. Lipoprotein oxidation in brain arteries. Nutr. Metab.
Cardiovasc. Dis. 7, 237–247.
11. Khder, Y., Bray, L., Boscs, D., Aliot, E., and Zannad, F. (1996) Endothelial, vis-coelastic and sympathetic factors contributing to the arterial wall changes during
ageing. Cardiol. Elderly 4, 161–165.
12. deWaart, F. G., Moser, U., and Kok, F. J. (1997) Vitamin E supplementation in elderly
lowers the oxidation rate of linoleic acid in LDL. Atherosclerosis 133, 255–263.
13. Parthasarathy, S., Grasse, B. J., Miller, E., Almazan, F., Khoo, J. C., Steinberg, D.,
and Witztum, J. L. (1991) Using an oleate-rich diet to reduce susceptibility of low-density lipoprotein modification in humans. Am. J. Clin. Nutr. 54, 701–706.
14. Scheek, L. M., Wiseman, S. A., Tijburg, L. M., and Vantol, A. (1995) Dialysis of
isolated low density lipoprotein induces a loss of lipophilic antioxidants and increases
the susceptibility to oxidation in vitro. Atherosclerosis 117, 139–144.
15. McDowell, I. F., McEneny, J., Trimble, E. R. (1995) A rapid method for measure-ment of the susceptibility to oxidation of low-density-lipoprotein.  Ann. Clin.
Biochem. 32, 167–174.
Analysis of Pentosidine 209
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Measurement of Pentosidine in Biological Samples
Jesus R. Requena, David L. Price, Suzanne R. Thorpe,
and John W. Baynes
1. Introduction
Pentosidine is a highly fluorescent advanced glycation end product (AGE)
and crosslink derived from one molecule of arginine and one of lysine bridged
in an imidazo-pyridinium structure (Fig. 1). It was first isolated from articular
cartilage by Sell and Monnier (1), and has now been detected and quantified
in a variety of human and animal tissues, including skin and kidney collagen
(2–5), lens crystallins (6,7), plasma (8,9), serum (10), urine (11), and synovial
fluid (12,13). Pentosidine is readily prepared from arginine, lysine, and a
pentose (hence its name). Dyer et al. (14) have also described its formation
from glucose, albeit at a slower rate and probably through oxidation of
glucose to arabinose  (15). Because its formation from either glucose or
ribose requires oxidation, pentosidine is both an AGE and a “glycoxidation”
product (16).
Pentosidine accumulates in long-lived tissue proteins with age (Fig. 2). The
kinetics of its accumulation in human skin collagen have been fitted to a linear
regression by Dyer et al. (µmol pentosidine/mol of lysine = 0.41 × age – 0.48
[r = 0.78, p < 0.001]) (3) and to an exponential one by Sell et al. (pmol pento-sidine/mg collagen = 0.008 × age2 + 0.325 × age + 3.45 [r = 0.927, p < 0.0001])
(5). The rate of accumulation of pentosidine in skin collagen is inversely
proportional to species life-span (5). Because it is a crosslink, it was hypoth-esized that pentosidine might contribute to the age-related increase in collagen
stiffness. However, the trace concentration of pentosidine in tissues, in the range
of a few millimoles per mole of triple helical collagen (Fig. 2), makes it unlikely
210 Requena et al.
that pentosidine would have a significant effect on collagen structure.
Pentosidine is, however, an excellent biomarker for nonenzymatic modifica-tion of long-lived proteins by the Maillard reaction, providing insight into over-all role of the Maillard reaction in aging and disease. Concentrations of
pentosidine in plasma protein and collagen are elevated in several pathological
conditions, especially in end-stage renal disease (2,8,10), and, to a lesser extent,
in diabetes  (2–4,6,8,11,16,17), rheumatoid arthritis  (12,13,18), atherosclero-sis, and neurodegenerative diseases (19). The elevation in pentosidine concen-tration in these diseases is attributed to increased precursor concentrations,
resulting from either increased oxidative stress and/or decreased detoxification
of reactive carbonyl intermediates in the Maillard reaction.
This chapter describes a procedure for the preparation, purification, and
calibration of a pentosidine standard, followed by a general method for mea-suring pentosidine by reverse-phase high-performance liquid chromatography
(RP-HPLC) with fluorescence detection. This method is suitable for collagen
samples, which produce clean chromatograms. In these cases, pentosidine is
normalized to the hydroxyproline (20) or lysine content of the sample, or to the
amount of protein. More complex samples, such as tissue extracts, plasma, and
serum, yield noisy chromatograms in which pentosidine cannot always be
resolved from other components. In these cases, the basic procedure is
modified to include a simple clean-up procedure using sulfopropyl–Sephadex
(SP-Sephadex) solid-phase extraction columns, based on a modification of
the method of Takahashi et al. (10). In these cases, pentosidine concentration
is expressed per mole of lysine, per milligram of protein, or per milliliter of
serum or plasma. In general, to facilitate comparisons between proteins, it
is more convenient to express the data as moles of pentosidine per mole of
Fig. 1. Structure of pentosidine.
Analysis of Pentosidine 211
2. Materials
1. Na-Acetyl-lysine, Na-acetyl-arginine,  D-ribose, trifluoroacetic acid (TFA) and
heptafluorobutyric acid (HFBA) are available from Aldrich (St. Louis, MO,
USA); SP-Sephadex C-25 from Pharmacia (Bromma, Sweden); C-18 solid phase
extraction columns (Sep-Paks) from Waters (Milford, MA, USA); and 0.22 µM
luer-adaptable filters from Millipore (Bedford, MA, USA).
2. HPLC system: Two-solvent gradient system, equipped with fluorescence and
diode array detectors. Reverse-phase, C-18 column, 15 × 0.46 cm, Vydac 218TP,
300A Pore Size; HPLC grade acetonitrile (CH3CN).
3. Speed-Vac Centrifugal evaporator system (Savant Instruments, Farmingdale, NY,
4. Heating block for heating 13 × 100 mm screw-cap test tubes at 65°C and 110°C.
5. Source of nitrogen gas (10 psi pressure).
3. Methods
3.1. Preparation of Pentosidine Standard
1. Dissolve  Na-acetyl-lysine (18.8 mg, 0.1 mmol),  Na-acetyl-arginine (21.6 mg,
0.1 mmol), and D-ribose (15 mg, 0.15 mmol) in 1 mL of 0.2 M sodium phosphate
buffer, pH 9.0, in a 13 × 100 mm screw-cap tube with Teflon-lined cap. Adjust
to pH 9 with 0.1 M NaOH.
2. Place in heating block at 65°C. Monitor pH, initially at hourly intervals, readjust-ing to pH 9.0, as needed.
Fig. 2. Effects of age and diabetes on concentration of pentosidine in human skin
collagen (3).
212 Requena et al.
3. After 48 h, dilute 300 µL of the resulting light brown solution to 3 mL with 1%
TFA and apply to a 3-mL C-18 solid-phase extraction minicolumn previously
equilibrated with 1% TFA.
4. Wash the column with 1% TFA. Collect fractions (2 mL) and monitor absorbance
at 226 and 326 nm. After the initially high absorbance at 226 nm decreases to
~0.2 and stabilizes (~25 fractions), elute the column with 5% CH3CN containing
1% TFA. Collect and pool fractions, with peak absorbance at 326 nm, corre-sponding to elution of N,N’-diacetyl-pentosidine, and dry in vacuo (see Note 1).
5. Redissolve sample in 500  µL of deionized water and purify  N,N’-diacetyl-pentosidine by RP-HPLC (Fig. 3). Apply to 15 × 0.46 cm C-18 column, equili-brated with CH3CN:0.1% HFBA in H2O (1:6), and elute isocratically at a flow
rate of 1 mL/min. Monitor absorbance at 326 nm. A single major peak corre-sponding to N,N’-diacetyl-pentosidine should appear. Collect and pool peak frac-tions; dry in vacuo.
6. Prepare pentosidine standard by hydrolysis of N,N’-diacetyl-pentosidine in 2 M
HCl for 4 h at 110°C. Dry the hydrolysate and redissolve in 1 mL of deionized
Fig. 3. RP-HPLC analysis of  N,N’-diacetyl-pentosidine, the precursor of the
pentosidine standard. Hydrolysis of this compound yields pure pentosidine, or a
slightly contaminated product that can be further purified by analytical RP-HPLC.
Analysis of Pentosidine 213
water. Check purity of pentosidine by RP-HPLC analysis, as described in Sub-heading 3.2., step 4. If fluorescent contaminants are present, repurify pentosidine
by RP-HPLC using analytical conditions, then pool pentosidine fractions. For
calibration, dissolve an aliquot of the pooled pentosidine in 0.1 M HCl, measure
absorbance at 326 nm, and calculate pentosidine concentration using a molar
extinction coefficient of 14,200 at 326 nm (Fig. 4; see Note 2). A convenient
concentration for a working standard is 0.5 pmol/µL. Store dilute, working stan-dards frozen at –20°C in 0.1% HFBA.
3.2. General Method for Measuring Pentosidine in Collagen
1. Place weighed sample of collagen (1–5 mg) in 13 × 100 mm screw-cap test tube.
Reduce by addition of 1 mL of 0.2 M sodium borate buffer, pH 9.2, followed by
100 µL of 1 M NaBH4 dissolved in 0.1 M NaOH. Reduction can be carried out for
4 h at room temperature or overnight at 4°C (see Note 3).
2. Recover collagen by centrifugation or decanting, discard supernatant, and wash
with deionized water.
3. Hydrolyze collagen in 5 mL of 6 M HCl at 110°C for 24 h in screw-cap tubes
flushed with N2 (see Note 3).
4. Dry hydrolysates in vacuo. Redissolve samples in 500 µL of 0.1 M HFBA using
vortex mixing or sonication, as required, and inject a 200-µL aliquot onto an
RP-HPLC column for analysis. HPLC solvents: A, 0.1% HFBA in H2O; B,
CH3CN. Gradient: 5 min in 10% B, followed by linear gradient of 10 to 22% B
Fig. 4. Absorbance spectrum of pentosidine in 0.1 M HCl.
214 Requena et al.
over 35 min, followed by 15-min wash with 95% CH3CN (see Note 4). Monitor
fluorescence at excitation λ = 328, emission λ = 378. Typical chromatograms are
shown in Fig. 5B,C. The limit of detection of pentosidine on our HPLC system is
approx 0.2 pmol, but will vary with instrumentation.
5. The concentration of pentosidine in the sample is determined using a standard
curve prepared with authentic pentosidine. A separate analysis of the hydrolysate
is required to measure hydroxyproline or lysine, depending on the method of
expressing the data. The intraassay coefficient of variation is typically 6–8% for
middle-aged human skin collagen.
3.3. Method for Measuring Pentosidine in Plasma Proteins
(see Note 5)
3.3.1. Sample Preparation
1. Mix 200 µL of plasma with an equal volume of 0.2 M borate buffer, pH 9.2, in a
13 × 100 mm screw-cap test tube. Add 40 µL of a freshly prepared 1 M solution of
NaBH4 in 0.1 M NaOH, and let the sample reduce at room temperature for 4 h.
Fig. 5. RP-HPLC chromatograms of analysis of  (A) normal human plasma and
(B,C) human skin collagen from 85- and 18-yr-old donors, respectively.
Analysis of Pentosidine 215
2. Precipitate protein by addition of 440 mL of 20% trichloroacetic acid (TCA)
while vortex-mixing. Centrifuge the sample at 1000g for 5 min (tabletop, clinical
centrifuge) and discard the supernatant. Wash pellet once by suspension in 10%
TCA and recentrifugation.
3. Hydrolyze protein in 5 mL of 6 M HCl for 24 h at 110°C in screw cap tube under N2.
4. Dry hydrolysate in vacuo and redissolve sample in 10 mL of deionized water and
filter through 0.22 µm luer-adaptable filters. Wash the filter with 1 mL of deion-ized water and pool with the filtrate.
3.3.2. Solid-Phase Extraction with SP-Sephadex Minicolumns
1. Swell SP-Sephadex overnight in deionized water at 4°C, according to the
manufacturer’s instructions.
2. Prepare minicolumns by filling small plastic columns with 3 mL of swollen gel.
3. Wash minicolumns with 15 mL of deionized water, apply filtered samples, and
wash with 20 mL of 0.1 M HCl. Elute with 7 mL of 1 M HCl. Pool and dry eluate
(see Note 6).
4. Redissolve eluate in 500 mL of 0.1% HFBA. Analyze a 200-µL aliquot by
RP-HPLC, as described in Subheading 3.2. A typical chromatogram of a human
plasma sample is shown in Fig. 5A. Pentosidine may be expressed as µmoles of
pentosidine per milliliter plasma or per mole of lysine, measured separately by
analysis of the plasma sample or hydrolysate. The intraassay coefficient of varia-tion is typically 8–10%.
4. Notes
1. The yield of pentosidine is typically ~1%, so the HPLC column could be over-loaded by impurities during purification. The preliminary clean-up step is
designed to eliminate excess reactants and salts that are not retarded by the
column, and brown products that are retained on the column after elution of
pentosidine. Column washing with 1% TFA is stopped when absorbance at
~226 nm is less than 0.2 or has ceased to decrease. Switch to eluting solvent (1%
TFA in 5% CH3CN) if absorbance at 326 nm begins to increase. Collected
fractions should be transparent or pale yellow. Brown color indicates overloading
of the minicolumn and calls for a decrease in the amount of sample applied. At
this stage, sacrifice yield for purity.
2. Dyer et al. (14) used radioactive lysine to calibrate their pentosidine standard. Sell
and Monnier  (1) used a gravimetrically calibrated standard (Monnier,  personal
communication). They reported a molar extinction coefficient of 4195 at 326 nm
in 0.1 M HCl. For determination of the extinction coefficient in the present study,
we prepared a sample of pure N-acetyl,N’- hippuryl-pentosidine (hippuryl = ben-zoyl-glycine) from  Na-hippuryl-lysine and  Na-acetyl-arginine. The product was
purified by C-18 solid extraction and RP-HPLC using procedures similar to those
described previously. Analysis by HPLC using a diode-array detector yielded a
single peak with an absorbance maximum at 326 nm (pentosidine) and secondary
maxima at 226 nm (amide carbonyl and carboxyl groups) and 278 nm (benzoyl
216 Requena et al.
group). Following hydrolysis in 6 M HCl for 4 h at 110°C, RP-HPLC yielded two
well-resolved absorbance peaks, with maxima at 226 nm (benzoic acid) and 326 nm
(pentosidine). The extinction coefficient for pentosidine was calculated from the
yield of glycine determined by amino acid analysis. The molar extinction coeffi-cient of 14,200 at 326 nm, measured in 0.1 M HCl, was in good agreement with
that for our previous pentosidine standard, based on the use of radioactive lysine
(14), but was 3.4 times higher than that reported by Sell and Monnier (1).
3. Reduction with NaBH4 prevents the interference from products of decomposition
of Amadori compounds and reactive AGEs on protein. This step, together with
hydrolysis under nitrogen, generally reduces the complexity of chromatograms.
4. The washing step is helpful for removing both fluorescent and nonfluorescent
impurities that may interfere with resolution of the pentosidine peak.
5. The method described is for analysis of human plasma proteins, but can be
adapted to analysis of other types of complex matrices.
6. Some pressure from a rubber bulb may be needed to start flow, but once started,
the minicolumns operate under gravity. Sample pH is ~2, and pentosidine is
retained on the column during the washing step. The gel collapses 10–20% during
elution of pentosidine in 1 M HCl and should be discarded after use.
This work was supported by Research Grant DK–19971 to J. W. B. from the
National Institutes of Diabetes, Digestive and Kidney Diseases. J. R. R. is the
recipient of a postdoctoral fellowship from the Juvenile Diabetes Foundation
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2. Sell, D. R. and Monnier, V. M. (1990) End-stage renal disease and diabetes catalyze
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Analysis of Pentosidine 217
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end product, in biological specimens. Clin. Chem. 40, 1766–1773.
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Horiuchi, K. (1993) Quantification of the cross-link pentosidine in serum from
normal and uremic subjects. Clin. Chem. 39, 2162–2165.
11. Takahashi, M., Ohishi, T., Aoshima, H., Kawana, K., Kushida, K., Inoue, T., and
Horiuchi, K. (1993) The Maillard protein cross-link pentosidine in urine from dia-betic patients. Diabetologia 36, 664–667.
12. Rodriguez-Garcia, J., Requena, J. R., and Rodriguez-Segade, S. (1998) Increased
concentrations of serum pentosidine in rheumatoid arthritis.  Clin. Chem.  44,
13. Miyata, T., Ishiguro, N., Yasuda, Y., Ito, T., Nangaku, M., Iwata, H., and Kurokawa,
K. (1998) Increased pentosidine and advanced glycation end product, in plasma
and synovial fluid from patients with rheumatoid arthritis and its relation with inflam-matory markers. Biochem. Biophys. Res. Commun. 244, 45–49.
14. Dyer, D. G., Blackledge, J. A., Thorpe, S. R., and Baynes, J. W. (1991) Formation
of pentosidine during nonenzymatic browning of proteins by glucose. J. Biol. Chem.
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(1995) Mechanism of autoxidative glycosylation: identification of glyoxal and ara-binose as intermediates in the autoxidative modification of proteins by glucose.
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J. W., and Lyons, T. J. (1993) Maillard reaction products and their relation to com-plications in insulin-dependent diabetes mellitus. J. Clin. Invest. 91, 2470–2478.
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Damage to Mitochondria 221
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Causes and Consequences of Damage
to Mitochondria
Morphological Aspects
Jaime Miquel and Carlo Bertoni-Freddari
1. Introduction
In recent years the role played by mitochondria in cellular aging has become
the focus of intensive research. The concept that these energy-producing orga-nelles are involved in aging derives from the views of Harman (1) and Gersch-man (2) linking senescence to the damaging effects of free radicals, especially
those released in the mitochondrial respiratory chain.
However, most gerontologists held the opinion that, because mitochondria
are self-replicating organelles, they should be able to counteract any age-related
loss. Our own electron microscopic studies have provided data in contradiction
with that former view. Thus, in our study of Drosophila aging we were the first
to demonstrate that the fixed-postmitotic cells of insects accumulate an age
pigment structurally similar to the lipofuscin present in mammalian cells and
that many of the pigment granules derive from degenerating mitochondria (3,4).
Further, our investigation of the testis of aged mice showed a striking mito-chondrial degeneration and loss in the fixed-postmitotic Sertoli and Leydig
cells, while the mitochondria of the fast-replicating spermatogonia (also present
in that organ) did not show any age-related change. This led us to propose the
mitochondrial theory of aging (5–7), according to which senescence is linked
to the injurious effects of oxy-radicals on the mitochondrial genome of neu-rons and other types of differentiated cells. This extranuclear somatic mutation
concept of aging is supported by the finding that mitochondrial DNA (mtDNA)
synthesis takes place at the inner mitochondrial membrane near the sites of
formation of highly reactive oxygen radicals and their products such as
222 Miquel
lipoperoxides and malonaldehyde. Further, mtDNA may be unable to counter-act the chronic oxygen stress because, in contrast to the nuclear genome, it
lacks histone protection and excision repair.
If oxidative injury is not limited to organellar membranes (as proposed by
early free radical theory advocates) but also occurs in mtDNA, the organelles
that have suffered oxidation-related genome mutation, inactivation, or loss will
be unable to rejuvenate themselves by normal replication. Further, there will
be an impaired renewal of the macromolecules coded by that genome, namely
the 13 hydrophobic polypeptides of the electron transport chain and of ATP
synthase as well as the mitochondrial rRNAs and tRNAs. This may lead to a
progressive decline in mitochondrial function with decrease in ATP production
and ATP-dependent protein synthesis.
It is very important from a clinical viewpoint that, because most cellular
energy is produced in mitochondria through the process of oxidative phospho-rylation, the age-related dysfunction and loss of these organelles must result in
bioenergetic decline. This may trigger apoptotic death (8) and play a key role
in the senescent decrease of physiological performance and in the pathogenesis
of some age-related degenerative diseases of the somatic tissues mainly com-posed of fixed-postmitotic cells such as the cardiac and skeletal muscle and the
central nervous system (CNS).
In our opinion, the preceding summary on the causes and effects of mito-chondrial aging justifies the present compilation of very detailed and practical
instructions for the electron microscopic study of mitochondria regarding tis-sue fixation, sectioning, staining, and quantitative morphological observation
of the organelles. We hope that this information will be especially useful to
researchers interested in furthering the understanding of the role of mitochon-dria as promoters or targets in the senescence of diverse cellular types and
animal models. The techniques described may also interest the workers explor-ing the modulation of the rate of animal aging by genetic manipulation, under-feeding, and pharmacological and antioxidant treatments.
2. Materials
1. Anesthetic solution: 2,2,2-Tribromoethanol (200 mg/kg body weight) in 10%
ethanol solution.
2. Perfusion solution: Prepare 0.1 M sodium cacodylate buffer at pH 7.4. If neces-sary, adjust the solution at the desired pH value by adding some drops of 0.1 N
HCl (see Note 1). Dissolve 2% paraformaldehyde in cacodylate (Note 2). The
weighed amount of paraformaldehyde is warmed in about half the final volume
of cacodylate buffer at a temperature of 55–60°C. This solution is then cooled
under running tap water before addition of glutaraldehyde. Add 5% glutaralde-hyde (Note 3) and bring the fixation solution to the final volume. As shown in
Damage to Mitochondria 223
Fig. 1, a simple apparatus to carry out fixation by perfusion can be easily assem-bled. In essence, it consists of an infusion set with two plastic flasks, 4 m of
small-sized plastic, transparent tubes, one Y-shaped tube, two clamps, and one
perfusion needle.
3. Osmium tetroxide preparation (see Note 4): Remove the label from the ampoule
containing the osmium tetroxide crystals, clean the ampoule carefully with etha-nol to erase debris of any kind due to the label, and wash with twice-distilled
water. A glass-stopped bottle, a glass tube, and a heavy glass rod must be cleaned
with concentrated nitric acid to remove the organic matter. Twice-distilled water
should be used to wash out traces of the acid. Never wipe with paper or cloth
towel, as they can let loose particles that reduce the solution to hydrated dioxide.
Add to the bottle a measured amount of twice-distilled water; place the ampoule
containing osmium tetroxide crystals gently in the bottle; use the glass tube and
the heavy glass rod to break the osmium ampoule and then quickly stop and shake
the bottle vigorously. Complete dissolution of osmium crystals in the water takes
at least half an hour. Wrap the bottle with aluminum foil and store in refrigerator
Fig. 1. This simple apparatus for vascular perfusion can be easily assembled using
inexpensive materials. Plastic flasks can be adapted to contain saline (I) and fixation
solution (II). (A,B) Clamps to stop either the saline or the fixative flow. (C) Y-Shaped
tube to regulate both flows. (D) Infusion needle with rounded tip.
224 Miquel
at 4°C. Protect this solution from exposure to light. Because of the high volatility
of osmium tetroxide, there is a rapid decrease in the concentration of its solu-tions. Therefore, they should be prepared in small quantities and stored in flasks
fitted with a glass stopper and Teflon sleeve (Note 5). We dissolve 1 g of osmium
tetroxide in 33 mL of twice-distilled water, and then this solution is diluted (usu-ally to 1%) and used according to our needs within a couple of weeks (Note 6).
4. Ethanol as a dehydrating agent for electron microscopy: The solutions with dif-ferent concentrations of ethanol (at 70% and 80%) should be freshly prepared
before their use by adding the proper volume of twice-distilled water. Ethanol at
95% for electron microscopy is commercially available at a reasonable price and
good purity. Therefore, it is advantageous to buy it instead of preparing a solution
starting from ethanol at 100%.
5. Propylene oxide: Used as transitional solvent from ethanol to the embedding
6. Epoxy resin (Araldite) preparation (Note 7): Araldite, a yellowish, transparent
epoxy resin, is prepared by mixing the following amounts of four components
(Note 8):
Araldite (Note 9) 10 mL
Dodecenyl succinic anhydride (DDSA) 10 mL
Dimetilaminometil-fenol (DMP-30) 0.5 mL
Dibutil-ftalate (Note 10)1 mL
7. Embedding molds.
8. Laboratory oven with facilities to program times and temperatures.
9. Knifemaker, glass strips, electric tape, copper grids, tweezers for electron micro-scopy, Petri dishes, filter paper, twice-distilled water, dental wax.
10. Preparation of glass knives (Note 11): Glass knives are made from glass strips
(thickness, 0.5 mm). They are prepared (just prior to usage) by means of a
knifemaker, a mechanical device that allows to obtain clean breaks, provided that
the glass strip is firmly blocked, First, the glass strip must be broken into several
squares, which, in turn, are scored and broken into two knives (Fig. 2) (Note 12).
11. Preparation of troughs: Choose a glass knife of good quality and wrap a piece of
adhesive electrical tape (Note 13) from one side to the other of the cutting edge of
the glass knife (Fig. 2) (Note 14). Seal the heel of the trough with melted paraffin
to make it waterproof. The trough is then mounted on the proper holder of the
ultramicrotome and it is filled up with distilled water, as flotation fluid (Note 15).
12. Ultramicrotome(s).
13. Uranyl acetate preparation: This chemical is supplied as a powder and it is used
as 3% aqueous or ethanol solution. It takes about 20 min to dissolve it completely
in water on an automatic shaker (Note 16).
14. Lead citrate preparation: This staining solution is prepared as follows in a 50-mL
volumetric flask: lead nitrate (1.33 g), sodium citrate (1.76 g), and twice-distilled
water (30 mL).
This mixture must be shaken vigorously several times for at least 30 min until
a milky solution is obtained. Add to this suspension 8 mL of 1 N NaOH previ-
Damage to Mitochondria 225
ously diluted with twice-distilled water to 50 mL and mix until the suspension
clears up completely (Note 17).
15. Transmission electron microscope.
3. Methods
Routine preparation of biological samples to be studied by electron micros-copy is carried out through the following steps: animal anesthesia, perfusion–
fixation, embedding, sectioning, and staining of the tissue samples.
Fig. 2. Upper left: A trimmed Araldite block ready to be sectioned at 0.06 µm thick-ness. The tissue specimen at the tip of the block can be clearly seen in osmium-treated
samples. Upper right: (1) Squares are obtained by scoring and breaking a glass strip
perpendicularly to its longitudinal axis. (2) Squares are scored and broken to obtain
knives (g: see Note 43). Lower left: a trough is made with a piece of electrical tape
wrapped around the cutting edge and sealed at the through heel. Lower right: A section
ribbon floating at the center of the trough provides evidence that the entire sectioning
procedure has been properly accomplished.
226 Miquel
3.1. Animal Anesthesia
Medium-sized laboratory animals (adult Wistar rats, average weight 250 g)
are easily anesthesized by injection with 200 mg/kg body weight of 2,2,2-tri-bromoethanol dissolved in 10% ethanol, which induces sleep within 5–7 min.
3.2. Perfusion
The thorax of the anesthesized animal is opened and the heart and major
blood vessels are exposed. A small incision is made in the left ventricle to
guide the perfusion needle to the aorta. As soon as the saline (to prevent clot-ting, a heparinized solution should be used) starts to flow (Note 18), a second
incision is made in the auricle to allow the blood to escape (Note 19). The
saline must flow up to a complete washout of the blood and should be followed
immediately by perfusion of the fixation solution (Note 20). The selected tis-sue samples are dissected out and a further fixation of the material is performed
by immersion in the same solution used to perfuse the animal (Note 21).
3.3. Embedding
Following fixation, to minimize any reaction between the fixative and the
dehydration agent, the excess fixative must be washed off (see Note 22). Wash-ing should be carried out in the same buffer as that used to prepare the fixative
mixture (see Note 23). For most tissues, two or three rinses in the buffer, for a
total period of 15 min, are needed (see Note 24). If fixation has been carried
out in a refrigerator at 4°C, wash the tissue samples at the same temperature.
Wide-mouth vials are suitable to process the tiny samples, which should not
be transferred from one vial to another. Each solution must be withdrawn
with a micropipet and immediately replaced with the other solution (Note 25).
The 3% aqueous solution of osmium tetroxide is diluted to 1% with cacodylate
buffer. This fresh-made reagent is poured into the vials containing the tissue
samples (Note 26). Immersion in osmium tetroxide lasts 2 h and is carried out
in a refrigerator at 4°C to reduce autolytic activity (Note 27). Dehydration by
ethanol follows osmium tetroxide postfixation (Note 28) according to the
following procedure:
1. Rapid wash in cacodylate buffer.
2. 70% Ethanol (Note 29): 10 min.
3. 80% Ethanol: 10 min.
4. 95% Ethanol: 10 min.
5. 100% Ethanol: 15 min (two changes).
Because most resins are not readily miscible with ethanol, tisssue samples
must be immersed in a transitional solvent that is completely miscible with
both ethanol and embedding resins, according to the following procedure:
Damage to Mitochondria 227
1. 100% propylene oxide: 20 min (three changes).
2. Propylene oxide and embedding resin (3:1): 30 min.
3. Propylene oxide and embedding resin (1:1): 30 min.
4. Propylene oxide and embedding resin (1:3): overnight (Note 30).
5. Embedding resin 100% (Note 31): 30 min.
Final embedding is carried out in rubber molds of different size which allow
an easy orientation of the tissue samples. Polymerization of the epoxy resin is com-pleted within 48 h at 60°C. Polymerized blocks can be sectioned best after 2 d.
3.4. Sectioning (Fig. 2)
Because sections are cut from a very small portion of the tissue specimens,
trimming of the blocks must be carried out. The block is mounted on the block-holder which is placed in the microtome (Note 32). Rough trimming of the
block must be performed with a sharp razor blade to remove the excess of
embedding medium around the tissue sample. Too large tissue samples should
be also reduced in size to obtain a block face of suitable size. Afterwards, fine
trimming is carried out with the aim of obtaining a block tip with smooth and
clean sides. In our experience, when making sections from a brain sample, it is
usually necessary to examine a semithin section (thickness: 1.5 µm) at the opti-cal microscope to select the area to be viewed at the electron microscope. When
the area of interest in the tissue sample is identified, the block is retrimmed to
obtain a trapezoid face with parallel lower and upper sides and two sloped
sides (Note 33). Because the shortest of the two parallel edges is mounted up,
each section will move the previous one from the cutting edge and a straight
ribbon of sections will be formed (Fig. 2; Note 15). Section collection (Note
34) is accomplished by the following steps:
1. Hold the edge of a grid with a pair of curved tweezers.
2. Bend the edge of the grid by about 100°.
3. Lower the grid onto the upper face of the ribbon (Note 35).
4. Dry the grid with the sections by placing it on filter paper which absorbs the
water (of course the side of the grid without sections is to be brought into contact
with the filter paper!) (Note 36).
3.5. Staining
To obtain a better contrast and resolution, the sections mounted on a grid are
stained by the deposition of heavy metal ions, such as uranium and lead (see
Note 37). The simplest method to stain sections is to let the grid (with section
side down) float on a drop of a staining solution in a Petri dish. The staining
surface is constituted by the lid of a glass Petri dish partially filled with melted
paraffin which is allowed to cool. This surface must be kept very clean and
should be protected from dust by keeping it covered during staining. The basic
228 Miquel
steps to carry out a conventional double staining with uranyl acetate (ethanol
solution) and lead citrate are:
1. An ethanol atmosphere is produced within the Petri dish by placing a piece of
filter paper wetted with ethanol 50% at one side of the staining surface.
2. A small volume of uranyl acetate is drawn up from below the surface of the stain-ing solution with the aid of a clean pipet and a small drop is placed on the paraffin
surface (Note 38).
3. The grid to be stained is wetted in distilled water (Note 39) and then floated on
the drop of uranyl acetate, where it should be left from 30 s to a few minutes
depending on whether or not it must undergo multiple staining.
4. The stained grid is then thoroughly washed by immersion in a small beaker con-taining a 50% aqueous solution of ethanol for at least 5 min. Washing can also be
carried out by holding the grid with tweezers and dipping and shaking it in the
solution, in the beaker (Note 40).
5. The washed grid is then dried by placing it on a filter paper, taking care that the
section side does not touch the paper.
6. Steps 1–5 can be repeated to stain the same grid with lead citrate. However, as
regards step 1, a carbon dioxide free atmosphere must be generated within the
Petri dish by placing a small amount of NaOH pellets at one side of the staining
surface. Washing should be carried out using twice-distilled water. Usually, the
duration of the lead staining (5–15 min) is longer than that with uranyl acetate
(Note 41).
3.6. Quantitative Mitochondrial Study
A mere qualitative evaluation by electron microscopy of tissue samples is
not enough to document reliably the age-related changes. Therefore, quantita-tive fine-structural determinations must be performed. With the introduction of
the dissector procedure (8–10), the methods of estimating subcellular particle
and organelle number and size have been changed, with a considerable improve-ment of the reliability of the findings by moving from assumption-dependent
to assumption-free methods (Note 42). Thus, suppositions about particle size,
shape, and orientation have been replaced by design-based random sampling,
which is a well grounded and reliable procedure to carry out quantitations. It
must be pointed out that, because these unbiased stereological techniques have
entered into common usage during the last decade, most published morpho-metric data have been obtained without the use of the dissector. This does not
mean that useful information was not obtained. Thus, to cite an example, in our
previous studies on the ultrastructure of nerve cell terminals in aged rats, the
morphometric techniques in use prior to the dissector method enabled us to
measure closely related structural parameters of functional significance. Thus,
an estimation of the remodelling process taking place in the synaptic regions of
the aged CNS was obtained. Namely we found that, while the numerical den-
Damage to Mitochondria 229
sity (no. of particles/µm3 of tissue) of synaptic mitochondria decreases signifi-cantly with aging, their average size (average mitochondrial volume) increases
to reach a complete recovery of the total volume in a µm3 of tissue (volume
density) (8–10). These findings appear to be significant if referred to the lim-ited specific volume where sampling has been carried out. In conclusion, it
seems reasonable to assume that the data obtained by many laboratories prior
to the introduction of the dissector procedure may still be trusted when the
conclusions drawn refer not to quantitation of the total number of structures
but to the relationships present among the structures (e.g., number of mito-chondria found in a synapse cross-section).
4. Notes
1. The buffer solutions should be prepared the day before their use. Freshly pre-pared solutions are stable for a week if stored at 4°C.
2. To prevent spontaneous polymerization during storage, commercial formalin con-tains more than 10% methanol. Thus, to yield a high-purity solution, it is pre-ferred to prepare the fixative for electron microscopy by depolymerizing
powdered paraformaldehyde.
3. The commercially available glutaraldehydes have varying amounts of impurities
which can affect tissue fixation to a high extent. Although it is more expensive, the
use of glutaraldehyde of optimal purity is more advantageous to obtain a good
preservation of biological samples. Glutaraldehyde is supplied as a 50% or 25%
solution that stays relatively stable for long periods of time if stored in the cold. If
the solution turns yellow, some polymerization has occurred and the chemical
should not be used unless it is purified with charcoal. However, the purification
procedure implies that the concentration of the final clear, oily solution must be
checked with a recording spectrometer. Therefore, we recommend purchasing the
right amount of glutaraldehyde to be used in a reasonably short period of time.
4. Osmium tetroxide is volatile, and its fumes are very toxic and able to fix biologi-cal tissues. Thus, any manipulation involving this chemical should be performed
in a fume cupboard. Exposure to the vapor should be kept to an absolute mini-mum and contact with hands and face should be avoided. Great care should be
taken to avoid any contamination by dust particles or light, which would result in
a striking decrease in the fixing capacity of the solution.
5. Osmium vapors readily leak out of many containers, and in many electron micro-scopy laboratories a typical sign of this contamination is the black precipitate
found on the inside of refrigerator walls. To prevent this leakage, osmium solu-tions should be stored in glass vacuum-type blood collection tubes. Plugging them
with rubber stoppers prevents contamination of the surrounding environment.
6. Before disposing of the used solutions of osmium tetroxide, neutralize them by
twice the volume of corn oil, a product that has a high percentage of unsaturated
bonds. Confirmation that the toxic osmium tetroxide has been completely inacti-vated can be obtained by soaking a piece of filter paper in corn oil and then
230 Miquel
suspending it over the solution. Blackening indicates that osmium tetroxide is
7. Araldite (as all epoxy resins) and propylene oxide are toxic. Therefore, any
manipulation involving these chemicals should be carried out under a fume
cupboard and wearing lattice gloves.
8. To facilitate the preparation of Araldite, the needed amount of the different com-ponents can be measured in grams instead of milliliters. Large amounts of
Araldite can be prepared to save time and disposable laboratory ware as well as
to minimize any mistake in the weighing of the chemicals. Routinely, we prepare
500 g of Araldite, storing the unused quantity in syringes (of 5, 10, and 20 mL) at
–20°C. Great care must be taken in filling each syringe with resin: remove the tip
cover, slowly draw in the resin, hold the syringe with the tip in an upward posi-tion, and remove eventual air bubbles by pulling the plunger slowly; push the
plunger upward until the resin appears at the end of the tip, cover the tip with
parafilm, and store in a refrigerator at –20°C. Thirty minutes before its use, the
required amount of Araldite must be warmed up to room temperature.
9. After pouring Araldite and DDSA, the mouth of their bottles must be carefully
wiped with filter paper, because with time, even at room temperature, small
residual amounts of these reagents can polymerize and cause a strong bonding
between the screw plug and the neck of the bottle.
10. The hardness of the blocks to be sectioned can be varied by changing the amount
of dibutyl phthalate (plasticizer); however, a low amount of this reagent will result
in brittle blocks.
11. Diamond knives can be used instead of glass knives. Diamond knives are com-mercially available at a rather high cost, but they are convenient as they are
supplied already sealed to a stainless steel or aluminum alloy trough. The shape
and color (usually black) of this trough make it possible to obtain an optimal
meniscus level (in relation to the cutting edge).
12. The quality of each glass knife must be checked according to the instructions for
using the knifemaker. The most useful (and sharpest) part of the knife is
represented by the left third of the whole cutting edge.
13. The fingers should not come into contact with the adhesive side of the tape, as
grease will be transferred to the flotation water. The black (or silver) color of the
tape will facilitate the viewing of the sections on the water surface, as these colors
can prevent interference due to reflection.
14. The lower edge of the tape must be parallel to the lower edge of the knife and the
upper edge of the tape must be precisely attached to the tip of the cutting edge.
These steps should be carried out with special care, as the resulting trough should
not be tilted toward or away from the cutting edge. The optimal trough will make
it possible to obtain a good meniscus level of the flotation water. As a conse-quence, the sections will glide smoothly away from the cutting edge and thick-ness can be easily checked on the basis of their interference color.
15. A straight ribbon of sections is easily obtained if the flotation water in the trough
is wetting the cutting edge of the knife to its very end. The shape of the meniscus
Damage to Mitochondria 231
of the flotation water is primarily responsible for the wetting of the knife facet.
The optimal wetting of the knife is obtained when the meniscus is flat (with a
knife angle of about 90°). To obtain the most advantageous meniscus level, first a
convex meniscus is developed and then, by withdrawing the water with a micropi-pet up to an initial drying of the cutting edge, the meniscus becomes concave.
16. A freshly prepared solution of uranyl acetate should be used, as the recommended
3% solution is close to saturation and gradually a precipitate forms with time.
However, in our experience, the solution can be used at least for a couple of
weeks, until it looks slightly cloudy.
17. The lead citrate solution can be used concentrated or diluted up to 5× with 0.01 N
NaOH according to the contrast to be obtained. To prepare lead citrate, the NaOH
solution should be fresh and carbonate free. If it is tightly stopped, the solution
will stay stable enough for 6–8 wk. When precipitates appear, the solution must
be discarded.
18. The blood washout by saline should be carried out in a short period of time (2–4
min) to minimize any tissue deterioration and ensure the best fixative preserva-tion.
19. The temperature of the perfusion solution should be kept as close to body tem-perature as possible to prevent vasoconstriction. The pressure head of the solu-tions (saline and the aldehyde-containing fixative), especially for fixation of CNS
structures, must be equivalent to 130–140 cm of water. This can be easily set up
by positioning the perfusion bottles shown in Fig. 1 about 130 cm higher than the
body of the animal to be perfused.
20. A fixation solution containing the right amounts of glutaraldehyde and paraform-aldehyde is more adequate than a solution containing just one of these aldehydes.
This is due to the complementary effects of the two chemicals: formaldehyde
diffuses rapidly and fixes slowly, whereas the glutaraldehyde action is character-ized by a slow penetration and a rapid fixation of tissues, due to the crosslinking
effects on the adjacent protein molecules.
21. The uniformity of fixation throughout the tissue sample is a good criterion for
adequate fixation. The size of the specimen exerts an important influence on the
homogeneity of fixation. As a general rule, the smaller the size of the biological
sample, the more complete and uniform will be the quality of fixation. With the
above described perfusion procedure, a nearly uniform fixation is obtained. How-ever, the size of the samples should not exceed 0.5 mm3, as osmium tetroxide
penetrates little in most tissues. Perfused tissues can be easily cut into thin strips
of about 0.5 mm thickness and then sectioned into smaller pieces before osmium
22. If washing of the tissue samples is improperly carried out, residual glutaralde-hyde will react with osmium tetroxide and produce a fine, dense precipitate of
reduced osmium.
23. If tissues fixed in buffered solutions are washed with water, dissolution and pro-gressive disintegration of some unfixed cellular structures takes place when the
added electrolytes (or nonelectrolytes) are removed by washing with water. By
232 Miquel
contrast, washing with buffer avoids a sudden, drastic change in the environment
of the tissue specimens. Because cellular membranes of aldehyde-fixed samples
maintain in part their selective permeability, a significant decrease in the osmo-larity of the washing solution would result in a marked ballooning of cellular
organelles, particularly mitochondria.
24. Washing should not be longer than necessary, as the crosslinks caused by glut-araldehyde are potentially reversible and therefore long washings may affect the
final appearance of the fine structure.
25. Solution changes (during fixation, washing, dehydration, and infiltration) should
be carried out with speed to avoid drying of the samples. The original vial serves
to perform fixation up to the transfer of the specimens to rubber molds, for
26. Because of the hypotonicity of the water in which osmium tetroxide is dissolved,
animal tissues exhibit a marked and rapid swelling when they are immersed in
the final buffer-diluted 1% solution of this reagent. However, this swelling is
neutralized by the shrinkage occurring during dehydration.
27. Rapid osmium tetroxide penetration lasts up to 1 h and reaches a depth of about
0.6 mm. After this period, the fixed outer layers of the tissue samples resist deeper
penetration of the fixative and the concentration of osmium tetroxide in solution
decreases. Consequently, the rate of penetration of the fixative slows down and it
takes 2 h to fix throughout a tissue sample of 1 mm thickness. Osmium tetroxide
is unable to make all cellular components insoluble in water and extraction of
cellular products (proteins) may occur during prolonged fixation. Thus, short fixa-tion times are more suitable, considering that the fixation time can be increased
when osmium tetroxide is used in postfixation.
28. Provided that the duration of each step is long enough to accomplish a gradual
replacement of water by the solvent, dehydration should be as short as possible to
minimize shrinkage and extraction of cellular constituents. A rapid dehydration
causes striking osmotic changes resulting in distortion of the structures. The dehy-dration time required will depend on the size and type of the tissue specimen.
29. Even in fixed tissue samples, the possible presence of soluble proteins is to be
taken into account and the concentration of ethanol at the start of dehydration
should be high enough to denature (and insolubilize) these proteins.
30. Because of the high volatility of propylene oxide, the vials containing the third
mixture (1:3, overnight) must be sealed carefully with a stopper and parafilm.
31. Residual propylene oxide can be completely eliminated from tissue samples by
storing for 2–4 min the open vials with 100% embedding medium into an oven
prewarmed at 60°C for subsequent polymerization.
32. To keep the trimmed block firmly attached to its holder, the block should not
extend more than 2–3 mm from the front edges of the holder jaws. The evident
results of a loose block are sections of uneven thickness, alternation of thick and
thin sections, or even total failure of sectioning.
33. Trimming of the block should be performed avoiding sharp, deep cuts. To obtain
the form of a short pyramid, the block should be trimmed by short cuts and its tip
Damage to Mitochondria 233
should not be too thin nor too long. Otherwise, it will vibrate at high frequency
when hitting the cutting edge.
34. Before collecting the sections, the compression due to the impact of the block
with the cutting edge of the knife can be relieved by exposing the sections very
briefly to the vapors of a strong solvent such as chloroform.
35. The grid should be pressed gently over the floating sections with the aim of
depressing (not breaking) the water surface. As soon as the grid reaches across a
ribbon, the sections stick firmly to the grid along with a drop of water. Because
the electrostatic charges on the grid surface are responsible for the attraction of
the ribbon, rinse the grid with acetone just prior to use, to reduce the charge and
get a better control in orienting and centering the ribbon.
36. The most frequent mistake in making sections for electron microscopy is
represented by periodic variations in the contrast of a section (commonly termed
“chatter”). Usually, this failure is not detected during sectioning, but only while
viewing the section at the electron microscope when it is too late to correct
the mistake. Usually, “chatter” is due to vibrations of microtome parts that include
the cutting edge, the specimen block, and the microtome arm. However, some
other causes may be involved, for example, too fast cutting speed, too large and/or
irregular block face, specimen or knife loosely held, too hard or too soft specimen
block, dull knife, and incomplete polymerization of the epoxy resin.
37. Contrast in an electron micrograph depends essentially on the density and
thickness of the different parts of the section. By combination with elements of
higher atomic weight (osmium, uranium, and lead), some cellular structures are
made denser than their surroundings.
38. The first one or two drops of stain from the pipet are discarded to avoid any
contamination. The drop used should be small enough to allow the grid to float
on it instead of sliding down to its sides.
39. Wetting of the sections decreases the risk of contamination due to extensive
stain–air contact.
40. To remove any excess of uranyl acetate, a quick rinse in 0.5% tartaric acid can be
performed after washing in a 50% aqueous solution of ethanol. This step must be
carried out very rapidly (2–3 s!) and it must be followed by repeated washings in
distilled water.
41. Keep the staining time as short as possible because there is a risk of overstaining,
which results in an overall increase in contrast with poor differentiation of cellu-lar structures.
42. The dissector method is designed to identify particles seen in one section (“look-up section”) but not in a following one (“reference section”), and serves to evalu-ate the number of particles per unit volume (numerical density: Nv). Serial
sections are needed to perform counting of particles or organelles. For instance,
the mitochondria present in a discrete area of the reference section, but not in the
look-up section are counted (Fig. 3, arrows) and referred to the height of each
dissector which takes into account the section thickness (further details in refs.
234 Miquel
43. When preparing knives, do not touch their sides with the fingers. When making
squares from the glass strip with the diamond-tipped shaft of the knifemaker,
check that the rough end of the strip (g) is positioned upward, whereas when
making knives it should be turned downward.
Dr. J. Miguel acknowledges the support of FIS-Grant 99/1264.r. The skillful
help of Mr. M. Solazzi and Mrs. F. Trucchia is greatly acknowledged by Dr. C.
l. Harman, D. (1956) Aging: a theory based on free radical and radiation chemistry.
J. Gerontol. 11, 298–300.
2. Gerschman, R. (1962) Man’s dependence on the earthly atmosphere, in Proceed-ings of the 1st Symposium on Submarine and Space Medicine (Schaeffer, K. S.,
ed.), MacMillan, New York, pp. 475.
3. Miquel, J. (1971) Aging of male Drosophila melanogaster: histological, histochemi-cal and ultrastructural observations, in Advances in Gerontological Research, vol.
3 (Strehler, B. L., ed.), Academic Press, London, New York, pp. 39–71.
Fig. 3. Electron microscopy pictures of rat cerebellar cortex processed according to
the procedure described in the text. These photos constitute a dissector, that is, A is the
“look-up” section, while B is the “reference” section. Arrows indicate the mitochon-dria which are present in A but not in B and therefore those to be counted. ×22,000.
Damage to Mitochondria 235
4. Miquel, J., Lundgren, P. R., and Johnson, J. E., Jr. (1978) Spectrophotometric and
electron microscopic study of lipofuscin accumulation in the testis of aging mice.
J. Gerontol. 33, 5–19.
5. Miquel, J. and Fleming, J. E. (1986) Theoretical end experimental support for an
“oxygen radical-mitochondrial injury” hypothesis of cell aging, in Free Radicals,
Aging and Degenerative Diseases (Johnson, J. E., Jr., Walford, R., Harman, D., and
Miquel, J., eds.), Alan R. Liss, New York, pp. 51–74.
6. Miquel, J., editor-in-chief (1989)  CRC Handbook of Free Radicals and Antioxi-dants in Biomedicine (3 vols.), CRC Press, Boca Raton, FL.
7. Miquel, J. (1998) An update on the oxygen stress-mitochondrial mutation theory of
aging: genetic and evolutionary implications. Exp. Gerontol. 33, 113–126.
8. Bertoni-Freddari, C., Fattoretti, P., Casoli, T., Spagna, C., Meier-Ruge, W., and
Ulrich, J. (1993) Morphological plasticity of synaptic mitochondria during aging.
Brain Res. 628, 193–200.
9. Bertoni-Freddari, C., Fattoretti, P., Caselli, U., Paoloni, R., and Meier-Ruge, W. (1996)
Age-dependent decrease in the activity of succinic dehydrogenase in rat CA1 pyra-midal cells: a quantitative cytochemical study. Mech. Ageing Dev. 90, 53–62.
10. Fattoretti, P., Bertoni-Fredari, C., Caselli, U., Paoloni, R., and Meier-Ruge, W. (1998)
Impaired succinic dehydrogenase activity of rat Purkinje cell mitochondria during
aging. Mech. Ageing Dev., 101, 175–182.
11. Sterio, D. C. (1984) The unbiased estimation of number and sizes of arbitrary par-ticles using the dissector. J. Microsc. 134, 127–136.
12. Mayew, T. M. and Gundersen, H. J. G. (1996) “If you assume, you can make an ass
out of you and me”: a decade of the dissector for stereological counting of particles
in 3D space. J. Anat. 188, 1–15.
13. Coggeshall, R. E. (1992) A consideration of neural counting methods. Trends Neurol.
Sci. 15, 9–13.
Damage to Mitochondria 237
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Causes and Consequences
of Damage to Mitochondria
Study of Functional Aspects by Flow Cytometry
Federico V. Pallardo, Juan Sastre, Jaime Miquel, and José Viña
1. Introduction
A rapidly increasing amount of data supports the view that progressive
bioenergetic loss caused by injury of the main energy-producing subcellular
organelles, that is, the mitochondria, plays a key role in aging. A link between
senescence and energy loss is already implied in Harman’s  (1) free radical
theory of aging, according to which oxygen-derived free radicals injure the
cells, with concomitant impairment of performance at the cellular and physi-ological levels. Further, Miquel and co-workers (2,3) have proposed a  mito-chondrial theory of aging, according to which aging results from oxygen stress
damage to the mitochondrial genome, with concomitant bioenergetic decline.
More recently, a number of laboratories, including our own (4–6), have pro-vided biochemical data in agreement with the above views. Thus, we have
shown that, as the result of age-related oxygen stress, mitochondrial glutathione
is oxidized in direct relation to injury of the mitochondrial DNA (5). Further,
our studies suggest that an antioxidant product extracted from Ginkgo biloba
may counteract in part the damaging effects of free radicals on mitochondrial
and cellular aging (7).
As reviewed elsewhere (6), age-related functional changes in mitochondrial
respiration and in transport systems have been reported. Nevertheless, because
of differences in the techniques and biological aging models used, the literature
on mitochondrial aging abounds in conflicting reports. Further, it is often diffi-cult to assess the functional significance of the mitochondrial changes shown by
standard biochemical techniques. This makes it advisable to study the effects of
238 Pallardo et al.
age on mitochondria by flow cytometry methods which allow a direct measure-ment of the organellar function as shown by its membrane potential value. This
value can be determined both on mitochondria isolated using standard tech-niques and on those that remain in their normal environment inside the cells.
Our own mitochondrial flow cytometry studies (6) on isolated rat hepatocytes
have shown, for the first time in intact cells, a correlation between age-related
changes in cell size and impaired mitochondrial function (Fig. 1). Specifically,
we have observed that age is accompanied by a decrease in mitochondrial mem-brane potential (MMP), an increase in mitochondrial size, and a loss of organellar
homeostasis (resulting in raised levels of peroxide generation). The pathoge-netic mechanisms responsible for these changes are not well understood, although
as pointed out by Hagen et al. (8), loss of the membrane phospholipid cardio-lipin (which is essential for respiratory chain work) may play a role in the age-related MMP decrease. Because MMP supports mitochondrial protein synthesis
(9), its decrease with age may be linked to a general decline in the biochemical
and functional competence of mitochondria (2).
It is well known that aging results in mitochondrial membrane changes that
increase the vulnerability of these organelles to the stress caused by the isola-tion procedure. Therefore, during isolation a considerable number of organelles
(precisely those most altered by aging) may be lost, and therefore the data
obtained will not provide an accurate measurement of the functional state of
the whole mitochondrial population. Nevertheless, because useful information
can be obtained on isolated mitochondria, we will describe the methods used in
our laboratory for flow cytometry study of isolated mitochondria and of mito-chondria present in their normal environment in cells previously isolated from
their tissue. A detailed presentation of these methods may be of interest to
workers interested in the mechanisms of senescence of mitochondria of differ-ent cell types, changes in the rate of aging caused by genetic or environmental
modulation of mitochondrial development and function, and protection of the
mitochondrial membranes and DNA by dietary antioxidants and free radical
The procedures presented here for the study of key parameters of mitochon-drial membrane function that change with age are as follows: (1) preparation of
the respiration buffer for suspension of isolated mitochondria or cells and (2)
determination of mitochondrial membrane mass, membrane potential, mem-brane lipid composition, and peroxide production.
Of course, a more complete understanding of the causes and effects of
mitochondrial aging will require determination of many other parameters,
such as fine structural changes (see Chapter 17), NADH/NAD redox state,
reduced and oxidized thiol contents, activity of the inner membrane respi-ratory enzymes, state 3/state 4 respiratory control ratios (which indicate
Damage to Mitochondria 239
the level of coupling of mitochondrial electron transport to ATP produc-tion), and oxidative damage and deletions of mtDNA. A number of recent
publications (including refs. 5–8 and 10–12) provide examples of methods
and age-related research on these parameters.
Fig. 1. Examples of application of flow cytometric methods to study mitochondrial
function in isolated whole cells (hepatocytes) of young (aged 3–4 mo) and old (aged
22–36 mo) male Wistar rats. (A) Distribution of mitochondrial size obtained using the
forward-angle light scatter. (The arbitrary units correlate positively with size.)  (B)
Peroxide generation in mitochondria (isolated from the above young and old rats)
measured with the DhRh123 technique. (Reproduced from ref. 6).
240 Pallardo et al.
2. Materials
1. Buffer for mitochondria or cell suspensions (“buffer,” pH 7.4): Solution in twice-distilled water of 5 mmol of KH2PO4, 0.3 mol of sucrose, 1 mmol of ethylene
glycol bis(β-aminoethyl ether)-N,N,N’,N’,-tetraacetic acid (EGTA), 5 mmol of
morpholinopropanesulfonic acid (MOPS), and 0.1% bovine serum albumin.
2. Stock solution of nonyl-acridine orange (NAO) (high purity, from Molecular Probes)
at 10 mg/mL of dimethylformamide. (Store at –20°C, protected from light.) (13).
3. Stock solution of rhodamine 123 (Rh123) (from SIGMA) at 5 mg/mL of
dimethylformamide. (Store at –20°C in 20  µL aliquots, using Eppendorf tubes
protected from light) (14).
4. Stock solution of nile red (from Molecular Probes) at 2 mg/10 mL of ethanol.
(Store at –20°C in 20-µL aliquots, using Eppendorf tubes protected from light.) (15).
5. Stock solution of dihydrorhodamine 123 (DhRh123) (from Sigma) at 10 mg/mL
of dimethylformamide. (Store at –20°C, protected from light.) (16).
6. EPICS ELITE cell sorter (Coulter Electronics, Hialeah, FL, USA).
3. Methods
3.1. Flow Cytometry
1. Flow cytometry tubes: Polypropylene tubes (3 mL).
2. Flow cytometry parameters: Sample volume = 50 µL; sample flow rate = 25 µL/min;
sheath pressure = 7.50 psi; laser power = 400 mW; parameter voltages: SS = 711 V;
FL1 = 1750 V; FL2 = 0 V; FL3 = 0 V; color compensation: FL1 = 5% of FL2; FL2
= 50% of FL3; FL2 = 70% of FL1; FL2 = 0% of FL3; FL3 = 43% of FL1; FL3
= 1% of FL2.
3. Windows: For all parameters used (LFS, FSL, FL1, and LSS), the maximal
window was 1023.
4. General fluorimetric procedure: The stained cells or mitochondria pass through
the flow chamber, where the fluorochromes are excited with an argon laser tuned
at 488 nm. Forward-angle and right-angle light scatter are measured and fluores-cence is detected through a 488 nm blocking filter, a 550 nm long-pass dichroic,
and a 525 nm band pass or a 575 nm long pass.
3.2. Sampling of Isolated Cells (see Note 1)
1. Prepare cellular suspensions containing approx 300,000 cells/mL. (Cell viability
is determined by the fluorescent dye propidium iodide at a final concentration of
5 µg/mL, at 630 nm fluorescence emission, by light-scatter properties. All studies
should be performed on viable cells.)
2. Place 50 µL of the suspension in the flow cytometry tube and mix with “buffer”
and reagents needed for each specific determination, as indicated below (Sub-headings 3.4. and 3.5.).
3.3. Sampling of Isolated Mitochondria (see Note 1).
1. Suspend gently in 2 mL of “buffer” a sample of the mitochondrial pellet contain-ing 1 mg of mitochondrial protein.
Damage to Mitochondria 241
2. To obtain the “working suspension,” mix 100 µL of the above suspension with
1900 µL of “buffer.”
3. Place 50 µL of the “working suspension” in the flow cytometry tubes and mix with
“buffer” and reagents needed for each specific determination, as indicated below
(Subheadings 3.4.–3.7.).
3.4. Mitochondrial Membrane Mass
This is measured with NAO at 525 ± 5 nm fluorescence emission (see Notes
2 and 3).
1. Dilute the NAO stock solution with “buffer” to a final concentration of 1 mg/mL
(“working solution”).
2. Place 10 µL of “working solution” in a flow cytometry tube.
3. Add 940 µL of “buffer.”
4. Add 50 µL of cell or mitochondria suspension.
5. Incubate in a water bath at 37°C for 20 min, protecting from light.
6. Assay in a flow cytometer.
3.5. Mitochondrial Membrane Potential
This is determined measuring Rh123 fluorescent emission at 525  ± 5 nm
(see Notes 4–7).
1. Mix 10 µL of Rh123 stock solution with 990 µL of “buffer” to obtain the “diluted
2. Add 10 µL of “diluted solution” to 90 µL of “buffer,” to prepare the “working
3. Place 10 µL of “working solution” in a flow cytometry tube.
4. Add 940 µL of “buffer.”
5. Add 50 µL of cell or mitochondria suspension.
6. Incubate at 37°C for 20 min, protecting from light.
7. Assay in a flow cytometer.
3.6. Mitochondrial Membrane Lipid Composition
Composition of polar and apolar lipids in the mitochondrial membranes is
assessed by fluorescence emission of nile red.
1. Place 10 µL of stock Nile solution in a flow cytometry tube.
2. Add 940 µL of “buffer.”
3. Add 50 µL of mitochondrial suspension.
4. Incubate at 37°C for 30 min, protecting from light.
5. Assay in a flow cytometer at 525 (green), 575 (orange), and 675 (red) nm.
3.7. Peroxide Production by Mitochondria (see Note 8)
For this determination, the fluorochrome DhRh123 is excited at 488 nm.
This fluorophore is oxidized by H2O 2-dependent reactions involving oxygen
species (16) (see Note 5).
242 Pallardo et al.
1. Mix 5  µL of DhRh123 stock solution with 495  µL of “buffer,” to prepare the
“diluted solution.”
2. Mix 5 µL of “diluted solution” with 95 µL of “buffer,” to prepare the “working
3. Place in a flow cytometry tube 10 µL of “working solution,” 940 µL of “buffer,”
and 50 µL of mitochondrial suspension.
4. Incubate at 37°C for 30 min, protecting from light.
5. Assay in a flow cytometer.
4. Notes
1. Cell and mitochondria isolation procedures are not presented here, because the
methods vary according to the organ used. For a wealth of information on stan-dard mitochondria techniques, we recommend the monograph Mitochondria, a
Practical Approach (10).
2. The fluorochrome NAO binds to all protein membranes regardless of their mem-brane potential (13) and, hence, the mitochondrial uptake of this metachromatic
dye does not depend on the mitochondrial energy status.
3. The forward angle light scatter represents the mitochondrial size, which contrib-utes to the refraction of light as it travels through the organelle. As an example of
age-related changes, we showed that the forward angle scatter for mitochondria
isolated from old rats (aged 22–36 mo) was about 140% of that of young rats
(aged 3–4 mo). Therefore, we concluded that the isolated organelles from old rats
were larger than those from young rats. In agreement with the mitochondrial
theory of aging (which assumes an impairment in mitochondrial division), a
senescent enlargement of the mitochondria of hepatocytes has been also shown,
using electron microscopy, by Miquel et al. (17) in hepatocytes, and by Bertoni-Freddari on nerve cells (see Chapter 17). As pointed out by Sastre et al. (6), the
increase in mitochondrial size with age probably affects mitochondrial function,
as volume-dependent regulation of matrix protein packing influences metabolite
diffusion and therefore mitochondrial metabolism.
4. The fluorescent dye DhRh123 is uncharged but it is oxidized to positively charged
Rh123 in the extramitochondrial space and then it is taken up by mitochondria,
where it accumulates. The MMP is the driving force for Rh123 uptake.
5. The MMP is monitored by fluorescence quenching of Rh123 in isolated mito-chondria under respiratory state 4. Under these conditions, the MMP is an index
of mitochondrial energy status. To induce state 4, mitochondria are incubated
with 5 mM sodium succinate.
6. As an example of gerontological application of the DHR123-flow cytometry
method to isolated cells, our work (6) on rat hepatocytes has shown that the MMP
of old rats (aged 22–36 mo) is 70% of that found in the hepatocytes of young rats
(aged 3–4 mo).
7. Hagen et al. (9) have also demonstrated an age-associated decline in the MMP of
hepatocyte mitochondria. Thus, evidence is building up in support of the view that
senescence is associated with an impairment of the mechanisms involved with ATP
Damage to Mitochondria 243
synthesis, that is, with the main function of mitochondria. In our opinion, this justifies
further application of the above techniques to elucidate the specific role played by
mitochondria in cellular aging under normal and experimentally altered conditions.
8. Another proof of the usefulness of flow cytometry study of mitochondrial aging
is provided by our finding of an increased DhRh oxidation in mitochondria iso-lated from the liver of old rats as compared to those isolated from young animals,
which suggests that senescence is accompanied by an increased oxygen stress.
1. Harman, D. (1956) Aging: a theory based on free radical and radiation  chemistry.
J. Gerontol. 11, 298–300.
2. Miquel, J. and Fleming, J. (1986) Theoretical and experimental support for an “oxy-gen radical-mitochondrial injury hypothesis of cell aging, in Free Radicals, Aging
and Degenerative Diseases (Johnson, J. E., Jr., Walford, R., Harman, D., and Miquel,
J., eds.) Alan R. Liss, New York, pp. 51–74.
3. Miquel, J. (l998) An update on the oxygen stress-mitochondrial mutation theory of
aging: genetic and evolutionary implications. Exp. Gerontol. 33, 113–126.
4. Yen, T. C., Chen, Y. S., King, K. L., Yeh, S. H., and Wei, Y. H. (1989) Liver mito-chondrial respiratory functions decline with age. Biochem. Biophys. Res. Commun.
165, 994–1003.
5. García de la Asunción, J., Millán, A., Pla, R., Bruseghini, L., Esteras, A., Pallardo,
F. V., Sastre, J., and Viña, J. (1996) Aging of the liver: age associated oxidative
damage to mitochondrial DNA. FASEB J. 10, 333–338.
6. Sastre, J., Pallardo, F. V., Pla, R., Pellin, A., Juan, G., O’Connor, E., Estrela, J. M.,
Miquel, J., and Viña, J. (1996) Aging of the liver: age associated mitochondrial
damage in intact hepatocytes. Hepatology 24, 1199–1205.
7. Sastre, J., Millan, A., García de la Asunción, J., Pla, R., Juan, G., Pallardó, F. V.,
O’Connor, E., Martin, J. A., Droix-Lefaix, M. T., and Viña, J. (1998) A  Ginkgo
biloba extract (Egb 761) prevents mitochondrial aging by protecting against oxida-tive stress. Free Radic. Biol. Med. 24, 298–304.
8. Hagen, T. M., Yowe, D. L., Bartholomew, J. C., Wehr, C. M., Do, K. L., Park, J.-Y.,
and Ames, B. N. (1997) Mitochondrial decay in hepatocytes from old rats: mem-brane potential declines, heterogeneity and oxidants increase. Proc. Natl. Acad.
Sci. USA 94, 3064–3069.
9. Chen, L. B. (1989) Mitochondrial membrane potential in living cells. Ann. Rev.
Cell Biol. 4, 155–181.
10. Darley-Usmar, V. M., Rickwood, D., and Wilson, M. T., eds. (1987) Mitochondria:
a Practical Approach, IRL, Oxford, pp. 1–16.
11. Ferrandiz, M. L., Martínez, M., De Juan, E., Díez, A., Bustos, G. and Miquel, J.
(1994) Impairment of mitochondrial oxidative phosphorylation in the brain of aged
mice. Brain Res. 644, 335–338.
12. Martínez, M., Ferrándiz, M. L., Díez, A., and Miquel, J. (1995) Depletion of cytosolic
GSH decreases the ATP levels and viability of synaptosomes from aged mice but
not from young mice. Mech. Ageing Dev. 84, 77–81.
244 Pallardo et al.
13. Maftah, A., Petit, J. M., Ratinaud, M. H., and Julien, R. (1989) 10-N nonyl-acridine
orange: a fluorescent probe which stains mitochondria independently of their ener-getic state. Biochem. Biophys. Res. 164, 185–190.
14. Petit, P. X., O’Connor, J. E., Grunawald, D., and Brown, S. C. (1990) Analysis of
the membrane potential of rat- and mouse-liver mitochondria by flow cytometry
and possible implications. Eur. J. Biochem. 194, 389–397.
15. Greenspan, P., Mayer, E. P., and Fowler, S. D. (1985) Nile red: a selective  fluores-cent stain for intracellular lipid droplets. J. Cell Biol. 100, 965–973.
16. Royal, J. A. and Ischiripoulos, H. (1993) Evaluation of 2′-7′-dichloro intracellular
H2O2 in cultured endothelial cells. Arch. Biochem. Biophys. 302, 348–355, 1993.
17. Miquel, J., Economos, A. C., and Bensch, K. G. (1981) Insect vs. mammalian aging,
in Aging and Cell Structure (Johnson, J. E. Jr., ed.), Plenum Press, New York, pp.
Mitochondrial DNA Mutations 245
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Analysis of Mitochondrial DNA Mutations
Robert W. Taylor, Theresa M. Wardell, Emma L. Blakely,
Gillian M. Borthwick, Elizabeth J. Brierley,
and Douglass M. Turnbull
1. Introduction
Although the precise mechanisms of the aging process remain poorly under-stood, a plausible theory for cellular dysfunction and deterioration during aging
involves mitochondria (1,2). The major function of mitochondria is to generate
energy for cellular processes in the form of ATP by oxidative phosphorylation.
Mitochondria contain their own DNA (mtDNA), a small 16.5 kb circular mol-ecule that encodes 13 essential polypeptides of the mitochondrial respiratory
chain, as well as 2 rRNAs and 22 tRNAs required for intramitochondrial pro-tein synthesis  (3). The mitochondrial respiratory chain is a series of five,
multisubunit protein complexes located within the inner mitochondrial mem-brane. The first four of these (complexes I–IV) reoxidize reduced cofactors
(NADH and FADH2) generated by the oxidation of foodstuffs, thereby gener-ating an electrochemical gradient across the inner mitochondrial membrane
which is harnessed by the fifth complex, the ATP synthetase, to drive the for-mation of ATP.
The mitochondrial aging hypothesis proposes that aging results from the
accumulation of detrimental mtDNA mutations during life, compromising the
cellular production of ATP to such a degree that it results in cellular dysfunc-tion and death. A number of features help to explain why mtDNA is particu-larly vulnerable to deleterious mutational events. The mitochondrial genome
has a mutation rate some 10-fold greater than that of nuclear DNA, lacks pro-tective histones, and possesses few and inefficient DNA repair mechanisms. It
246 Taylor et al.
is also highly vulnerable to nucleolytic attack by free radicals, a natural
byproduct of oxidative phosphorylation. Moreover, because mtDNA has no
introns and little redundancy, any mutational event within the genome is likely
to affect a coding sequence, and therefore biochemical function.
An increasing body of scientific evidence has arisen supporting a mitochon-drial involvement in the aging process, in particular the finding of pathogenic
mtDNA mutations in healthy, elderly subjects. Numerous mtDNA deletions
(4–15) have been shown to appear and accumulate with age in a variety of
human tissues. Point mutations have also been detected (16), although whether
they increase with age is debatable  (17). The accumulation of age-related
mtDNA deletions has also been shown to correlate with increased cellular lev-els of 8-hydroxy-2-deoxyguanosine, an indicator of free radical induced
mtDNA damage. Furthermore, cytochrome c oxidase (COX)-deficient muscle
fibers, a pathological hallmark of mitochondrial disease, have also been shown
to accumulate in an age-related manner (18,19). Although an apparent decline
in the activity of several respiratory chain enzymes has been reported with age,
these changes (as much as a 50% decline in activity in a tissue homogenate) are
hard to equate with the low levels of mtDNA mutations (<0.1% of the total
mtDNA) observed in tissues of elderly individuals. The mitochondrial genome
is present in multiple copies in individual mitochondria, and depending upon
the oxidative demand of the tissue in question, there may be up to several thou-sand genomes within individual cells. Because mtDNA mutations have been
shown to be incredibly recessive, with high levels (often >85% mutant mtDNA)
required before any biochemical dysfunction is apparent (20,21), a more subtle
molecular mechanism must exist to enable the low levels of mutations reported
in various tissues to impair biochemical function.
Recent studies in single muscle fibers from patients with autosomal domi-nant progressive external ophthalmoplegia (adPEO) (22) and healthy elderly
individuals (23) have shown that the clonal expansion of somatic mtDNA dele-tions in individual cells occurs to the extent that it results in a biochemical
(COX activity) defect. It has been proposed that a replicative advantage (either
at the level of the genome or for the mitochondria itself) allows mutant mtDNA
to accumulate within a cell over time to such a level that it surpasses the critical
threshold required to express a biochemical defect. Because somatic mtDNA
mutations may occur anywhere within the genome, one would expect to find
individual COX-deficient fibers harboring high levels of different deleted
mtDNA species, which has been shown to be the case (22,23).
In view of these findings, the molecular genetic techniques that are applied
to the study of somatic mtDNA mutations in aging must allow either the accu-rate quantification of these mutations at very low levels within tissue
homogenates, or be applicable to single-cell studies in which the level of mutant
Mitochondrial DNA Mutations 247
mtDNA is high compared to wild-type. Because of the number of polymerase
chain reaction (PCR)-based techniques available, we describe these protocols
in two chapters. In this, the first chapter, we discuss the methods used to inves-tigate mtDNA deletions, whereas the second chapter (Chapter 20) focuses on
the analysis of mtDNA nucleotide substitutions.
1.1. Long-Range PCR of mtDNA
Long-range PCR is now commonly used as an initial screen for the presence
of large-scale rearrangements (deletions or duplications) of mtDNA  (24). In
this respect, it has many advantages over Southern Blotting. Low levels of rear-ranged mtDNA species can be detected from small amounts of DNA template
within one day, whereas Southern blotting requires 2–3 µg of total DNA, takes
a number of days to complete, and has a detection threshold of about 5% mutant
mtDNA. For these reasons long-range PCR has been used to study the accumu-lation of age-related deletions in muscle and other tissues (25,26) as well as
characterizing pathogenic rearrangements in blood (27) and muscle (Fig. 1).
Although there are a number of commercially available kits for long-range
PCR, we and others favor the Expand™ Long Template PCR system produced
by Boehringer Mannheim. This uses a mixture of thermostable Ta q and Pwo
Fig. 1. 13 kb long-range PCR of mtDNA using primers L3200 and H16215.
M, λ-HindIII DNA markers;  C, control subject showing a single wild-type band;
lanes 1–5, subjects demonstrating different mtDNA rearrangements.
F. P.O.
248 Taylor et al.
(with a 3’→5′ exonuclease proofreading activity) DNA polymerases to enable
the accurate and efficient amplification of PCR products up to 40 kb in size.
1.2. Single-Cell PCR
The clonal expansion of mtDNA deletions within individual cells calls for
sensitive PCR-based techniques to enable the identification and characteriza-tion of these mtDNA mutations. Single-cell PCR on COX-deficient (biochemi-cally compromised) cells highlights the focal nature of these mutational events,
otherwise masked if analyzing DNA from tissue homogenates, and permits the
comparative study of mtDNA rearrangements in different cell types within the
same organ.
This section describes both the histochemical analyzes required to identify
those cells that are biochemically affected as demonstrated by COX activity,
and protocols for isolating total DNA from these single cells.
1.3. Quantitative analysis
of the 4977 Basepair Common Deletion in Single Cells
The PCR-based techniques described in the preceding section will detect
any rearrangement between nt 8355 and nt 13832, but will not quantify the
absolute levels of mutant mtDNA (deletion). When searching for high levels of
deleted mtDNA due to clonal expansion, competitive PCR (using three oligo-nucleotide primers to simultaneously amplify both wild-type and mutant
mtDNA) is a useful method for detecting and quantifying the common 4977-basepair deletion (mtDNA4977) in single cells (23). The method described below
is essentially that of Sciacco and colleagues (21).
1.4. Semiquantitative PCR of the 4977 Basepair Common
Deletion in Tissue Homogenates
Although the age-related accumulation of mtDNA deletions in various tis-sues has been demonstrated, the absolute levels of rearrangement in total cellu-lar DNA extracted from tissue homogenates are very low. Consequently,
semiquantitative PCR-based methods have been described to permit the inves-tigation of these accumulating mutations in different cell types. The protocol
we describe is based extensively on the method of Corral-Debrinski and col-leagues (7,8), in which the PCR amplification of wild-type mtDNA and
mtDNA4977 from serially diluted DNA samples allows the estimation of the
amount of deletion in a tissue.
1.5. Primer-Shift PCR
Primer-shift PCR was first described by Ozawa et al. (28) as a means to fine
map differentially deleted mtDNA species from total cellular DNA isolated from
tissue homogenates. The rationale is straightforward; using pairs of primers
Mitochondrial DNA Mutations 249
designed to amplify both (H) and (L) strands and relatively short extension times,
PCR products are obtained only if a deletion is present, thereby shifting the
PCR primers closer together, facilitating amplification. These PCR products will
contain the specific deletion breakpoint, and as such can either be cloned or
sequenced directly to map the precise location. Furthermore, because this tech-nique excludes nonspecific PCR amplification due to the misannealing of PCR
primers, it amplifies only deleted mtDNA species. Consequently, primer-shift
PCR has been used to characterize the multiple mtDNA associations observed
in patients with inclusion body myositis (29,30) and demonstrate the clonal expan-sion of mtDNA deletions in COX-deficient skeletal muscle fibers of patients
with adPEO (22). This study highlights the power of this technique to screen for
mtDNA deletions associated with disease or the aging process in DNA isolated
from a single cell. The protocol described in the following sections describes
the use of primer-shift PCR to investigate the presence of mtDNA deletions in
DNA isolated from single muscle fibers or individual neurons. The same method
is easily applied to the characterization of deleted mtDNA species in total cellu-lar DNA isolated from a tissue homogenate.
2. Materials
2.1. Long-range PCR of mtDNA
1. Expand™ Long Template PCR system (Boehringer Mannheim): This is supplied
with the enzyme mix and three buffers. Reaction buffer 3 is suitable for most
applications as it contains dimethyl sulfoxide (DMSO) (20% [v/v]) which pre-vents DNA depurination and intrastrand secondary structure formation. The
enzyme can be stored at –20°C for approx 3 mo. Reaction buffer 3 should be
checked for the appearance of crystals that may have precipitated before use.
2. Deoxynucleoside triphosphates (dNTPs): Separate 10 mM working solutions of
dATP, dCTP, dGTP, and dTTP are recommended. These are made from 100 mM
lithium salt dNTP stocks purchased from Boehringer Mannheim. Store at –20°C.
3. Bovine serum albumin (BSA): Although not essential for the reaction, the addi-tion of BSA (200 µg/mL final concentration) may increase the efficiency of the
long template amplification. A 10 mg/mL stock solution of BSA that is often
supplied with restriction endonucleases can be diluted to a 1 mg/mL working
solution for this purpose, and stored at –20°C.
4. Oligonucleotide primers: Any primers designed to amplify mtDNA may be used;
however, successful amplification may require testing various combinations of
primers and annealing temperatures. Increasing the annealing temperature and
reducing primer concentrations may help to reduce any non-specific PCR ampli-fication. We find that short primers (20–24 bases long) give good results, whereas
longer primers (>30 bases) do not. Interestingly, this inefficiency of amplifica-tion may be overcome by using a short primer in combination with a long primer.
For much of our routine screening, we amplify a 13-kb fragment of the mitochon-
250 Taylor et al.
drial genome using a forward primer L3200 (nt 3200–3219) and a reverse primer
H16215 (nt 16215–16196), numbered according to the Cambridge sequence (3). For
whole genome amplification, a number of articles have been published with primer
sequences; we have successfully used those described by Kovalenko and colleagues
(26). Stock solutions (10 µM) of primers are stored at –20°C (see Note 1).
5. DNA template: Only 10–50 ng of total DNA is required for amplification of the
genome in part or whole. The quality of the DNA template is crucial for success-ful amplification, and protein contamination (determined by measuring the A260/
A280 ratio should be no less than 1.8. We recommend that solutions of DNA
(10–50 ng/µL) be made fresh from concentrated DNA stocks, although these may
be stored for 3–4 d at 4°C and for 1–2 wk at –20°C (see Note 2).
6. Sterile water.
7. Ice: The reaction should always be set up on ice to avoid hot start.
8. Mineral oil if required to overlay the reaction.
9. Thin-walled PCR tubes: These permit a more efficient transfer of heat, and as
such are crucial for amplifying long templates. Use 0.2- or 0.5-mL thermotubes
(Applied Biosystems) depending upon thermal cycler used.
10. Thermal cycler: Both the Hybaid Omnigene and Perkin–Elmer GeneAmp® PCR
System 2400 thermal cyclers give good, reproducible results.
11. Horizontal gel electrophoresis equipment.
12. Agarose gel containing ethidium bromide; 1× TAE (40 mM Tris-acetate, 1 mM
EDTA, pH 8.0) running buffer.
13. UV transilluminator.
2.2. Single-Cell PCR
1. Tissue sections: Fresh muscle and neuronal tissue are frozen in liquid nitrogen
cooled isopentane and stored at –85°C. Fresh frozen sections (30  µm) are cut
using a Brights OTF cryostat and air-dried at room temperature for 30 min. These
sections can be used immediately or stored at –80°C in air-tight slide containers.
2. Histochemical staining: The assay of COX activity requires stock solutions
of 5 mM 3,3′-diaminobenzidine tetrahydrochloride (light sensitive) (DAKO) in
0.1 M phosphate, pH 7.0, and 500 µM cytochrome c (light sensitive) (Sigma) in
0.1 M phosphate, pH 7.0. The assay of succinate dehydrogenase (SDH) activity
requires stock solutions of 1.875 mM nitroblue tetrazolium (Sigma), 1.30  M
sodium succinate (Sigma), 2 mM phenazine methosulfate (light sensitive)
(Sigma), and 100 mM sodium azide (BDH) all in 0.1  M phosphate, pH 7.0.
Toluidine blue staining is performed with a 1% solution of toluidine blue (light
sensitive) (Sigma) in 1% sodium borate (Sigma).
3. Capillaries: Standard wall borosilicate glass capillaries without filament, 1.0 mm
outer diameter  × 0.58 mm inner diameter (GC100–15 Clark Electromedical
Instruments) are used for muscle dissection. Standard wall borosilicate glass cap-illaries without filament, 1.5 mm outer diameter  × 0.86 mm inner diameter
(GC150–10 Clark Electromedical Instruments) are used for neuronal dissection.
Micropipets are produced using a Narishige PC-10 micropipet puller.
Mitochondrial DNA Mutations 251
4. Micromanipulation: An inverted microscope (Zeiss Axiovert 25) fitted with 10×
and 20× objectives is used for muscle dissection. A Leica inverted microscope
(DMIRB/E) fitted with 4×, 10×, 20×, and 40× long-distance working objectives
and a Leica mechanical micromanipulator fitted with a Narishige injection holder
(HI–6) is used for neuronal dissection. A stock solution of 10 mM Tris-HCl,
1 mM EDTA, pH 7.4 (TE buffer) is required.
5. Cell lysis: Stock solutions of 1% Tween-20 (Sigma); 100 mM EDTA, pH 8.0;
0.5 M Tris-HCl, pH 8.5; and 50 mg/mL of proteinase K (NBL Gene Sciences).
6. PCR amplification: The following stock solutions are required: 2 mM (10×)
dNTPs (Boehringer Mannheim), 10× GeneAmp® PCR buffer containing 100 mM
Tris-HCl, pH 8.3; 500 mM KCl and 15 mM MgCl2 (Perkin–Elmer); 20 µM stock
solution of each forward and reverse oligonucleotide primer, and 5 U/µL of
AmpliTaq® DNA polymerase (Perkin–Elmer). Each PCR amplification is
performed in a 0.5-mL thin-walled thermo-tube (Applied Biosystems).
7. Ice: All PCR reactions are set up on ice.
8. Thermal cycler.
9. Horizontal gel electrophoresis equipment.
10. UV transilluminator.
2.3. Quantitative Analysis of the 4977 Basepair Common
Deletion in Single Cells
1. Sterile water.
2. Sterile 500-µL Eppendorf tubes for PCR.
3. Ice: All PCR reactions are set up on ice.
4. Lysis solution: 200 mM KOH, 50 mM dithiothreitol (DTT). Combine 200 µL of
1 M KOH (make fresh weekly and store at 4°C), 100 µL of 500 mM DTT (store
aliquots at –20°C), and 700 µL of water.
5. Neutralization solution (900 mM Tris-HCl, pH 8.3; 200 mM HCl). Combine
900 µL of 1 M Tris-HCl (store at 4°C), 40 µL of 5 M HCl (room temp), and 60 µL
of sterile water just before use.
6. Heat block.
7. Oligonucleotide primers: A set of three primers are used to amplify both wild-type and deleted mtDNA simultaneously. These are L8273 (nt 8273–8289),
H9028 (nt 9028–9008), and H13720 (nt 13720–13705). The primers L8273 and
H9028 amplify 755 basepairs corresponding to wild type mtDNA and the prim-ers L8273 and H13720 amplify a 470-basepair fragment corresponding to
mtDNA4977. Stock solutions (20 µM) are stored at –20°C.
8. PCR amplification: 2 mM (10×) dNTPs are from Boehringer Mannheim. Ther-mostable DNA polymerase and the associated 10× reaction buffer (containing
MgCl2) are from Advanced Biotechnologies.
9. [α-32P] dATP (3000 Ci/mmol) (Amersham Life Science Products).
10. Mineral oil.
11. PCR thermal cycler.
12. Vertical polyacrylamide gel electrophoresis system (SE 600, Hoefer)
252 Taylor et al.
13. 5% nondenaturing polyacrylamide gel.
14. Whatman 3MM filter paper.
15. Saran Wrap®.
16. Running buffer (1× TBE): 90 mM Tris-borate, 2 mM EDTA, pH 8.0.
17. Gel dryer (Model 543, Bio-Rad Laboratories).
18. PhosphorImager and ImageQuant software (Molecular Dynamics).
2.4. Semiquantitative PCR of the 4977-Basepair Common
Deletion in Tissue Homogenates
1. Template DNA.
2. BamHI restriction endonuclease (10 U/µL) and SuRE/Cut Buffer B for restric-tion (10×) (Boehringer Mannheim).
3. PCR amplification: The following stock solutions are required: 2 mM (10×)
dNTPs (Boehringer Mannheim), 10× GeneAmp® PCR buffer containing 100 mM
Tris-HCl, pH 8.3; 500 mM KCl and 15 mM MgCl2 (Perkin–Elmer), AmpliTaq®
DNA polymerase (Perkin–Elmer). Each PCR amplification is performed in a
0.5-mL thin-walled thermo-tube (Applied Biosystems).
4. Oligonucleotide primers: Two pairs of primers are used to amplify a rarely deleted
region of mtDNA (wtDNA) and mtDNA containing the common deletion
(mtDNA4977). The primers L3108 (nt 3108–3127) and H3717 (nt 3717–3701)
amplify a 610-basepair fragment corresponding to wtDNA, whereas L8282
(nt 8282–8305) and H13851 (nt 13851–13832) amplify a 593-basepair fragment
corresponding to mtDNA4977. Stock solutions (20 µM) are stored at –20°C.
5. Thin-walled PCR tubes: 0.5-mL thermotubes (Applied Biosystems) are recommended.
6. Ice: All PCR reactions are set up on ice.
7. PCR thermal cycler.
8. Large horizontal gel electrophoresis unit (Maxi unit, Scotlab) with well-forming
combs to produce wells of 50 µL volume.
9. 1.5% Agarose gel containing ethidium bromide, 1× TAE running buffer.
10. UV transilluminator.
11. Digital imaging system and imaging software for PCR quantitation (AlphaImager
and AlphaEase, Flowgen).
2.5. Primer-Shift PCR
1. Template DNA: The isolation of total DNA from single muscle fibers and indi-vidual neurons is described in detail in Subheading 3.2.
2. PCR reagents: Primer-shift PCR is performed on a Perkin–Elmer GeneAmp®
PCR System 2400 thermal cycler using AmpliTaq® DNA polymerase (Perkin–Elmer)
and the 10× reaction buffer supplied. dNTPs are made as a 2 mM (10×) stock.
Thin-walled PCR tubes (0.2 mL) are also from Perkin–Elmer.
3. Oligonucleotide primers: Although any combinations of mtDNA-specific PCR
primers can be used to map mtDNA deletions, we have essentially used those
described by Moslemi et al.  (22), in the combinations shown in  Fig. 2. The
sequences of these primers are as follows: L1 (nt 1–20), L7901 (nt 7901–7920),
Mitochondrial DNA Mutations 253
L8197 (nt 8197–8216), H466 (nt 466–447), H9050 (nt 9050–9031), H13640 (nt
13640–13621), H14840 (nt 14840–14821), H15260 (nt 15260–15241), H16150
(nt 16150–16131), and H16560 (nt 16560–16541). Stock solutions (20 µM) are
stored at –20°C.
Fig. 2. Primer-shift PCR from single muscle fibers showing the combination of
PCR primers (pairs a–h) used to screen the mtDNA genome. A shows the products
generated from total DNA isolated from skeletal muscle of a patient with multiple
mtDNA deletions. B and C represent the products generated from the DNA isolated
from two individual COX-deficient muscle fibers, highlighting the presence of differ-ent deleted mtDNA species in each fiber.
254 Taylor et al.
4. Sterile water.
5. Ice: All reactions are set up on ice.
6. Horizontal gel electrophoresis equipment.
7. 1.5% Agarose gel containing ethidium bromide; 1× TAE running buffer.
3. Methods
3.1. Long-Range PCR of mtDNA
1. Thaw all components stored at –20°C and bring reaction buffer 3 to room tem-perature. Vortex well to mix, checking the reaction buffer for crystals. If crystals
are present leave at room temperature overnight and mix well again. Place all the
components on ice.
2. Label two, sterile 1.5-mL Eppendorf tubes A and B; the master mix is made up
as two separate mixes to avoid degradation of the primers or template due to the
3’→5′ exonuclease activity of the Pwo enzyme.
3. Combine the following components to form mix A for each 50 µL PCR:
2.5 µL 10 mM dATP
2.5 µL 10 mM dCTP
2.5 µL 10 mM dGTP
2.5 µL 10 mM dTTP
1.5 µL10 µM Forward primer
1.5 µL10 µM Reverse primer
12 µL Sterile water
4. Vortex-mix briefly, centrifuge the mix, and aliquot 25 µL into each thin-walled
tube on ice.
5. Add 1 µL of DNA template to each separate tube.
6. Prepare master mix B; for each 50-µL reaction combine the following:
5 µL10× Reaction buffer 3
10 µL1 mg/mL of BSA
8.65 µL Sterile water
*0.35 µL Expand™ enzyme
*While the kit recommends using 0.7 µL of enzyme per 50 µL of PCR reaction,
we find that the same efficiency of amplification is achieved using only 0.35 µL.
Vortex-mix and briefly centrifuge the mix, aliquoting 24 µL into each thin-walled tube, mixing gently to avoid introducing air bubbles.
7. If necessary overlay the reactions with 30 µL of mineral oil.
8. Centrifuge the samples briefly, and place back on ice.
9. Begin the following PCR program to heat the block to 92–94°C:
Initial denaturation 92–94°C* 3 min
10 Cycles 92–94°C* 10–30 s
55–68°C** 30 s
Mitochondrial DNA Mutations 255
68°C† x min
20 cycles 92–94°C* 10–30 s
55–68°C** 30 s
68°C† x min + 5 s per cycle
Final extension 68°C 15–20 min
*Use lower denaturing temperatures and shorter times for longer products to
avoid damaging the template.
**Annealing temperature is dependent upon the oligonucleotide primers used
in the PCR reaction. For the amplification of a 13-kb product using L3200 and
H16215, we use an annealing temperature of 58°C.
†x depends on the length of product to be amplified; generally allow 1 min for
every 1 kb to be amplified.
10. Place tubes in thermal cycler when there is 90 s remaining of the initial denatur-ation step.
11. When the program is complete, electrophorese 15–20 µL of the PCR products
through a 0.7% agarose gel at 65 V for 2–4 h (see Notes 3–5). The remaining
product may be used for further analysis such as restriction enzyme digest or
primer-shift PCR to map the rearrangement (31).
3.2. Single Cell PCR
1. Histochemical staining: Tissue sections that have been stored at –85°C should be
equilibrated at room temperature for 30 min, then removed from the air-tight
container and air-dried for a further 30 min prior to histochemical analysis. COX
activity is detected using incubation medium containing 4 mM 3,3′-diaminobenzidine tetrahydochloride and 100 µM cytochrome c. Each section is
incubated with 100–200 µL of incubation medium at 37°C for up to 60 min in a
humid chamber. Any excess medium is rinsed using distilled water. The activity
of SDH in the sections is detected using incubation medium containing 1.5 mM
nitroblue tetrazolium, 130 mM sodium succinate, 0.2 mM phenazine methosulfate
and 1 mM sodium azide. Each section is incubated at 37°C with 100–200 µL of
incubation medium, in a humid chamber for 30 min. Sections are rinsed in dis-tilled water to remove excess medium. COX-deficient cells are detected by a
double activity assay; sections are initially assayed for COX activity, as mea-sured by the production of a brown end product. After removal of the excess
medium, the sections are subsequently assayed for SDH activity. COX-deficient
cells are easily identified as they do not produce the brown reaction product asso-ciated with COX activity, but do react for SDH activity, which gives a character-istic blue appearance (32). The neuronal cell harvesting is performed immediately
after histochemistry; however, muscle sections can be stored in 50% alcohol at
4°C for several months.
2. Micropipets: Micropipets of a specific diameter are produced from heat-steril-ized glass capillaries, using a micropipet puller in the double pull mode. A range
of tip diameters (1–50 µm) can be made, depending on the size of the cells to be
256 Taylor et al.
harvested. The diameter of the micropipet is determined using a light microscope
fitted with a graticule. Crude micropipets for muscle fiber work can be prepared
by heating and pulling glass capillaries in a Bunsen burner flame. These micropi-pets can then be heat sterilized. Prior to using these crude micropipets the tip is
gently broken against the microscope slide to yield a tip of appropriate diameter
(see Note 6).
3. Dissection of single muscle fibers: Single muscle fibers can be picked from appro-priately stained sections (30 µm) by hand, using a standard inverted microscope.
The muscle section is kept under 50% alcohol during this procedure. The micro-pipet is held by hand, and the tip is used to tease around the edge of an individual
muscle fiber and pick up the fiber. The presence of a cell on the capillary is deter-mined visually with a hand held eyepiece.
4. Dissection of neuronal cells: An inverted microscope with long-distance working
objective lenses is used in conjunction with a micromanipulator. The micropipets
are held in position with an injection holder attached to an instrument holder on
the micromanipulator. Stained sections (30 µm) of the neuronal tissue under inves-tigation are moistened prior to micromanipulation with TE buffer. Single neurons
are then picked up on the end of a micropipet. Visual confirmation of the pres-ence of a cell is obtained by focusing on the micropipet while it is still held in the
injection holder (Fig. 3) (see Note 7)
5. Cell lysis: The tip of the micropipet containing the single cell is broken off into
sterile microfuge tubes, containing 20 µL of TE buffer. At this point the cells can
be stored at 4°C for several weeks, prior to lysis. The microfuge tubes containing
the individual cells are centrifuged at 7000g for 10 min. The supernatant is
removed and the cells are lysed with 10 µL of 50 mM Tris-HCl, pH 8.5; 1 mM
EDTA, pH 8.0; 0.5% Tween-20, and 200 µg/mL of proteinase K (33). The cells
are incubated for 2 h at 55°C, with agitation every 30 min. This is followed by
heat inactivation of the proteinase K at 95°C for 10 min (see Note 8).
6. PCR amplification of single cells: PCR amplification of a rarely deleted region of
mtDNA (ND1 and 16S rRNA genes) is performed to demonstrate the presence of
template DNA (wtDNA) within the single cell lysate. The primers are L3108 (nt
3108–3127) and H3717 (nt 3717–3701) which amplify a 610-basepair product.
The common deletion (mtDNA4977) is amplified using the primers L8282 (nt
8282–8305) and H13851 (nt 13851–13832) which yield a 593-basepair product.
The cell lysate is used as the template in a single PCR reaction. A standard PCR
procedure is performed using AmpliTaq® (1 U/reaction), PCR buffer, 0.6 µM for-ward and reverse primer, and 200 µM dNTPs in a 50-µL reaction volume. PCR
amplification is performed under the following conditions: initial denaturation at
94°C for 2 min, denaturation at 94°C for 45 s, annealing at 51°C (wtDNA) or
56°C (mtDNA4977) for 45 s, and extension at 72°C for 1 min for 34 cycles fol-lowed by a final extension at 72°C for 8 min  (7). PCR products (40  µL) are
electrophoresed through a 1.5% agarose gel in 1× TAE buffer containing ethidium
bromide, and visualized by UV transillumination.
Mitochondrial DNA Mutations 257
7. Seminested PCR: The PCR products from the PCR reactions described in the
previous section can be amplified by performing a semi-nested PCR using 1 µL
of the primary PCR product as template and one internalized primer. The PCR is
performed as described in step 6 using the primers L3275 (nt 3275–3306) and
H3717, and an annealing temperature of 56°C for wtDNA. The primers L8333
(nt 8333–8355) and H13851 to13832 are used to amplify deleted mtDNA in the
mtDNA4977 PCR, annealing at a temperature of 56°C.
3.3. Quantitative Analysis of the 4977 bp Common Deletion
in Single Cells
1. Isolate cells as described in the previous section and then place in a sterile 500 µL
Eppendorf containing 20 µL water. Centrifuge tube for 10 min, remove the water
carefully then add 5 µL of the lysis solution. Place tubes into the heat block at
Fig. 3. Microdissection of an individual Purkinje cell for PCR analysis. The neu-ronal tissue section is stained with 1% toluidine blue, a single cell targeted (A, arrow)
and removed by micromanipulation (B) onto the end of a micropipet (C). The asterisk
(*) highlights the position from where the single cell was removed.
258 Taylor et al.
65°C for 1 h, add 5  µL of the neutralising solution to each tube and remove
from heat.
2. Prepare a PCR mastermix by combining, for 10 tubes, the following:
582.5 µL Sterile water
100 µL10× Reaction buffer
100 µL2 mM (10×) dNTPs
50 µL20 µM L8273
50 µL20 µM H13720
12.5 µL20 µM H9028
5 µL Thermostable DNA polymerase
5 µL[α-32P] dATP (3000 Ci/mmol)
3. Add 90 µL of the PCR master mix to each tube containing 10 µL of single cell
lysis and overlay with mineral oil.
4. Place tubes into thermal cycler and begin the following PCR program:
Initial denaturation 94°C1 min
25 Cycles 94°C1 min
55°C1 min
72°C2 min
Final extension 72°C 15 min
5. Electrophorese samples through a 5% nondenaturing polyacrylamide gel in 1×
6. Place polyacrylamide gel on filter paper, cover with cling film, and dry on gel
dryer. When dry, lay gel on PhosphorImager cassette overnight, and analyze
and quantitate the radioactive fragments using a PhosphorImager and
ImageQuant software (see Notes 9–11).
3.4. Semiquantitative PCR of the 4977-Basepair Common
Deletion in Tissue Homogenates
1. Linearization of DNA: A 125 ng/µL solution of the DNA sample under investi-gation is digested with the restriction endonuclease BamHI at 37°C for 90 min
(see Notes 12 and 13). Set up the following reaction:
8 µL DNA
1 µL BamHI (Boehringer Mannheim)
1 µL SuRE/Cut Buffer B (10×) (Boehringer Mannheim)
2. Serial dilution of DNA: A serial dilution (1:2) of DNA is prepared for each
PCR (wild-type and common deletion). The range for the wild-type PCR is
10–3.81 × 10–5 ng/µL, comprising a total of 19 concentrations. The common
deletion PCR has a range of DNA concentrations from 100 to 0.003 ng/µL,
comprising a total of 16 dilutions (see Note 14).
Mitochondrial DNA Mutations 259
3. PCR amplification: For each concentration of serially diluted DNA (1 µL), set up
the following reaction:
35.8 µL Sterile water
5 µL10× GeneAmp® reaction buffer
5 µL2 mM dNTPs
1.5 µL L3108 or L8282
1.5 µL H3717 or H13832
1 µLTemplate DNA
0.2 µL AmpliTaq® DNA polymerase (see Note 15)
When setting up a number of PCR reactions to investigate a number of con-centrations, make an appropriate volume of a PCR master mix for each primer
pair containing all the components except template DNA. Aliquot these into the
respective tube, and add 1 µL of the appropriate serially diluted DNA.
4. Perform 34 cycles of amplification as follows:
Initial denaturation 94°C2 min
34 Cycles 94°C 45 s
51°C for 30 s (wtDNA) or
56°C for 30 s (mtDNA4977)
72°C1 min
Final extension 72°C8 min
5. Electrophorese the PCR products (40 µL) through a 1.5% agarose gel in 1× TAE
buffer using a large horizontal electrophoresis unit.
6. Visualize bands by UV transillumination.
7. Quantitation of PCR: The gel image is stored using a digital imaging system, and
the optical densities of the PCR products are quantified using image analysis
software. An integrated density value (IDV) for a set area is obtained for each
PCR product. The DNA concentration at which the IDV is zero is obtained for
the wtDNA and mtDNA4977, respectively. The percentage of mtDNA4977 deletion
in the DNA sample is calculated by dividing the DNA concentration at which the
wtDNA IDV is zero by the DNA concentration at which the mtDNA4977 IDV is
zero (see Note 16).
3.5. Primer-Shift PCR
1. PCR amplification: Prepare a master mix containing all the components of the
PCR reactions with the exception of oligonucleotide primers. For eight reactions
(final volume of 50 µL) you will need the following:
282.8 µL Sterile water
40 µL10× Reaction buffer
40 µL2 mM dNTPs
10 µLTemplate DNA (single cell lysis)
3.2 µL AmpliTaq® DNA polymerase
260 Taylor et al.
2. Vortex-mix briefly, centrifuge, and aliquot 47  µL into each of eight tubes
labeled a–h.
3. Add 1.5 µL of each of the appropriate forward and reverse primer.
4. Centrifuge briefly and place in PCR thermal cycler
5. Perform 35 cycles of amplification as follows:
Initial denaturation 94°C2 min
35 cycles 94°C 30 s
53°C 30 s
72°C 30 s
Final extension 72°C8 min
6. Electrophorese samples (30–40 µL) through a 1.5% agarose gel for 1 h, and visu-alize by UV transillumination (see Notes 17–19).
4. Notes
1. Ensure working solutions of oligonucleotide primers are prepared regularly, as
they are prone to degrade more rapidly at lower concentrations and after repeated
cycles of freeze–thawing.
2. Identify a DNA sample that can be used as a control for each reaction; this will
allow identification of nonspecific products.
3. Nonspecific products can be avoided by raising annealing temperatures or
decreasing primer concentrations.
4. If no product is amplified from a particular DNA sample, raise the concentration
of DNA added to the reaction up to 100 ng. If there is still no visible product
check protein contamination and treat the sample with phenol/chloroform  if nec-essary (this may also be required for DNA samples extracted more than a year
prior to amplification).
5. If smearing occurs reduce the number of cycles or the amount of DNA added.
6. The tips of the micropipets are very fragile and can be damaged during autoclav-ing; therefore it is important to sterilize the capillaries prior to pulling. The pro-cedure then needs to be performed maintaining the sterility of the micropipets.
During the production of micropipets it is essential to check while measuring the
diameter of the tips that a smooth edge has been produced.
7. Initially using this single cell method it is recommended that a number of cells
are pooled and lysed as the template to establish the technique.
8. There are several alternatives to this lysis method. Individual cells can be lysed
by adding 10 µL of 200 mM KOH, 50 mM DTT, and incubated at 65°C for 1 h
(21,23). Samples are neutralized by the addition of 10 µL 900 mM Tris-HCl, pH
8.3, and 200 mM HCl, and the DNA phenol/chloroform extracted and precipi-tated at –85°C in 2 vol of 100% ethanol, 1/10 vol of 3 M sodium acetate, pH 5.2,
and 5 µL of 0.3 mg/mL glycogen (Boehringer Mannheim). The resulting pellet is
centrifuged, air-dried, and resuspended in a volume of sterile water. Alternatively,
cell lysis can be performed in the PCR mix by the addition of 5 µL of 10% Triton
Mitochondrial DNA Mutations 261
X-100. This mix is then heated at 94°C for 5 min prior to adding 0.2  µL of
AmpliTaq® DNA polymerase and commencing the PCR amplification as
described in Subheading 3.2., step 6.
9. It is essential that preliminary control studies are done using samples containing
a varying proportion of wild-type mtDNA and mtDNA4977, proportions deter-mined by Southern blotting. This ensures that the PCR reflects the two popula-tions of mtDNA in the original sample.
10. The concentration of the template DNA affects the ratios of wild-type to
mtDNA4977 especially at very low concentrations (21).
11. A control sample containing wild-type mtDNA and mtDNA4977 of known propor-tion must be run and quantitated on each PCR run.
12. The template DNA must be of good quality, with an A260/A280 ratio of no less
than 1.8.
13. Other restriction enzymes that linearize the mitochondrial genome (e.g., PvuII)
can be used as an alternative to BamHI. Linearized DNA samples can be stored
successfully at –20°C for 2–3 d. Over longer periods of time, degradation of the
DNA sample will lead to the appearance of additional, nonspecific bands follow-ing PCR amplification.
14. Serial dilution of DNA: It is essential that great care is taken at this stage to
obtain an accurate dilution. This requires accurate pipetting and thorough mixing
of each sample.
15. DNA polymerase: We strongly recommend that AmpliTaq® DNA polymerase
(Perkin–Elmer) is used. Cheaper alternatives do not give reliable amplification in
our hands.
16. Quantitation of PCR: When analyzing the gels of the PCR products, it is impor-tant to link the IDV values to a local background setting.
17. Primer pairs a and b are designed to amplify normal mtDNA from regions of the
genome that are rarely (primer pair a) and often (primer pair b) deleted (Fig. 3).
They serve as good controls for effective cell lysis.
18. PCR products generated by amplification with primer pairs c–h are specific to
the deletion found in that particular cell. Figure 2 shows the amplification of two
different deletions from individual COX-deficient muscle fibers, and highlights
the presence of a single mutation at high levels in these fibers. These PCR prod-ucts encompass the deletion breakpoint, the sequence of which can be determined
by directly sequencing the primer-shift PCR product.
19. If no visible products are amplified using any of the primer pairs, a secondary
PCR can be performed exactly as described using 1 µL of the initial PCR reaction
as template.
The financial support of the Northern Regional Health Authority, Research
into Ageing, the Muscular Dystrophy Group of Great Britain, and the Wellcome
Trust is acknowledged.
262 Taylor et al.
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Mitochondrial DNA Mutations 265
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Analysis of Mitochondrial DNA Mutations
Point Mutations
Robert W. Taylor, Richard M. Andrews, Patrick F. Chinnery, and
Douglass M. Turnbull
1. Introduction
Since the first demonstration that mutations of the mitochondrial genome
were associated with human disease, more than 100 pathological mitochon-drial DNA (mtDNA) defects have been characterized in patients with a broad
spectrum of clinical manifestations (1). Single-point mutations, involving either
protein-encoding genes or more commonly RNA (rRNA and tRNA) genes,
represent a substantial proportion (more than one third) of the pathogenic
mtDNA mutations described in the literature, and this number is steadily
increasing (2,3). Although some of the more common mtDNA point mutations
can be screened using simple polymerase chain reaction (PCR)-based tech-niques (e.g., restriction digest analysis), an increasing number of pathological
point mutations are identified only when large-scale sequencing of either all 22
tRNA genes or the whole mitochondrial genome is performed (4–7).
Any point mutation that is identified by sequence analysis must fulfil a num-ber of accepted criteria before pathogenicity is ascribed, particularly as the
mitochondrial genome is highly polymorphic, with any two individuals differ-ing by as much as 40–50 basepairs from the reference Cambridge sequence
(8). The majority of pathogenic mtDNA mutations are heteroplasmic, that is,
both mutant and wild type mtDNA are present within the same cell and tissue.
Because the proportion of mutant to wild-type mtDNA can vary between dif-ferent tissues, this can cause problems when sequencing if the level of mutant
mtDNA is particularly low, and as such these studies are performed using DNA
extracted from postmitotic tissues such as skeletal muscle in which the level of
266 Taylor et al.
mutant mtDNA is often high. However, the demonstration of heteroplasmy
is only one of a number of accepted criteria for a pathogenic role, particularly
as mtDNA heteroplasmy in both coding and noncoding regions of mtDNA is
increasingly being found in nondisease subjects (9). The mutation can be shown
to be disease specific by demonstrating high levels of mutated mtDNA segre-gating with a biochemical defect (e.g., cytochrome  c oxidase [COX] defi-ciency) in single cells (5–7).
This chapter describes the strategies developed in our laboratory to amplify
and sequence the entire mitochondrial genome, and the methods with which
we confirm and quantify the level of mtDNA heteroplasmy in a DNA sample.
1.1. Automated Sequencing of mtDNA
The automation of DNA sequencing makes it possible to undertake large-scale sequencing projects to detect mtDNA mutations involved in disease and
the aging process. Although a number of protocols exist in the literature that
differ in both the method of template preparation and sequencing chemistries
used (10), any strategy adopted must be capable of detecting mtDNA hetero-plasmy. We have therefore chosen to amplify segments of the genome by PCR
using forward and reverse mtDNA primers that have 18 bases of universal M13
primer sequence at their 5′ ends (Tables 1 and 2). Dye primer cycle sequencing
with universal M13 primers using ABI PRISM Ready Reaction kits is subse-quently performed, and samples separated electrophoretically on an ABI 373
DNA sequencer (Appled Biosystems). This approach is efficient in both labor
and time while providing high-quality sequence data for the whole mitochon-drial genome including the noncoding region from 28 primary PCR amplifications.
Two methods of cycle sequencing are currently available. Dye terminator
sequencing involves using unlabeled sequencing primers with fluorescently
labeled dideoxynucleotides. Although this is the most rapid and convenient
approach, individual peak heights are typically variable which can make the
interpretation of base calling difficult. Dye primer cycle sequencing utilizes
fluorescently tagged primers, while the dideoxynucleotides are unlabeled.
Although this method requires four separate extension reactions that are subse-quently pooled, dye primer sequencing has the advantage that it provides a
cleaner signal with more even peak heights which facilitates the identification
of heteroplasmic bases changes (see Note 1).
To generate sufficient mtDNA template for sequencing, we have chosen to
use PCR amplification rather than bacterial cloning on account of both ease of
use and speed, while still giving comparable results. The purity of the prepared
template is extremely important in determining the quality of the final sequence
data, and each PCR amplification will require optimization (i.e., a single prod-uct of the correct size in appropriate amounts). We have found that the main
Mitochondrial DNA Mutations 267
problem is the formation of PCR artefacts, particularly “primer–dimers.” If
carried through to the sequencing reaction, dye primer labeling of these
artefacts will result in an increase in the background noise and the formation of
“stop peaks” at the start of the sequence, rendering some of the data unread-able. Such artefacts are reduced by performing a “hot start” in the PCR. We
therefore use AmpliTaq Gold™ (Perkin–Elmer), which requires a pre-PCR heat
activation at 94°C for 12 min to provide a “hot start.”
Although it is possible to cycle sequence PCR products directly, we have
found both quantification and purification of the amplified products necessary
Table 1
L-strand PCR Primers Used for Amplification of mtDNA
for Dye Primer Sequencing
Primer Position Sequence (5′-3′)
268 Taylor et al.
to give consistently high quality sequence data. QIAquick PCR Purification
columns (Qiagen) are both quick and easy to use and have the advantage that
the final elution volume can be varied to give a template concentration appro-priate for the length of fragment to be sequenced.
1.2. Last Hot Cycle PCR
The demonstration of mtDNA heteroplasmy associated with a biochemical
defect remains the most compelling piece of evidence of pathogenicity for any
mutation as it implies a recent mutational event. In view of the high levels of
Table 2
H-strand PCR Primers Used for Amplification of mtDNA
for Dye Primer Sequencing
Primer Position Sequence (5′-3′)
Mitochondrial DNA Mutations 269
mutant mtDNA required for the expression of a biochemical defect (often >85%
for point mutations), a number of different methods have been developed
to accurately measure the level of mtDNA heteroplasmy, thus providing more
clues about the molecular basis of their pathogenesis. These include PCR-single-strand conformation polymorphism (PCR-SSCP) analysis, multiple
clonal analysis, allele-specific oligonucleotide hybridization, and more recently
a fluorescence-based primer extension method  (11–14). However, for many
laboratories, the method of choice remains PCR-restriction fragment length
polymorphism (RFLP) analysis, especially as this is readily applicable to the
study of mtDNA heteroplasmy in single cells. Essentially, the region of mtDNA
containing the point mutation under investigation is amplified by PCR,
radiolabeled, and digested by a restriction endonuclease that can discriminate
between wild-type and mutant mtDNA molecules at the site of the point
mutation. Digestion products are electrophoresed through either agarose or
nondenaturing polyacrylamide gels and the level of heteroplasmy calculated
by determining the incorporation of radiolabel into each of the restriction prod-ucts. The formation of heteroduplex molecules during the PCR reaction can
make quantitation difficult because these are not digested by restriction endo-nucleases. However, the addition of radiolabel to the last cycle of amplification
(hence “last hot cycle PCR”) avoids the detection of these heteroduplexes,
allowing an accurate determination of heteroplasmy (15,16).
We describe in the following section the methodology associated with deter-mining the level of mtDNA heteroplasmy in both tissue homogenates and single
cells. As an example, we describe the strategy used to investigate a patient
previously described as heteroplasmic for a T10010C transition in the tRNAGly
gene (17).
2. Materials
2.1. Automated Sequencing of mtDNA
1. 2 mM (10×) dNTP mix for PCR. Store at –20°C
2. 10× GeneAmp PCR buffer: 100 mM Tris-HCl, pH 8.3, 500 mM KCl, 15 mM
MgCl2, 0.01% (w/v) gelatin (Perkin–Elmer). Store at –20°C.
3. AmpliTaq Gold™ DNA polymerase (5 U/µL) (Perkin–Elmer). Store at –20°C.
4. Template DNA: Dilute to 200 ng/µL for PCR amplification.
5. Oligonucleotide primers for PCR: Owing to the length of read that is now pos-sible using automated DNA sequencers, the mitochondrial genome is amplified
in fragments of between 650 and 750 basepairs using a set of forward (L) and
reverse (H) primer pairs (e.g., 01F and 01R) that have been designed to anneal at
58°C. Both sets of primers have 18 bases of M13 sequence at their 5′ end, allow-ing the products to be cycle sequenced. The M13 sequences used are: forward (L)
primers are prefixed by 5′ TGTAAAACGACGGCCAGT 3′; reverse (H) primers
270 Taylor et al.
are prefixed by 5′ CAGGAAACAGCTATGACC 3′ (see Tables 1 and 2). Owing
to the length of these PCR primers (35–40 basepairs), we recommend that they
are hplc purified following synthesis. Stocks for PCR amplification (20 µM) are
stored at –20°C (see Note 2).
6. DNA thermal cycler: This is required for the initial PCR amplifications and the
subsequent cycle sequencing. To achieve a high throughput of samples, a thermal
cycler capable of running 96 samples is desirable. We use the GeneAmp® PCR
System 9700 (Perkin–Elmer) which has a heated lid facility, thereby negating the
need for mineral oil overlay for either the PCR amplification or cycle sequencing
7. Sterile water.
8. 0.2 mL—MicroAmp® Reaction tubes (Perkin–Elmer).
9. Horizontal gel electrophoresis equipment for running 1% agarose gels; 1× TAE
(40 mM Tris-acetate, 1 mM EDTA, pH 8.0) running buffer containing ethidium
10. UV transilluminator.
11. QIAquick PCR Purification Columns (Qiagen).
12. Microfuge capable of holding 1.5-mL Eppendorf tubes.
13. Dye primer cycle sequencing ready reaction kits, –21 M13 and M13Rev (Perkin–Elmer).
Because both of the primers used in the PCR amplification of the fragment of
interest have M13 tails, the product can be sequenced in both (forward and
reverse) directions. Store these kits at –20°C.
14. Bucket microfuge capable of spinning a 96-well PCR tray.
15. Ethanol, 95% (v/v).
16. Glycogen, 0.3 mg/mL solution. Store at –20°C.
17. Sample buffer: A 5:1 (v/v) mixture of deionized formamide and 25 mM EDTA,
pH 8.0, containing blue dextran (50 mg/mL).
2.2. Last Hot Cycle PCR
1. Template DNA: Although initial studies are likely to be performed on total DNA
extracted from a tissue homogenate, this technique is perfectly suited to the inves-tigation of mtDNA mutations in single cells. Methods describing the isolation of
DNA from single cells are found in the preceding chapter.
2. PCR amplification: The following stock solutions are required: 2 mM (10×)
dNTPs (Boehringer Mannheim), GeneAmp® PCR buffer containing 100 mM
Tris-HCl, pH 8.3; 500 mM KCl; and 15 mM MgCl2 (Perkin–Elmer), AmpliTaq®
DNA polymerase (Perkin–Elmer).
3. Oligonucleotide primers: These will amplify a region of the mitochondrial
genome containing the mutation of interest. In this case, a 350-basepair fragment
is amplified using L9695 (nt 9695–9717) and H10044 (nt 10044–10022). Stock
solutions (20 µM) are stored at –20°C.
4. Sterile water.
5. PCR tubes: 0.5-mL Thermotubes (Applied Biosystems) are recommended.
6. Ice: All PCR reactions are set up on ice.
Mitochondrial DNA Mutations 271
7. PCR thermal cycler.
8. Horizontal gel electrophoresis equipment.
9. 1% Agarose gels containing ethidium bromide, 1× TAE running buffer.
10. UV transilluminator
11. [α-32P] dCTP (3000 Ci/mmol) (Amersham Life Science Products).
12. Phenol (Molecular Biology grade).
13. Chloroform:isoamyl alcohol (24:1 [v/v]).
14. 7.5 M Ammonium acetate, sterile.
15. Ethanol, 100%.
16. Ethanol, 70% (v/v).
17. Cerenkov counter.
18. Appropriate restriction enzyme supplied with 10× reaction buffer: For the inves-tigation of the T10010C mutation, the restriction endonuclease RsaI (Boehringer
Mannheim) is required.
19. Heat block with variable temperature setting.
20. Vertical electrophoresis system: We regularly use a 16 cm unit (SE 600) manu-factured by Hoefer (Pharmacia Biotech).
21. 5% Nondenaturing polyacrylamide gel: This is made using a 30% polyacryla-mide stock solution (29:1 acrylamide/bisacrylamide [w/w]) and contains 1× TBE
(45 mM Tris-borate, 1 mM EDTA, pH 8.0) as the buffering component.
22. 1× TBE (45 mM Tris-borate, 1 mM EDTA, pH 8.0) running buffer.
23. Gel drying equipment (Bio-Rad Laboratories).
24. Phosphorimage cassette and imaging system, including ImageQuant software
(Molecular Dynamics).
3. Methods
3.1. Automated Sequencing of mtDNA
1. For each PCR amplification, prepare the following reaction at room temperature:
35.75 µL Sterile water
5 µL10× PCR reaction buffer
5 µL10× (2 mM) dNTPs
1.5 µL20 µM forward primer
1.5 µL20 µM reverse primer
1 µLTemplate DNA (200 ng/µL stock)
0.25 µL AmpliTaq Gold™
2. Centrifuge briefly and place in PCR thermal cycler.
3. Perform 30 cycles of amplification as follows:
Initial denaturation 94°C 12 min
30 cycles 94°C 30 s
58°C 30 s
72°C 30 s
Final extension 72°C8 min
272 Taylor et al.
4. Electrophorese samples (5 µL) through a 1% agarose gel containing 0.4 µg/mL of
ethidium bromide for about 1 h, and visualize by UV transillumination.
5. Purify the remaining sample using a QIaquick PCR purification column accord-ing to the manufacturer’s instructions. At the last step, elute the DNA into a vol-ume of water to give a DNA concentration appropriate for the length of fragment
to be sequenced (e.g., a sample of 500 basepairs will need to be at a concentration
of about 7 ng/µL). Once purified, these samples can be stored at –20°C prior to
cycle sequencing.
6. When sequencing, thaw the dye primer cycle sequencing ready reaction mixes
slowly on ice, and vortex to mix well.
7. Combine the following into four separate PCR tubes:
Reagent Reaction: A (µL) C (µL) G (µL) T (µL)
Ready Reaction Premix 4488
PCR product 1122
8. Vortex briefly to mix, and centrifuge to bring the sample to the bottom of the
9. Place the tubes in a thermal cycler, set the reaction volume to “10 µL,” and cycle
sequence using the following linked cycles:
15 Cycles 96°C 10 s
55°C5 s
70°C 60 s
linked to
15 Cycles 96°C 10 s
70°C 60 s
10. Aliquot 80 µL of 95% ethanol and 5 µL of glycogen into a fresh sterile PCR tube.
11. Add the four extension reactions (A, C, G, and T) into the ethanol/glycogen mix,
cover the tubes with aluminum foil, and vortex-mix briefly.
12. Place the tubes on ice and leave for 15 min to precipitate the extension products.
13. Centrifuge in the bucket centrifuge for 15 min (1800gav).
14. Discard the foil and decant the supernatant by inverting the tubes over a paper
towel and centrifuging for a further minute at 100gav.
15. Air-dry the resulting pellets at room temperature for 5 min and replace on ice.
These can be stored at –20°C for several months prior to resuspension and elec-trophoresis.
16. Just prior to electrophoresis, the pellet is dissolved in 3 µL of sample buffer (see
Subheading 2.1.). The DNA sample is heated at 90°C for 2 min, placed on ice
immediately and loaded onto a 6% polyacrylamide gel (29:1 [w/w] acrylamide/
bisacrylamide, Bio-Rad Laboratories) containing 8  M urea that has been
preelectrophoresed for 15 min in 1× TBE buffer. Samples are electrophoresed at
39 W for 14 h, and sequence data collected on an ABI Model 373A automated
DNA sequencer (Applied Biosystems). Factura and Navigator sequence analysis
software (Perkin–Elmer, Applied Biosystems Division) are used to compare and
align sequence files (see Note 3).
Mitochondrial DNA Mutations 273
3.2. Last Hot Cycle PCR
1. For each PCR amplification, prepare the following reaction:
35.75 µL Sterile water
5 µL10× PCR reaction buffer
5 µL10× (2 mM) dNTPs
1.5 µL20 µM forward (L9695) primer
1.5 µL20 µM reverse (H10044) primer
1 µLTemplate DNA (200 ng/µL stock)*
0.25 µL AmpliTaq® DNA polymerase
*If template DNA is from a single cell lysis, reduce the volume of water accord-ingly. A master mix of these components without the template DNA can be made
if multiple samples are being investigated. Overlay with mineral oil if required.
2. Centrifuge briefly and place in PCR thermal cycler
3. Perform 30 cycles of amplification as follows:
Initial denaturation 94°C8 min
30 Cycles 94°C1 min
56°C1 min
72°C1 min
Final extension 72°C8 min
4. Electrophorese samples (5 µL) through a 1% agarose gel containing 0.4 µg/mL
of ethidium bromide and visualize by UV transillumination.
5. If the amplification is successful, add another 1.5  µL of each primer, 0.25  µL
AmpliTaq® DNA polymerase, and 0.5  µL of [α-32P] dCTP (3000 Ci/mmol) to
each reaction, and perform the following cycle:
Denaturation 94°C8 min
Annealing 56°C1 min
Final extension 72°C 12 min
6. Extract the products with 50 µL of phenol, followed by 50 µL of phenol:chloro-form:isoamyl alcohol (25:24:1).
7. Precipitate the labeled products by the addition of 25 µL of 7.5 M ammonium ace-tate (1⁄2 volume) and 100 µL of 100% ethanol (2 vol) and place at –80°C for 1 h.
8. Centrifuge the precipitated products, wash with 70% ethanol, and air-dry.
9. Count the pellets using the Cerenkov counter, and resuspend in sterile water so
that equal amounts (2000–8000 cpm) are digested.
10. Set up the restriction digest in a final volume of 20  µL using the appropriate
restriction endonuclease (5–10 U) and 10× reaction buffer as recommended by
the manufacturer.
11. Separate the digested products through a 5% nondenaturing polyacrylamide gel,
dry the gel, and expose to a PhosphorImage cassette (see Note 4).
12. Using the available software, determine the level of heteroplasmy in the sample
by calculating the amount of radiolabel in each restriction fragment, ensuring
274 Taylor et al.
Fig. 1. PCR-RFLP analysis of mtDNA heteroplasmy in a patient with a pathogenic
T10010C mutation. U, uncut PCR product; C, control subject; M, skeletal muscle show-ing a high level of mutant mtDNA; B, blood that has a much lower level of heteroplasmy.
In the presence of the mutation, the 297-basepair fragment remains uncut.
Mitochondrial DNA Mutations 275
that the digestion has gone to completion. For the T10010C mutation analysis,
there are two  RsaI recognition sites in the wild type product, which generate
fragments of 263, 53, and 34 basepairs. In the presence of the mutation, a site is
lost, leaving two fragments of 297 and 53 basepairs. For quantitation, the 263-basepair
fragment is normalized to the 297-basepair fragment for deoxycytosine content, and
the level of heteroplasmy calculated as a percentage of the amount of radiolabel
in the 263-basepair fragment relative to the combined amount in the 263 and
297-basepair fragments (Fig. 1) (see also Notes 5–7).
4. Notes
1. Although dye primer cycle sequencing typically provides an even peak height
making it easier to detect heteroplasmic base changes, peak height is not directly
proportional to the level of mtDNA heteroplasmy within a sample. Consequently,
proportions of mutant or wild-type mtDNA below 30% are unlikely to be
detected. If a novel change from the Cambridge sequence  (8) is detected that
does not appear to be a recognized polymorphism, last hot cycle PCR-RFLP
analysis should be performed to exclude or confirm the presence of heteroplasmy
and its level within the DNA sample.
2. As previously mentioned, we recommend that both PCR primers are synthesized
with the universal sequencing primer sequences at their 5′ ends, thus allowing
each PCR product to be sequenced bidirectionally and any base substitutions
confirmed. Moreover, there are occasionally small lengths of sequence (usually
less than 5 basepairs) that cannot be read in one direction, presumably due to the
nature of the DNA secondary structure. Sequencing of these samples in the
opposite direction resolves this problem.
3. The complementary DNA strands of the mitochondrial genome have an asym-metric distribution of G’s and C’s, generating a heavy purine-rich H-strand, and a
light pyrimidine-rich L-strand. Sequence data from the H-strand typically show
higher levels of background noise and poorer base-calling than that obtained from
sequencing the L-strand.
4. A sample of the undigested, labeled PCR product must always be run on the gel
to ensure that the restriction digest is 100% efficient.
5. The most critical part of this method is the design of the RFLP to analyze the
mutation. Ideally, the PCR-generated fragment should have two restriction sites
for the enzyme used, one of which is unique to the mutation. This can be either
gain or loss of a site. If the putative mutation does not create or destroy a natu-rally occurring restriction site, it is possible to engineer a site that is specific for
the mutation into the PCR fragment using a mismatch primer (7,18).
6. Instead of performing a last hot cycle, the use of radioactivity can be avoided by
the addition of fluorescent-labeled deoxyuridinetriphosphates (dUTPs) to the
final cycle of PCR, separating the restriction products by nondenaturing poly-acrylamide gel electrophoresis (PAGE) (ABI Model 373A automated DNA
sequencer [Applied Biosystems]), and quantitating the level of heteroplasmy
using Genescan software (Applied Biosystems) (19).
276 Taylor et al.
8. Quantitating the level of heteroplasmy in single cells (e.g., individual muscle
fibers) will confirm whether high levels of the mutation above the required thresh-old for disease expression precipitate an observable biochemical defect. In this
case, single-fiber PCR analysis will reveal significantly higher levels of mutant
mtDNA in COX-deficient fibers than in COX-positive fibers. The protocols
describing the isolation of template DNA from single cells are described in detail
in the preceding chapter.
The financial support of the Muscular Dystrophy Group of Great Brit-ain, the Medical Research Council, and the Wellcome Trust is gratefully
1. Chinnery, P. F. and Turnbull, D. M. (1997) Clinical features, investigation, and
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8. Anderson, S., Bankier, A. T., Barrell, B. G., de Bruijn, M. H., Coulson, A. R.,
Drouin, J., Eperon, I. C., Nierlich, D. P., Roe, B. A., Sanger, F., Schreier, P. H.,
Smith, A. J., Staden, R., and Young, I. G. (1981) Sequence and organisation of the
human mitochondrial genome. Nature 290, 457–465.
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Mammalian mitochondrial genetics: heredity, heteroplasmy and disease.  Trends
Genet . 13, 450–455.
10. Tanaka, M., Hayakawa, M., and Ozawa, T. (1996) Automated sequencing of
Mitochondrial DNA. Methods Enzymol. 264, 407–421.
Mitochondrial DNA Mutations 277
11. Tanno, Y., Yoneda, M., Tanaka, K., Tanaka, H., Yamazaki, M., Nishizawa, M.,
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T-Cell Function in the Aged 281
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Assessment of T-Cell Function in the Aged
T-Cell Proliferative and T-Cell Adherence Assays
Ian Beckman
1. Introduction
Certain immunological activities, particularly cell-mediated immunity,
decline with advancing age. Gaining insight into the underlying mechanism(s)
is complicated by the fact that human T-cells comprise several functionally and
phenotypically distinct populations and subpopulations. Proliferative studies
designed to identify differences between old and young subjects that are based
entirely on unfractionated peripheral blood lymphocytes (PBLs) or pan T-cell
responses run the risk, therefore, of missing subtle but perhaps crucial changes
in a particular T-cell type.
Although it is outside the scope of this chapter to describe T-cell purification
procedures, it is important to emphasize several key considerations before ven-turing down this road.
First, choose several different procedures that operate sequentially rather
than one method repeated several times over. For example, my laboratory
employs simple nylon wool columns to obtain an initial 90–95% purity (nylon
wool-adherent cells also make an excellent source of blood monocytes if
required in subsequent cell mixing experiments). The partially purified T-cells
are then further depleted of residual monocytes by layering over plastic Petri
dishes before being divided into various subpopulations (e.g., CD4+CD45RO+
or CD8+CD28+) either by negative selection, using two rounds of antibody
panning with an appropriate cocktail of monoclonal antibodies (MAbs), or by
positive selection, via flow microfluorimetry sorting (1,2). Magnetic beads are,
of course, another option.
282 Beckman
Whatever the mechanism employed, it is important that the final T-cell frac-tion is free of B cells (and other T-cell subpopulations if desired), monocytes,
and natural killer (NK) cells. For example, “young” T-cells in particular can
respond well to a single mitogenic signal such as phytohemagglutinin (PHA)
with < 1% contaminating monocytes. NK cells, on the other hand, are often
overrepresented in the circulation of aged individuals and because NK cells
can be purified along with T-cells, they not only may skew the final “T” cell
count but also are a potent source of interferon-γ when activated.
Some of the advantages of working with purified populations of T-cells are
that functional changes can be related to a specific cell type(s), and most impor-tantly, when comparing young and aged subjects, the actual number of T-cells
(and indeed, accessory cells) seeded into a given well can be controlled in all
The disadvantages of multiple purification procedures are low cell yields
(despite relatively large volumes of blood) and increased risk of contamination.
It is also important that the purification process per se does not preactivate the
cells. For example, avoid using strongly mitogenic MAbs or sheep red blood
cells (SRBCs) during positive selection. Although T-cell binding to SRBCs
(E-rosettes) is a well-known means of enriching for T-cells, very low numbers
of SRBCs provide sufficient stimulus to activate purified T-cells in the presence
of PHA.
Human peripheral T-cells are normally quiescent. They can be activated to
secrete interleukin-2 (IL-2) and proliferate by a variety of stimuli. In general,
complete T-cell activation requires two signals. The first is delivered through
the T-cell receptor (TCR)–CD3 complex by small antigenic peptides in asso-ciation with HLA molecules. This is best achieved in the laboratory using
reagents such as superantigens or MAbs directed against TCR or CD3. The
first signal can also be triggered by ligands that bind the surface receptor CD2.
Second signals follow engagement of ligands on the surface of antigen present-ing cells (APCs) with T-cell surface receptors (TCRs) such as CD28. However,
a variety of molecules and in vitro systems can bypass the need for APCs or
other accessory cells (ACs), making it possible to dissect specific T-cell–AC
interactions and identify actual sites of dysfunction.
Two potent pan T-cell mitogens, namely MAb OKT3 (anti-CD3) and the
superantigen, staphylococcal enterotoxin B (SEB), have an obligatory require-ment for ACs. However, OKT3-induced activation can be achieved in the
absence of ACs either using wells precoated with anti-mouse IgG, which pro-motes crosslinking of the TCR–CD3 complex, or using plate-bound OKT3 and
fibronectin. The latter combination is a particularly strong T-cell stimulus.
Alternatively, plate-bound 64.1 (an IgM anti-CD3 MAb) alone can induce
proliferation in highly purified T-cells.
T-Cell Function in the Aged 283
With respect to CD2-induced stimulation, mitogens such as PHA and sev-eral anti-CD2 MAbs that trigger the CD2 receptor are powerful activators.
Interestingly, CD2-induced activation is markedly diminished in the elderly
and because PHA is relatively nonspecific, CD2 activation is probably best
observed using the anti-CD2 MAb pair T112 and T113. Together they are
strongly mitogenic for T-cells in the presence of ACs. More importantly, when
used with purified T-cells at relatively low concentrations, this pair of MAbs
provides a potent signal 1 without causing IL-2 secretion. Full activation is
achieved, however, when used in combination with any one of a number of
different costimulatory factors or cells. To this end, various cofactors can eas-ily be tested for their ability to provide an effective signal 2.
Proliferation or activation can be determined in a number of ways. Clearly, the
choice of the primary readout system is best governed by the type of information
required. Other factors, including the availability of technical expertise, the sensi-tivity and specificity of the assay, cost, and time are also important considerations.
The simplest approach is to measure tritiated thymidine incorporation. Prob-ably the best marker of T-cell activation, however, is the production of IL-2 and
other T-cell-derived cytokines. Resting unstimulated human T-cells remain
transcriptionally silent even after several days of in vitro culture. Cytokines
can be detected at the mRNA level by reverse transcriptase-polymerase chain
reaction (RT-PCR),  in situ  hybridization using labeled probes, or by  in situ
PCR. The RT-PCR method (described in detail below) is relatively simple to
perform; it is also fast, sensitive and semiquantitative (if required). Moreover,
it allows the expression of a large range of activation-associated genes to be
examined simultaneously using very small cell numbers, for example,
cytokines and their receptors, other immune function genes (e.g., HLA DR,
LFA–1, transferrin receptor, adhesion receptors, CD80, CD86, CD40), cell-cycle-associated genes, proto-oncogenes etc.
At the protein level, cytokines are readily detected by enzyme-linked
immunosorbent assay (ELISA) (using cell supernatants) and in situ immunohis-tochemistry. The latter technique is very impressive (particularly when coupled
to double or triple labeling and flow cytometry allowing the accurate enumera-tion of specific cell types, e.g., IL-4 producing CD4+CD45RO+ T-cells) but the
trick to getting it to work consistently is obtaining the right MAbs (e.g., see refs.
3–5). It is definitely a procedure worthwhile pursuing in the aging area.
Immunophenotyping and cell-cycle analysis using BrdU or propidium iodide
could also be used to complement the above readouts. Wherever possible, uti-lize a number of different detection systems when comparing T-cell prolifera-tion in aged and young subjects. Finally on the subject of read-out, if “old”
T-cells do not appear to respond to a particular stimulus, check if they have
actually apoptosed.
284 Beckman
T-cell adherence to vascular endothelium is, I believe, an important issue in
immunogerontological studies. To egress the circulation and enter tissues,
T-cells must interact with, and adhere to, endothelial cells. Changes in T-cell–
endothelium interactions may well have implications for atherogenesis and
the observed age-related decline in cell-mediated immunity, particularly
delayed type hypersensitivity (DTH). For example, activated “old” T-cells may
display an increased propensity to “stick” to endothelial cells (EC) and facili-tate inflammation, or conversely, they may exhibit a diminished ability to bind
and transmigrate through the endothelium. To this end, I have included in this
chapter a simple assay that provides a measure of the capacity of various
T-cells to bind human umbilical vein endothelial cells.
2. Materials
1. Complete culture medium (CCM); RPMI-1640 (Flow Labs) supplemented with
10% heat-inactivated fetal calf serum (FCS), 2 mM glutamine (added just prior to
culture) and penicillin/streptomycin (10,000 U/mL) (see Note 1).
2. Sterile flat, 96-well microtiter (Linbro) and 24-well tissue culture (Costar) plates
with lids (see Note 2).
3. Phosphate buffered saline (PBS).
4. 50 mM Tris-HCl, pH 9.5.
5. Tritiated thymidine ([3H]-TdR) (Amersham).
6. Exogenous activators: MAbs (OKT3 [anti-CD3] [from American Type Culture
Collection], 64.1[anti-CD3] and 9.3 [anti-CD28] [from Bristol–Myers–Squibb,
Seattle], T112 and T113 [anti-CD2] [from E. Reinherz, MIT, Boston MA, USA]);
PHA (Sigma), phorbol 12-myristate 13-acetate (PMA) (Sigma), SEB (SEB)
(from Sigma), fibronectin (Collaborative Research Incorporated, or Integrated
Sciences), and various cytokines (Boehringer Mannheim and Genzyme).
7. Goat anti-mouse IgG (GAM) (Dakopatts).
8. Mitomycin C (Sigma).
9. Collagenase (Sigma).
10. Fluorescein-labeled (FITC-) goat or sheep anti-mouse IgG (Becton-Dickinson).
11. Vanyl ribonucleoside complexes (Gibco-BRL).
12. 123-basepair DNA ladder (Gibco-BRL).
13. Endothelial cell growth factor supplement (Integrated Sciences).
14. ECV304: Transformed immortal human endothelial cell line from umbilical cord
(ATCC CRL-1998).
15. Trypsin/EDTA solution (1×) in PBS (Boehringer Mannheim).
3. Methods
3.1. T-Cell Activation Using Unfractionated PBL
1. Add 20 µL or 100 µL of MAb OKT3, at a single predefined optimal concentra-tion (e.g., 50 ng/mL; note the final concentration in each well is 5 ng/mL) or use
T-Cell Function in the Aged 285
a range of concentrations, to triplicate wells of a 96- or 24-well plate, respec-tively (see Note 3).
2. Alternatively or in parallel experiments, add titrated amounts of SEB (a final
concentration of 10 ng/mL is optimal in my laboratory) or MAbs T112 and
T113 for anti-CD2 stimulation (as a guide we use a final dilution each of 1:500
ascites fluid).
3. Dispense 180 µL or 900 µL of PBL (prewashed twice in PBS and resuspended in
CCM at 5.5 × 105/mL) to each 96- or 24-well plate, respectively, and incubate at
37°C in a 5% CO2 controlled atmosphere for 3–4 d. Try a range of cell concentra-tions ranging from 2 × 105/mL to 1 × 106/mL (see Note 4).
4. At the designated time(s), appropiate cultures are spiked with 0.5 µCi [3H]TdR
for 4 h before harvesting the cells using a semiautomated sample harvester. The
amount of [3H]TdR incorporation is measured in a beta scintillation counter (sub-tract the cpm recorded in negative control wells, i.e., wells containing nonac-tivated cells).
Cells are also harvested from parallel cultures at identical times for other analy-ses, for example, RT-PCR, cell cycle, and/or immunohistochemical analysis (see
Note 5). Remember to store the centrifuged culture supernatants at –80°C for
cytokine ELISAs, if required.
Alternatively, T-cell responses generated in one-way mixed lymphocyte reac-tions (MLR) using PBLs are analyzed by coculturing 1  × 105 cells (responder
cells) from one individual with an equal number of irradiated (2000 rads) or mi-tomycin C-treated cells (stimulator cells) from one or more unrelated individuals
in triplicate round-bottom 96-well plates in a final volume of 200 µL (see Note 6).
Incubate for 7 d and spike with 0.5  µCi [3H]TdR per well for 16–18 h before
harvesting as described previously.
3.2. T cell Activation Using Purified T Cells and AC
1. Cultures are basically set up as described previously except purified T-cells are
employed (see Note 7); however, in these experiments the proportion of ACs or
monocytes to T-cells is controlled for each subject.
2. We usually add 10% irradiated (2000 rads) AC-enriched cells (i.e., nylon wool-adherent and plastic-adherent peripheral blood mononuclear cells) to each well
containing 1 × 105 or 1 × 106 T-cells (see Note 8).
Again, it is often informative to try a range of AC concentrations (i.e., 5%
to 30%).
Furthermore, these experiments provide a degree of flexibility. That is, the
ACs can be either (1) irradiated autologous ACs (2) irradiated heterologous
ACs (a MLR is avoided here by culturing for 3 d) (3) transformed cell lines
such as K562 or U937 (irradiated with 4000 rads to prevent outgrowth) (4)
irradiated CHO transfectants expressing specific human ligands singularly or
in combination, for example, HLA DR, CD80, CD86, LFA-3, CD40, Fas, etc.,
or (5), irradiated OKT3–hybridoma cells. Interestingly, the OKT3 expressing
286 Beckman
T3 hybridoma cells satisfy the dual requirements for full activation by provid-ing both first and second signals.
Judicious use of transfectants can help accurately dissect specific T-cell–AC
interactions and thus provide real insight into potential age-related deficiencies.
3.3. AC-Independent T-Cell Activation
3.3.1 Anti-Mouse IgG-Coated Plates
1. Precoat wells with 750 ng/well goat anti-mouse IgG (GAM) prepared in 50 mM
Tris-HCl, pH 9.5.
2. Leave overnight at 4°C, then wash the plates 3× with PBS and store at –20°C
until required.
3. Incubate T cells at 1 × 106/mL with OKT3 (100 ng/mL) for 1 h at 4°C.
4. Wash once in PBS, resuspend in CCM at 5 × 105/mL, and dispense 200 µL per
GAM-coated well. Culture for 3–4 d.
3.3.2. Plate-bound OKT3 + Fibronectin (FN)
1. Precoat wells with 100 ng/well OKT3 in 50 mM Tris-HCl, pH 9.5, and leave
overnight at 4°C.
2. Flick off the supernatant and without washing the plates, add a freshly prepared
solution of FN (1 µg/50 µL/well, diluted in PBS).
3. Incubate at room temperature for 2 h, wash 3× in PBS, and store at 4°C until
4. Dispense 200 µL of T cells at 5 × 105/mL to each well. Culture for 3–4 d.
3.3.3. Immobilized 64.1
MAb 64.1 is diluted in PBS to 1 mg/mL (aliquot and store at –20°C).
1. Add titrated amounts of 64.1 (1000 ng–20 ng/50 µL/well) to flat-well plates and
leave at room temperature for 5 h.
2. Wash 3× in PBS and leave the plates wrapped in alfoil at 4°C (long-term storage
at –20°C).
3. Start the experiment by adding 200 µL of T cells at 5 × 105/mL and culture for
3–4 d (see Note 9).
3.3.4. Anti-CD2-Induced T-Cell Activation in the Presence of Cofactors
1. Titrate both T112 and T113 together to determine a concentration that does not
induce activation in pure T-cell preparations, that is, a submitogenic dose (in our
hands this is usually a 1:1000 or 1:2000 dilution of each ascites fluid).
2. Full activation, including IL-2 secretion, requires a second costimulatory
stimulus. To this end, we have used a variety of cofactors including cytokines
IL-2 (20 U/mL), IL-1β (20 U/mL), IL-6 (200 U/mL), and IL-7 (100 U/mL); PMA
(1 ng/mL); MAb anti-CD44 (1:1000 ascites); and the anti-CD28 MAb 9.3 (1:1000
ascites) (see Note 10). The above concentrations are a guide only.
T-Cell Function in the Aged 287
3. Dispense the anti-CD2 MAbs and graded amounts of cofactor(s) to microtiter
wells, add 2 × 105 T-cells per well, and culture for 3–4 d. Thus the capacity of
each cofactor to induce activation is easily tested. Anti-CD28 MAbs are excellent
costimulatory reagents, presumably because they mimic the key CD28–CD80
signal transduction pathway.
3.4. RT/PCR Analysis
We have established a reliable and semiquantitative RT-PCR technique that
is based on taking several 5-µL aliquots during the linear phase of the PCR and
relating the amount of target product to two control genes, actin and CD3 δ, for
a given starting cell number. However, good commercial kits are now available
for quantitative RT-PCR that are designed for a number of cytokines.
1. After 4–48 h of culture, harvest as few as 2 × 105 and up to 1 × 106 stimulated and
nonstimulated (control) T cells from 24-well Costar plates and wash twice in
PBS-containing 0.01% diethylpyrocarbonate (DEPC) (an RNase inhibitor) (see
Note 11).
2. To extract cytoplasmic RNA, pellet the cells in Eppendorf tubes and remove as
much PBS as possible. Lyse the cell pellets in 0.1 mL of solution A (10 mM
Tris, pH 7.5; 150 mM NaCl; 0.65% NP-40; and 10 mM vanyl ribonucleoside
complexes). After about 10 s, centrifuge the tubes at 12,000g for 1 min and
transfer the supernatant containing the cytoplasmic fraction (be careful not to
disturb the pellet) to a new tube containing 0.3 mL of solution B (10 mM sodium
acetate, pH 5.0; 50 mM NaCl; 5 mM EDTA; and 0.5% sodium dodecyl sulfate
3. Vortex-mix the mixture and then extract twice with 800 µL phenol/chloroform
(1:1) and finally once more with chloroform alone.
4. Remove about 40 µL of the aqueous phase and use an aliquot for the cDNA reac-tion. Store the remainder at –80°C as 1250 or 2500 cell equivalents per microliter
of supernatant. The RNA is stable for at least 12 mo.
5. First-strand cDNA is synthesized from 5 to 10 µL of RNA supernatant, using
M-MLV reverse transcriptase (BRL) and oligo dT priming according to the
manufacturer’s directions, in a final volume of 40 µL. The reaction is carried out
at 37°C for 1 h.
6. One eighth of the cDNA is then added to 45 µL of PCR mix and the tubes sub-jected to 27–34 cycles of PCR amplification using a thermal cycler with a 1 min/
95°C denaturation, 2 min/60°C annealing, and 3 min/72°C extension profile.
7. A 5 µL aliquot is taken at the end of several cycles (e.g., 27, 29, and 31), mixed
with loading buffer, and then analyzed by electrophoresis on 2% agarose gels
and subsequently stained with ethidium bromide (Et Br). A 123-basepair DNA
ladder (Gibco-BRL) is used as a marker. Compare and contrast each gene prod-uct with the actin and CD3 δ product. Thus its possible to screen about 30–40
different genes from a single cell preparation. For relevant primer sequences
see (1).
288 Beckman
Having identified a candidate gene(s) it is important, however, to examine
not just message but where possible, protein levels (ELISA) and functional
integrity (bioassay).
3.5. T-Cell Adherence to Endothelial Cells (ECs)
This simple assay measures the capacity of resting and activated T-cells to
adhere to activated or resting endothelium (see Note 12).
3.5.1. Preparation of Human EC
1. Human ECs are prepared by washing intact human umbilical veins, one end
closed with a sterile stopcock and valve, with 20 mL of Hanks’ balanced salt
solution (HBSS) containing 0.08% collagenase.
2. After 10 min at 37°C, 20 mL of medium 199 is injected into the vein and the
collagenase solution washed out. The vein is massaged with fresh medium and
the stripped EC collected by centrifugation.
3. ECs are cultured in medium 199 supplemented with 20% FCS and 20  µg/mL
Endothelial Cell Growth Factor Supplement (Integrated Sciences) in flasks
(Costar) precoated with 2.5 mL of gelatin (a 2% solution in HBSS).
4. To remove the ECs from a flask, treat with 5 mL of trypsin/EDTA solution for
5 min, then wash twice and resuspend the cells in 3 mL of the above medium.
5. Add 1 mL per well to a 24-well culture plate containing a sterile round glass
coverslip (sterilized by soaking in ethanol, flaming, and washing in sterile PBS).
After 2 d the cells form a new monolayer.
6. Alternatively, monolayers can be derived from the spontaneously transformed
immortal human endothelial cell line, namely ECV304, by culturing cells at
2 × 105/mL in 24-well plates in medium 199 plus 10% FCS.
7. To prepare activated EC monolayers, incubate with 5 ng/mL of tumor necrosis
factor-α (TNF-α) for 4 h and then wash the cells three times with PBS.
3.5.2 Adherence Assay
1. Dispense half a million resting or activated pan T cells (e.g., activation can
achieved by incubating the cells with plate-bound MAb 64.1 for 4 h) to the
EC-coated glass coverslips.
2. After 90 min at 37°C wash the coverslips twice in PBS and incubate with 200 µL
of MAb; for example, anti-CD3, anti-CD4 (helper T cells), anti-CD8 (supressor/
cytotoxic T cells), anti-CD45RA (naive T cells), anti-CD45RO (memory T cells),
or irrelevant MAbs (negative controls).
3. Incubate for 20 min at 4°C and then wash the coverslips twice with PBS.
4. Add 200 µL of pretitrated FITC-labeled goat anti-mouse IgG per well for 20 min
at 4°C.
5. Wash the coverslips thrice and remove them from the wells using a needle with a
bent tip and place on a glass slide for subsequent examination by UV micros-copy. T-cells bound to the EC monolayers are easily identified by their staining
pattern with the appropriate MAb.
T-Cell Function in the Aged 289
4. Notes
1. Test several different FCS batches and select one that is neither inhibitory in the
presence of known activators nor mitogenic in the absence of activators. It should
also be relatively cheap and available in sufficient quantity to complete all experi-ments. FCS can be replaced with pooled human AB sera, however, varying
amounts of platelet-derived growth factor (PDGF) in human sera may alter T-cell
activity, particularly cytokine production. “Old” T-cells are reported to be sensi-tive to PDGF. If possible, therefore, use a proven serum-free medium or supple-ment, such as Stratagenes Cell/Perfect PBL serum-free media supplement, that
consistently supports T-cell proliferation in your hands.
2. I recommend flat 96-well microtiter plates (Linbro) or 24-well plates (Costar)
with lids but different types and brands should be compared. Some plates are
clearly better at supporting cellular proliferation than others. (Note that round-bottom plates are optimal for MLR).
3. Identify the optimal and suboptimal concentrations of all exogenous stimuli by
running dose–response curves and kinetic studies over several days. The impor-tance of these experiments cannot be overstated, and they should be performed
on cells from both aged and young control subjects. Such experiments also help
to define the experimental conditions, and indeed help exploit the differences
that may exist between age groups. For example, “old” T-cells that do not respond
to a particular stimulus at one concentration may respond well at a higher con-centration or a day later, when compared to “young” cells.
4. Determine cell viabilities (should be >95%). If <90%, either abort the experi-ment (and potentially save on time and money in the long term) or try to remove
the damaged cells (e.g., by Percoll gradient centrifugation or flow cytometry).
Dead or dying cells can influence the final outcome of cultures significantly.
5. In my experience, some of the different readout systems do not always correlate
with each other and vital information may be lost if just one or two systems are
used to measure the proliferative response. For example, I have observed acti-vated T-cells from aged subjects that produced very low levels of IL-2 but still
displayed normal levels of tritiated thymidine incorporation, and vice versa.
6. Prepare a solution of mitomycin C in PBS at 0.5 mg/mL, filter sterilize, and store
the bottle in the dark (mitomycin C is light sensitive). Incubate PBL at 1 × 106/mL
in PBS with 50 µg/mL of mitomycin C for 20 min at 37 C. Wash the cells twice
with large volumes of CCM and resuspend the pellet in CCM at 1 × 106/mL.
7. To ascertain that accessory cells or monocytes have been removed, albeit func-tionally, culture the T-cell preparations alone with PHA (5 µg/mL) for 3–4 d and
measure the amount of tritiated thymidine uptake. Residual NK cells, B cells,
and monocytes can also be checked by immunophenotyping and flow cytometry
using appropriate MAbs.
8. AC-enriched populations are obtained from spent nylon-wool columns (i.e., after
the T-cells have been eluted) by loading the column with warm medium and vig-orously and continously proding the nylon-wool with a sterile 10-mL pipet. Col-lect the adherent cell eluate and plate out on plastic Petri dishes for 1–2 h. Wash
290 Beckman
the plates vigorously with medium and harvest the plastic-adherent cells by gen-tly scraping with the rubber end of a 10-mL syringe plunger.
9. Proliferative T-cell responses to immobilized 64.1 are significantly enhanced by
the addition of IL-1β (20 U/mL), but only at the relatively lower antibody con-centrations of 100 ng–20 ng/well. In our hands this is a simple but very effective
bioassay for IL-1β.
10. It is advisable to use well-defined reagents (e.g., antibodies, cytokines, etc.) that
can be purchased or prepared in purified form. Dialyze extensively against PBS
to remove preservatives and other potentially inhibitory factors and then filter
sterilize. Check for endotoxin activity and confirm, where possible, the specific-ity of the reagents before aliquoting and placing in long-term storage.
11. IL-2 (and other cytokine transcripts) may be detected in unstimulated or control
T-cells by RT-PCR if the medium contains even trace amounts of mycoplasma or
nonspecific serum-derived mitogenic factors. This is another good reason to use
serum-free medium.
12. We have tried several methods based on differential prestaining of T-cells and
ECs with a number of dyes to detect T-cells bound to EC monolayers but with
mixed success, largely due to background staining from the ECs. However, an
indirect immunofluorescence method proved to be fast and effective, and allowed
identification of specific T-cell subtypes without the need to first purify these
cells before addition to the EC monolayers. If background staining is a problem,
block with normal goat serum or try other second antibodies.
1. Beckman, I., Shepherd, K., Dimopolous, K., Ahern, M., Firgaira, F., and Bradley, J.
(1994) Differential regulation and expression of cytokine mRNA’s in human CD45R
T cell subsets. Cytokine 6, 116–123.
2. Beckman, I., Shepard, K., Ahern, M., and Firgaira, F. (1995) Age-related defects in
CD2-receptor induced activation in human T cell subsets. Immunology 86, 533–536.
3. Sewell, W. A., North, M. E., Webster, A. D., and Farrant, J. (1997) Determination of
intracellular cytokines by flow-cytometry following whole blood culture. J. Immunol.
Methods 10, 67–74.
4. Morita, Y., Yamamura, M., Kawashima, M., Harada, S., Tsuji, K., Shiuya, K.,
Maruyama, K., and Makino, H. (1998) Flow cytometric single cell analysis of
cytokine production by CD4+ T cells in synovial tissue and peripheral blood from
patients with rheumatoid arthritis. Arthritis Rheum. 41, 1669–1676.
5. Collins, D. P., Luebering, B. J., and Shaut, D. M. (1998) T-lymphocyte functionally
assessed by analysis of cytokine receptor expression, intracellular cytokine expres-sion, and femtomolar detection of cytokine secretion by quantitative flow cytometry.
Cytometry 33, 249–255.
Dendritic Cells in Old Age 291
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Dendritic Cells in Old Age
Beatrix Grubeck-Loebenstein, Maria Saurwein-Teissl,
and Nikolaus Romani
1. Introduction
1.1. General Background
Dendritic cells (DCs) are powerful antigen-presenting cells that have the unique
capacity to stimulate naive T-cells (1,2). DCs are identified by a triad of criteria:
Morphologically, they exhibit pronounced cytoplasmic veils that are mobile and
can easily be observed under a phase-contrast microscope. These veils become
apparent only in the mature state. Phenotypically, they express high levels of
major histocompatibility class (MHC) (class I and II), adhesion (CD11c, CD54,
CD58), and costimulatory (CD80, CD86, CD40) molecules on their cell sur-faces. They also express CD1a and CD83, but lack CD14. On cytocentrifuge
smears stained with anti-CD68, a marker of the endocytic system that is abun-dant in macrophages, DCs display spotlike staining whereas typical macrophages
are strongly positive all over the cytoplasm. When looking at forward/side scat-ter profiles in the fluorescence-activated cell sorter (FACS), DCs show high
light scattering and are outside the typical lymphocyte gate. Functionally, they
are potent stimulators of resting T lymphocytes in the allogeneic mixed leuko-cyte reaction. DCs derived from various tissues have been shown to undergo a
complex maturation process during which their morphology, phenotype, and
function change. DCs are derived from bone marrow progenitors and circulate
in the blood as immature precursors before they migrate into peripheral tissues,
such as the epidermis, heart, lung, liver, gut, thymus, spleen, and lymph nodes.
DCs of myeloid as well as of lymphoid origin have been described (3–5). Within
tissues DCs take up and process antigen which is then presented in the context
of MHC molecules. Upon appropriate stimulation they undergo further matura-tion and migrate to secondary lymphoid tissue where they present antigens to
292 Grubeck-Loebenstein, Saurwein-Teissl, and Romani
T-cells and induce an immune response (6). Recent knowledge on DCs and how
they control immunity has been summarized in excellent reviews (4,6–9). It is
the aim of the present chapter to give a brief overview on what is known on DCs
in aged humans and animals. Particular emphasis is given to outline the method-ology required to purify, culture, and characterize this interesting cell type.
1.2. How Studies on DCs Can Be Performed
In the past, DCs from various sources could be isolated only by using sophis-ticated protocols  (10–12). These protocols are still being used for studies in
which freshly isolated DCs have to be analyzed. Rapid progress in the cha-racterization of DCs has, however, been made in recent years, as simple
methods to generate large numbers of DCs from precursors have been devel-oped. CD34+ progenitor cells from cord blood or bone marrow or monocytes
from the peripheral blood can hereby be used as starting populations (13–15).
Depending on the nature of these populations and on the stimulatory agents
applied, DCs with a varying degree of maturation can be propagated (16,17).
Although granulocyte/macrophage colony-stimulating factor (GM-CSF) and
tumor necrosis factor-α (TNF-α) have mainly been used to induce the genera-tion of a typical DC progeny from CD34+ cells, combinations of GM-CSF and
interleukin (IL-4/IL-13) have been shown to trigger the differentiation of mono-cytes into typical DCs. Various factors, such as serum, monocyte conditioned
medium (18,19), transforming growth factor-β (TGF-β) (20,21), flt-3 (22), and
stem cell factor  (23) have been shown to support the in vitro generation of
DCs. Organ cultures to study the outgrowth of DCs from tissue samples have
additionally been developed (18).
1.3. DCs in Aged Rodents
1.3.1. Murine DCs in Aging
Inbred strains of laboratory mice represent well-defined and suitable sys-tems to study effects of aging on DCs. It is easy to obtain large numbers of
animals at defined ages. There is even a special strain of mice that may be used
for senescence-related work: the SAMP1 (senescence-accelerated) mouse (24).
In spite of such good preconditions, little research has been done with regard to
DCs and aging. Yet, some interesting data are available and are reviewed here.
1.3.2. Ontogeny of DCs
As mentioned previously, DCs arise from progenitors in the bone marrow.
One study has addressed whether the capacity of the bone marrow to give rise
to Langerhans cells (LCs) changes with age  (25). When bone marrow cells
from young and old mice were transplanted into irradiated recipients of the
same age it was observed that the density of LCs was lower in those mice that
Dendritic Cells in Old Age 293
had received marrow cells from aged animals than in those mice that had been
grafted with marrow from young mice. This suggests that the frequency of DC
progenitors may be lower in aged mice. Alternatively, the migration of pro-genitors from the marrow to their target tissues or the homing of progenitors
may be impaired in aged animals. These possibilities are yet to be clarified.
1.3.3. Distribution of DCs
LCs, that is, the epidermal variant of DCs, have been studied in some detail,
the most obvious question being whether the numbers of LCs change with age.
Immunohistochemistry (using monoclonal antibodies [MAbs] against MHC class
II) and enzyme histochemistry (using ATPase staining) were generally applied to
address this issue. Several authors found a decline of LC densities in the epider-mis with age (25–27). The extent of LC reduction was about one third. This was
also observed with LCs in the oral mucosa: a reduction by 30–60% was reported
in the mouse (28) and a reduction of LC foci and interfocal LCs in the hamster
cheek pouch (29). In contrast, a study in the rat reported an unchanged density of
DCs (reactive with MAb OX-6) in the mucosa of the tooth pulp  (30). So did
another investigation of the mouse gingiva (31). No detailed studies are available
concerning age-related changes in numbers and distribution of DCs from sources
other than the skin (spleen, lymph nodes, thymus).
1.3.4. Function of DCs
The experiments reviewed here are too few in number to allow well-founded
conclusions. Important questions have not yet been addressed. It has not been
dissected which component of DC function is reduced with age. The three broad
functions of DCs as delineated by Steinman (1; see Subheading 1.1.) are sepa-rated from each other in space and time. They are down- and up-regulated in
the course of the maturation process. Therefore, the experiments discussed
below also imply an analysis of the maturation process in relation to age.
1.3.5. T-Cell Stimulatory Properties of Murine DCs
Antigen processing may be affected by age. The observations by Haruna et
al. (24) that the impaired T-cell stimulatory function of DCs did not correlate
with the expression of such crucial molecules as the costimulators CD80 and
CD86 leads one to speculate that perhaps the generation of immunogenic MHC
class II/peptide complexes may be diminished as well. This aspect of immuno-genicity could easily be studied by means of peptide-specific T-cell hybrido-mas (32). The development of T-cell-sensitizing capacity of DCs with age is
another important point. Early work showed an increase of the immuno-stimulatory capacity of spleen DCs from C57BL/6 mice in the syngeneic mixed
leukocyte reaction, but not in the allogeneic mixed leukocyte reaction or in the
294 Grubeck-Loebenstein, Saurwein-Teissl, and Romani
polyclonal response to concanavalin A (33). A recent and detailed study uti-lized a substrain of SAMP1 mice. The authors investigated the allogeneic mixed
leukocyte reaction as a measure for the capacity of DCs to stimulate resting
T-cells. Spleen DCs from aged SAMP1 mice stimulated less well in this assay
than DCs derived from age-matched BALB/c mice or from young SAMP1 mice
(24). These changes correlated with the expression of MHC class II and adhe-sion molecules (CD54/ICAM–1) but, interestingly, not with the expression of
costimulator molecules (CD80/B7–1 and CD86/B7–2) on DCs. Clearly, more
studies are needed to construct a conclusive picture. Other factors contributing
to the sensitizing potential of murine DCs have not been analyzed to date.
Examples are the capacity to cluster T-cells in vitro (34) and in vivo (35) or the
ability to secrete IL-12 (36).
1.3.6. Migratory Function
Migration of DCs from the sites of antigen uptake (e.g. the skin) to the
regional lymphatic organs is essential for the successful generation of an
immune response. DCs typically migrate through lymphatic vessels. Recently,
a novel pathway of DC migration was identified, namely a transition of DCs
from the blood into the lymph in the liver (37). It would be interesting to know
whether migration is impaired in aged mice. Indirect evidence for this stems
from contact hypersensitivity experiments. The induction of contact hyper-sensitivity correlates with an efflux of LCs from the epidermis and their entry
into dermal lymphatics (38). Sprecher et al. (25) noted that the capacity of LCs
to transport antigen (the contact sensitizer rhodamine) from the skin to the
draining lymph nodes was not impaired in old mice. The clinical outcome of
this antigen transport, that is contact hypersensitivity, was found to be reduced
in one study with aged mice (39), and more variable but not reduced in another
study (27). No consensus has thus yet been reached regarding DC migration
during aging. Skin organ cultures (38,40) may be a suitable experimental sys-tem to directly study this issue in more depth.
1.4. DCs in Aged Humans
1.4.1. Ontogeny and Distribution of DCs
Only limited information is at present available on DCs in aged humans. It is
therefore not clear whether the frequency of DCs progenitors decreases during
the aging process. No studies have ever addressed the question whether the
capacity of the bone marrow to generate DCs changes with age. Early studies
have demonstrated a decreased density and functional activity of LCs in the
aged skin (41,42). This may, however, partly be due to UV irradiation, as sig-
Dendritic Cells in Old Age 295
nificantly fewer LCs were observed in exposed vs covered skin in old individu-als, while no such disparity was noted in the younger subjects.
1.4.2. Characterization of DCs Generated from Blood Monocytes
A detailed analysis of functional and phenotypic characteristics of DCs from
aged humans has only recently been performed, when large numbers of DCs
were generated from blood monocytes of healthy aged persons (43). After 1 wk
of culture in GM-CSF and IL-4 these cells have a typical dendritic morphology,
have an intact capacity of phagocytosis, and express DC surface marker mol-ecules in the same way as corresponding populations from young persons
(43,44). They represent a relatively homogeneous population of intermediate
maturity, which matures further in response to stimulation with pathogens.
1.4.3. Maturation of Human DCs
Factors such as pathogens (45–47) and cytokines (48,49) have been demon-strated to stimulate the maturation of DCs in vitro and in vivo. The responsive-ness of DCs from elderly humans to stimuli has been tested in a recent study, in
which it has been demonstrated that inactivated influenza virus induces DC
maturation (50). Whereas unstimulated blood-derived DCs from old and young
healthy individuals express MHC class II, CD54, CD80, and CD86 on their
surface at medium density and secrete moderate amounts of IL-12 and TNF-α,
stimulation with inactivated influenza virus leads to a marked increase in the
production of surface molecules and cytokines. These changes are equally pro-nounced in cells from old and young individuals. The results demonstrate that
DC responsiveness to stimulation with certain vaccines is unimpaired in old
age. DCs may therefore represent a suitable tool for immunotherapy and may
increase the efficacy of vaccines in the elderly.
1.4.4. T-Cell Stimulatory Properties of DCs from Aged Humans
Monocyte-derived DCs from aged persons are equally effective as young
control cells in presenting antigen to tetanus-specific T-cell clones  (43) and
have even been shown to rescue aged T-cells in an in vitro senescence model
from terminal apoptosis (44). In this model, tetanus toxoid (TT)-specific T-cell
lines are derived from young and old individuals and kept in long-term culture.
T-cell proliferation in response to stimulation with antigen presented by either
irradiated autologous peripheral blood mononuclear cells (PBMCs) or DCs is
assessed soon after the initiation of the cultures and after 20 and 30 population
doublings. At this late stage T-cell growth is characteristically slow and pro-grammed cell death imminent. Antigen presentation by DCs enhances T-cell
proliferation at each time point and reinduces proliferation in T-cell popula-tions that have stopped dividing. Terminal apoptosis is thus prevented. DCs
296 Grubeck-Loebenstein, Saurwein-Teissl, and Romani
from old individuals are hereby as effective as cells from young donors. The
results of the study demonstrate that DCs from aged individuals may still stimu-late the clonal expansion and postpone the clonal elimination of antigen-spe-cific T-cell populations. As a consequence DCs may increase immunoreactivity,
prolong immunological memory, and be of particular importance for the main-tenance of the T-cell repertoire in old age.
It must, however, be pointed out that the aforementioned studies concen-trated only on the stimulation of cloned T-cell lines, which usually do not return
to a completely resting stage, but remain partly activated during long-term
culture. The described system, therefore, rather represents a model of a
secondary than of a primary immune response. No data are presently available
on the capacity of DCs of aged humans to activate naive and resting T cells. In
view of the increased clinical use of DCs as vaccine carriers, studies on this
topic would be of utmost importance.
1.4.5. Migratory function
Interestingly, higher numbers of DCs can be derived from the blood of aged
than of young individuals (43). In view of the low numbers of DCs in the aged
skin (41,42), this result suggests that the migration of DCs/DC precursors from
the bloodstream to peripheral organs is affected by the aging process. As the
expression of surface molecules is unimpaired in DCs from elderly persons —
at least following in vitro culture — it seems possible that the DCs themselves
meet the necessary requirement for migration, but that their mobility is reduced
owing to age-specific changes of the endothelium or the extracellular matrix.
1.5. Methods Described in this Chapter
Depending on the aim of the study, a choice has to be made of which method
to use for the purification or generation of DCs. As only few studies on lym-phoid DCs have so far been published (3–5) and there is virtually no informa-tion on lymphoid DCs in the aged, this chapter concentrates on DCs of the
myeloid lineage and describes seven short protocols that we consider particu-larly suitable for the characterization of DCs from aged humans and mice. We
would still like to point out that the methodologies described here represent
only a very small part of the published literature. We would therefore like to
refer the interested reader to some new excellent reviews and textbooks on
methodological aspects of DCs (18,51–53).
2. Materials
2.1. Purification of DCs (LCs) from Murine Skin
1. Utensils: Two pairs of thin but strong forceps with rounded tips (anatomical type);
two pairs of thin, curved, and pointed forceps; flat-bottom tea strainer with handle
Dendritic Cells in Old Age 297
to fit into a 100 mm Petri dish (e.g. “cell dissociation sieve” CD-1, Sigma, St.
Louis, MO, USA); nylon gauze with a mesh size of about 40 mm (NITEX
3–325–44; Tetko, Elmsford, NY, USA; alternatively, nylon gauze with a wide
range of mesh sizes may be obtained from suppliers for the graphic arts); tissue
culture 100 mm dishes (e.g. Falcon, Oxnard, CA, USA, cat. no. 3003) and bacte-riological 100 mm Petri dishes (e.g. Falcon no. 1029).
2. Culture medium: RPMI-1640 supplemented with 5% or 10% fetal calf serum
(FCS), 50 mM 2-mercaptoethanol, 200 mM L-glutamine, and 20 mg/mL of
3. Cytotoxicity medium: For complement-mediated cytotoxicity RPMI-1640 con-taining 25 mM N-2-hydroxyethylpiperazine-N ‘-2-ethanesulfonic acid (HEPES)
buffer and 0.3% bovine serum albumin (BSA), pH 7.2, is used.
4. Complement: Rabbit Low-Tox-M complement from Cedarlane Laboratories,
Hornby, ON, Canada.
5. Salines: Phosphate-buffered saline (PBS) and Hank’s balanced salt solution
(HBSS), both without calcium and magnesium salts.
6. Trypsin: 2.5% commercial stock, aliquoted and frozen at –20°C.
7. Deoxyribonuclease (DNase I): (Boehringer Mannheim, Mannheim, Germany;
cat. no. 104 159). Prepare a stock solution of 5 mg/mL in PBS, sterile filter, and
store at 4°C for up to 3 mo.
8. Antibodies used for enrichment of LCs: Mouse IgM anti-Thy-1/CD90 (ATCC
no. TIB99); rat IgG2b anti-I-Ab,d/clone B21–2 (no. TIB229), and mouse IgG2a
anti-I-Ek,d/clone 14-4-4S (no. HB32) can be used as hybridoma culture superna-tants. Hybridoma cells can be obtained from the American Type Culture Collec-tion (ATCC), Rockville, MD, USA.
Although LCs can be prepared from body skin, the ears are clearly the best
source for obtaining highly enriched LCs (see Note 1).
2.2. Purification of DCs (LCs) from Human Skin
One best uses split thickness skin (300–400 µm thick) that is obtained from
patients undergoing reconstructive plastic surgery of breasts or abdomen, or
less preferably from cadaver skin (within 24 h of death). Aged skin samples
may sometimes be obtainable as a byproduct of hernia operations. Use mate-rials, utensils, and reagents as for murine LCs.
2.3. Purification of DCs from Human Blood
This protocol works only for the purification of human DCs, as murine blood
is available in too small amounts.
1. Culture medium: RPMI-1640 supplemented with 10% heat-inactivated (56°C,
30 min) human serum, 20 µg/mL of gentamicin, and 10 mM HEPES.
2. Chelating washing medium: HBSS without Ca2+ and Mg2+ containing 1% BSA
and 1 mM EDTA.
298 Grubeck-Loebenstein, Saurwein-Teissl, and Romani
3. Neuraminidase-treated sheep erythrocytes for E rosetting: Sheep red blood cells
(SRBC; Cocalico, Reamstown, PA, USA) are washed 3× with RPMI-1640 at
room temperature and centrifuged with 460g. A 5% SRBC solution is prepared.
Add neuraminidase (Calbiochem, La Jolla, CA, USA; or Behring, Marburg, Ger-many) to give a final activity of 0.01 U/mL. Incubate for 1 h at 37°C. Centrifuge
and wash two more times at 460g with warm PBS without Ca2+ and Mg2+. Store
on ice and use on the same day.
4. Antibodies for sorting DC: A “cocktail” of MAbs against CD3 (Becton-Dickinson Immunocytometry Systems, Mountain View, CA, USA), CD11b
(OKM1, ATCC, Rockville, MD, USA), CD16 (3G8; Immunotech-Coulter,
Marseille, France), and CD19 (J4.119; Immunotech) is prepared. Antibody
concentrations in the cocktail should be chosen according to manufacturers’
instructions. Antibodies with well-defined CD specificities from other reli-able manufacturers may also be used. Furthermore, phycoerythrin (PE)-con-jugated anti-HLA-DR, fluorescein isothiocyanate (FITC)-conjugated goat
anti-mouse IgG + IgM (Jackson Immunoresearch, Westgrove, PA, USA), and
mouse IgG (Jackson) are used to identify the HLA-DR-positive DC within
the “cocktail negative” population.
5. Monocyte-conditioned medium: Fresh PBMCs are adhered to Petri dishes that
have been coated with human γ-globulin. 50 × 106 PBMC are plated in 10-mL
culture medium per 100 mm dish. After 30 min at 37°C, nonadherent cells are
washed away and adherent cells are cultured in culture medium for 24 h. Super-natants are collected, centrifuged, tested for activity, aliquoted, and stored at
–20°C until needed.
6. Coating: Bacteriological Petri dishes (e.g., Falcon no. 1029) are covered with a
solution of human Ig (Calbiochem, La Jolla, CA, USA) in PBS at a concentration
of 10 µg/mL and incubated at room temperature for 1 h. After four washes with
PBS the dishes are ready. Dishes may be coated on the day before use; they should
not be stored longer, however.
2.4. Generation of DCs from Human Blood Monocytes
1. Culture medium: RPMI-1640 supplemented with 5–10% FCS,  L-glutamine,
2-mercaptoethanol, and gentamicin (all from Seromed-Biochrom, Berlin, Ger-many). To generate DCs for clinical use FCS must be avoided as well as any other
nonhuman proteins.
2. Multiwell tissue culture plates (24-well or 6-well).
3. Cytokines: Recombinant human cytokines are used throughout. GM-CSF
(Novartis, Basel, Switzerland) and IL-4 (Genzyme, Cambridge, MA, USA).
2.5. Generation of DC from Human Blood CD34+  Progenitors
1. Culture medium: RPMI-1640 supplemented with 10% FCS.
2. MiniMACS Multisort kit (Miltenyi Biotech, Bergisch-Gladbach) for purification
of CD34+ cells.
3. Antibody for purity control: CD34 mAB (HPCA-2, PE conjugated).
Dendritic Cells in Old Age 299
4. Cytokines: Recombinant human cytokines are used: GM-CSF (Novartis, Basel,
Switzerland), TNF-α (Bender, Vienna, Austria), flt-3 (Neupogen; Amgen, Thou-sand Oaks, CA, USA), stem cell factor (rhSCF, Amgen, Thousand Oaks, CA,
USA), TGF-β (British Biotechnology, Abington, UK).
2.6. Generation of DCs from Murine Bone Marrow (53)
1. Utensils: Autoclaved strong scissors, anatomical forceps, 5- or 10-mL syringe
with a 25-gauge needle.
2. Culture medium: RPMI-1640 supplemented with 10% FCS, 50 mM 2-mercapto-ethanol, and 20 mg/mL gentamicin.
3. Cytokines: Recombinant murine cytokines are used: GM-CSF (e.g. from
Immunex, Seattle, WA, USA; specific activity 4 × 107 U/mg); TNF-α (e.g. from
Bender, Vienna, Austria; specific activity 2.6 × 107 U/mg)
4. Culture vessels: 24-well plates and 60 or 100 mm tissue culture dishes.
5. Cytotoxicity medium: For complement-mediated cytotoxicity RPMI-1640 con-taining 25 mM HEPES buffer and 0.3% BSA, pH 7.2, is used.
6. Complement: Rabbit Low-Tox-M complement from Cedarlane Laboratories,
Hornby, ON, Canada.
7. Antibodies used for enrichment of DC: anti-B220 (RA3-3A1, no. TIB146, ATCC)
for B cells, anti-CD4 (GK1.5, no. TIB207) and anti-CD8 (HO-2.2, no. TIB150)
for T-cells, RB6-8C5 for granulocytes, and anti-MHC class II (e.g., B21-2, no.
TIB229 for BALB/c mice).
2.7. Generation of DCs from Human Bone Marrow
1. Culture medium: RPMI-1640 supplemented with 10%FCS.
2. Cytokines: Recombinant human cytokines are used: GM-CSF (Novartis, Basel,
Switzerland), TNF-α (Bender, Vienna, Austria), SCF (rhSCF, Amgen, Thousand
Oaks, CA, USA).
3. Methods
3.1. Purification of DCs (LCs) from Murine Skin
1. Rinse mice ears briefly twice in 70% ethanol, place on sterile gauze in a Petri
dish and allow to air-dry for approx 20–30 min. Split ears. Place the ear halves
separately into two Petri dishes with dermal side down in a way that they float
and do not submerge.
2. Incubate the ear halves in 1% (ventral sides) and 0.33% (dorsal sides) of trypsin
in HBSS at 37°C for 30 (dorsal) to 60 (ventral) min (see Note 2).
3. Carefully aspirate the trypsin solution. Peel off the epidermal sheets from the
underlying dermis. The extent of trypsinization is optimal when the sheets can be
pulled off in one piece. Put the sheets dermal side down in a tissue culture dish
containing a tea strainer. Shake the tea strainer for 3 min to release basal layer
cells into the medium.
4. After removing the strainer, resuspend the cells using a 5- or 10-mL pipet. Filter
the suspension through a nylon mesh and centrifuge it in 50-mL tubes at 300g for
300 Grubeck-Loebenstein, Saurwein-Teissl, and Romani
10 min. Wash twice in culture medium afterwards. At this point the cells may
either be processed further to enrich LCs (step 5a) or be put in culture (step 5b).
5a. To preenrich LCs, epidermal cells can be treated with anti-Thy-1 and comple-ment to remove most keratinocytes and lymphocytes. In a 50-mL polypropylene
tube, resuspend 100–150 × 106 epidermal cells (the expected yield from 60 ears)
in 3 mL of hybridoma culture supernatant of MAb anti-Thy–1 (ATCC no. TIB99).
Then add 10 mL of a sterile-filtered solution consisting of 8.5 mL of cytotoxicity
medium, 1 mL of reconstituted complement, and 0.5 mL of DNase stock solu-tion. After incubation for 1 h at 37°C in a shaking water bath, the resulting epi-dermal cell suspension (viability 10–20%) is washed twice with cold PBS. Treat
cells for 10 min at 37°C with 0.125% trypsin and 80 µg/mL of DNase in PBS at a
cell concentration of 1–2  × 106 viable cells/mL. To stop the digestion, add an
equal volume of culture medium and centrifuge the cells for 10 min at 4°C, at
200g with brakes off. This simple procedure (“trypsin trick”) removes most dead
cells and results in viable (>90%) epidermal cell suspensions containing about
15% (range 10–28%) LCs, which can then be further purified by positive selec-tion by sorting (by FACS or MACS, Miltenyi, 1990) for, for example, MHC class
II (see Note 3).
5b. Untreated or anti-Thy–1/C’ treated (i.e. LC preenriched) epidermal cell suspen-sions can be cultured for 2–4 d after which the LCs have matured and can be
enriched to 60–90% by simple Ficoll–Hypaque/Lymphoprep density gradient
centrifugation according to standard methodology of the nonadherent fraction
and harvesting of the low-density fraction (see Note 4).
3.2. Purification of DCs (LCs) from Human Skin
1. Incubate skin for about 1 h in culture medium containing 25 mM HEPES and
10-fold concentrated gentamicin at room temperature (see Note 5).
2. Rinse the pieces of skin twice in PBS in 100 mm Petri dishes. Cut it into pieces
(1 cm2 if thin, 15 mm2 if thick) and put them into another 100 mm Petri dish
containing 11.25 mL of PBS. When correctly placed epidermal side up, the skin
will spread on the PBS. When all pieces are floating on PBS add 1.25 mL of 2.5%
3. Incubate with trypsin overnight at 4°C.
4. Next morning aspirate the medium from the dishes and remove epidermal sheets
from the dermis with two thin forceps. Put epidermal sheets into a tea strainer
placed in another 100 mm Petri dish with 30 mL of culture medium containing
80 µg/mL of DNase I and 25 mM HEPES buffer. The sheets should not overlap.
When the strainer is filled with sheets, stir  vigorously  to release single cells,
remove the strainer from the dish, filter the cell suspension through a Nitex nylon
mesh (54 µm mesh size), and centrifuge it in 50-mL tubes at 300g for 10 min
followed by two washes in culture medium. The primary epidermal cell suspen-sion contains between 1% and 3% LCs (see Note 6).
5. Culture of LCs: Plate epidermal cells in culture medium in culture vessels of
your choice.
Dendritic Cells in Old Age 301
6. Harvest of cultured LCs: After 3 d of culture nonadherent cells are rinsed off the
tissue culture vessels and collected. This cell suspension has a very low viability
(sometimes <20%). Under the hemocytometer one can readily identify very
“hairy” LCs. Further enrichment of cultured LCs can be achieved by a variety of
different methods such as fluorescent cell sorting, panning, magnetic bead tech-niques, and density separations (see Note 7).
3.3. Purification of DCs from Human Blood
1. PBMCs are obtained by standard Ficoll–Hypaque/Lymphoprep centrifugation
(see Note 8).
2. T-cell depletion by E rosetting: PBMCs are resuspended in 10% human serum at
a density of 20  × 106/mL. Add an equal volume of 5% neuraminidase-treated
SRBC. Centrifuge gently at 25g at 4°C for 5 min and incubate on ice for 1 h.
Resuspend very carefully so as not to disrupt the rosettes that have formed. Layer
resuspended cells on a Ficoll–Hypaque/Lymphoprep gradient (20–30 mL of cell
suspension per gradient tube). Harvest nonrosetted cells (Er-) from the interface.
3. B, NK, and monocyte depletion by panning: Er- cells are mixed with a cocktail of
MAb’s against CD11b, CD16, CD19, and CD3. Cells are incubated with the
cocktail for 30 min on ice, washed with chelating medium and added to dishes
coated with goat anti-mouse IgG. A maximum of 30 × 106 cells may be loaded
onto one 100 mm Petri dish. Dishes are incubated on ice for 30 min. Nonadherent
cells are collected in a tube and processed further for cell sorting. Instead of
panning one can, of course, also use immunomagnetic depletion, for example, by the
Dynabead (Dynal, Oslo, Norway) or MACS (Miltenyi Biotec GmbH, Bergisch
Gladbach, Germany) technology, although this approach is more expensive.
4. Positive selection of DCs by two-color cell sorting: Nonadherent cells are incu-bated for 30 min on ice with a FITC-conjugated goat anti-mouse IgG + IgM
antibody, to label any cells that carry a mouse monoclonal antibody but have not
been depleted by the preceding panning procedure. This is followed by a
quenching step of 15 min on ice with 100 µg/mL of mouse Ig (to quench free
FITC-goat anti-mouse IgG + IgM binding sites) and finally PE-conjugated anti-HLA-DR (Becton Dickinson, Mountain View, CA, USA). Again, cells are washed
with chelating medium and are subjected to the fluorescent cell sorting proce-dure. DCs are those that are FITC-cocktail negative and PE-HLA-DR positive
(see Notes 9 and 10).
3.4. Generation of DCs from Human Blood Monocytes
1. PBMCs are prepared from buffy coats from the blood bank, or from freshly drawn
whole blood, by standard Ficoll–Hypaque/Lymphoprep density centrifugation.
PBMC are seeded into 6-well plates at a density of 10 × 106 per well in 3 mL of
culture medium without cytokines (see Note 11).
2. Enrichment of monocytes: After 2 h the nonadherent cells (mostly lymphocytes)
are aspirated and discarded. Warm medium is added back to the adherent frac-tions and the plates are rinsed very gently (see Note 12).
302 Grubeck-Loebenstein, Saurwein-Teissl, and Romani
3. DC differentiation: Monocytes/monocyte-enriched fractions are cultured at a den-sity of about 3–5  × 105/mL in GM-CSF (800 U/mL) + IL-4 (1000 U/mL) for
6–9 d. The cells have then acquired the characteristics of DCs of intermediate
maturity. Cell markers of mature DCs are expressed, but at low density. Further
differentiation/maturation can be achieved by a 2–3-d exposure to either mono-cyte-conditioned medium (see Subheading 2.3.), or, alternatively, a cocktail
(TNFα± IL-1 ± IL-6 + prostaglandin E2) that mimics monocyte-conditioned
medium (54,55) in the continued presence of GM-CSF ± IL-4. When TGF-β is
added together with GM-CSF and IL-4 from the beginning of the cultures, LC
development is most pronounced (20).
4. Feeding of the cultures: The cultures have to be fed every other day, starting on
d2 of culture. To this end 1 mL of culture medium is carefully aspirated from the
wells. To compensate for evaporation it is replaced by 1.5 mL of fresh culture
medium containing 1600 U/mL of GM-CSF and 1000 U/mL of IL-4.
3.5. Generation of DCs from Human Blood CD34+  Progenitors
1. PBMCs are obtained from buffy coat by standard Ficoll–Hypaque/Lymphoprep
density centrifugation, washed twice, and centrifuged through 10% BSA to
remove platelets.
2. Purification of CD34+ cells: 5 × 108 cells are incubated for 5 min at 4°C in 500 µL
of FcR blocking reagent (included in the MiniMACS multisort kit). Five hundred
micoliters of CD34 Multisort microbeads are then added and the cells left to
incubate at 4°C for 60 min. After this incubation period cells are centrifuged and
resuspended in PBS-EDTA. Microbead-labeled cells (108 cells) are loaded onto
each column of the kit in 500 µL and inserted into the magnetic field. Columns
are washed 3× and then removed from the magnetic field. CD34+ cells bound to
beads are cosequently eluted with 1.5 mL buffer and then loaded onto a further
column. The procedure is repeated. Finally, the microbead-labeled CD34+ cell
population is incubated with Multisort Release Reagent for 10 min at 4°C to
release the beads from the cells and loaded onto another column in a magnetic
field. The eluate which contains the unbound CD34+ cells is centrifuged through
10% BSA in PBS for 10 min at 600g and the pellet is resuspended. Purity control
by FACS analysis (see Note 13).
3. Differentiation of DCs from stem cells: CD34+ cells are put into 24-well culture
plates (1–105 cells/well) in culture medium and stimulated every 2 d with
GM-CSF (400 U/mL) and TNF-α (500 U/mL). A greater yield will be achieved
when flt-3 (100 ng/mL), SCF (20 ng/mL), and TGF-β (0.5 ng/mL) are addition-ally present. The latter cytokine works particularly well under serum-free condi-tions, under which it prevents apoptosis (21).
4. Harvesting of DCs: DCs are best harvested after 7–14 d of culture.
3.6. Generation of DCs
from Murine Bone Marrow CD34+  Progenitors
1. Preparation of bone marrow: Remove muscles from the femurs and tibias.
Immerse the bones in a Petri dish with 70% ethanol for 1 min, wash twice in
Dendritic Cells in Old Age 303
Ca2+- and Mg2+-free PBS, and put them into a Petri dish with RPMI-1640. Cut
off both ends (epiphyses) of each bone with scissors. Make a marrow suspension
by flushing out the shafts with a syringe containing about 2 mL of RPMI-1640
per bone. Larger clumps are removed by passage through nylon mesh. Red blood
cells are depleted by hypotonic lysis with NH4Cl. The marrow is then depleted of
mature leukocytes by treatment with antibodies for B cells, T cells, granulocytes,
and complement. For the exact procedure consult Subheading 3.1. (see Note 14).
2. Culturing the cells: Bring the cells to 5–6 × 105/mL in culture medium containing
GM-CSF (final concentration of 200 U/mL) and plate 1–1.5-mL volumes into the
wells of a 24-well plate.
3. Propagation and differentiation of DCs: The wells are washed and fed every 2 d.
Early in the course of the culture (d 2–4) one rinses off the many nonadherent
granulocytes that are developing. On d 4 the rinsing step with RPMI-1640 is
omitted. By d 4 one begins to  see aggregates of growing DCs attached to the
adherent stroma. The aggregates are round, in contrast to macrophage colonies,
which are flattened and dispersed. One can also recognize many cell processes
(“veils”) extending from the periphery of the aggregates, giving them a “hairy”
appearance. By d 6, the wells are usually covered with many aggregates (“balls”)
of proliferating DCs. TNF-α (500 U/mL) when added during the last 2 d of culture
significantly increases the percentage and yield of mature DCs (MHC class II+;
4. Harvesting of DCs: Between d 6 and 8, dislodge the growing aggregates from the
adherent stroma and transfer them to tissue culture dishes. Cells are pooled,
centrifuged, and plated in 10 mL of fresh culture medium containing GM-CSF
per 100 mm Petri dish at a maximal cell number of 10 × 106 per dish. During the
24 h following this transfer, free, nonproliferating, mature DCs are released from
the aggregates. They can be collected by gently swirling the cells from the plate.
Harvesting the mature progeny is best performed after 8 or 9 d of culture (see
Note 15).
3.7. Generation of DCs from Human Bone Marrow
Sufficient amounts of bone marrow for the generation of DCs will only rarely
be available from healthy aged person. Upon suspension culture in GM-CSF +
TNF-α + SCF for 12–14 d an effective yield of about 1.7 × 106 mature DCs per
single milliliter of adult human bone marrow can be obtained. The generation
of DCs from bone marrow CD34+ cells has yet to be optimized with respect to (1)
increasing the low percentage of DCs in the bulk cultures and (2) avoidance of FCS.
4. Notes
4.1. Purification of DCs (LCs) from Murine Skin
1. Cell yields: Depending on the mouse strain (56), 3–5 × 106 primary epidermal
cells, containing 1.4–3% LCs, can be obtained per mouse. Lower yields have to
be expected in aged animals (25).
304 Grubeck-Loebenstein, Saurwein-Teissl, and Romani
2. Reagents: Standard tissue culture reagents including FCS may be obtained from
many different manufacturers with no differences in results. An exception to this
is trypsin, where batch-to-batch variability may be of importance. It might be a
good idea to use one batch for a series of experiments.
3. Pre-enriched (10–15%) fresh LCs are as good as highly enriched LCs (>90%) for
many experimental purposes. For instance, the antigen-processing capacity can
be measured in such populations without interference from the majority popula-tion of keratinocytes (57).
4. According to a recently published method (18,40), DCs can also be obtained from
an organ culture system in which whole ear halves are cultured in one well of a
24-well plate in 2 mL of culture medium for 3 d. DCs migrate into the culture medium
from where they can be harvested. This is a good test to assess the migratory capac-ity of DCs, and the simplest procedure to obtain highly potent mature DCs.
4.2. Purification of DCs (LCs) from Human Skin
5. Cell yields: From 1 cm2 of “young” epidermis, one may expect up to 3  × 106
viable epidermal cells containing 1–3% LCs. This percentage may be consider-ably lower when skin from aged individuals is used.
6. It should be noted that, in contrast to the murine system, there is no simple way to
preenrich human LCs by depletion of a majority of keratinocytes.
7. Organ culture: As in the mouse, DCs can also be readily obtained from the super-natants of skin explants (size 0.5 × 0.5 to 3 × 3cm) cultured for 3 d (58).
4.3. Purification of DCs from Human Blood
8. Cell yield: One obtains 1 × 106 virtually pure DCs after sorting a 500 × 106 start-ing population.
9. The obtained DC population contains functionally mature (CD11c+, CD83+,
CD86+) as well as functionally immature (CD11c-, CD83-, CD86dim) cells.
The functionally immature population develops into mature DCs with typical
dendritic morphology and potent T-cell stimulatory function provided that
monocyte-conditioned medium is present. Otherwise the CD11c- subset dies if
cultured further.
10. An alternative and rather simple method to isolate DCs from fresh blood is based
on the fact that both, functionally mature as well as immature DCs, express sig-nificant levels of CD4. A blood DC isolation kit is available from Miltenyi Biotec
GmbH (Bergisch Gladbach, Germany). DCs are enriched by depletion of T, B,
and natural killer (NK) cells and then enriched using CD4 MicroBeads.
4.4. Generation of DCs from Human Blood Monocytes
11. Cell yields: After 1 wk of culture about 2–5 × 106 DCs can be retrieved from a
starting volume of 40 mL of whole blood (59). When blood from healthy aged
persons is used the yield of DCs is usually as high as 8–10 × 106 per 40 mL of
blood. DC yields are generally lower when PBMCs are used following freezing
and rethawing.
Dendritic Cells in Old Age 305
12. DC precursors are only weakly adherent. Therefore it is important to avoid strong
washing after the initial 2 h of adherence lest the adherent fraction is depleted of
DC progenitors. If done correctly, control cultures of the nonadherent fraction,
which is normally discarded, should not show aggregates.
4.5. Generation of DCs from Human Blood CD34+  Progenitors
13. The percentage of proliferating CD34+ precursors is extremely low (0.06% in the
peripheral blood of healthy adults).
4.6. Generation of DCs
from Murine Bone Marrow CD34+  Progenitors
14. It is essential to proceed quickly with the procedure. Avoid delays and interrup-tions. Initially, do not try to handle more than three mice at one time.
15. Cell yields: Generally, we obtain up to 20–30 × 106 MHC class II-negative cells
from one mouse (two femurs and two tibias), of which most are washed away at
d 2 of culture. By d 7, the yield of DCs is about 5  × 106 per mouse  (60).
Cell yields are dependent on the age of the mice. Five to six weeks is optimal.
DC yields decline substantially at an age of more than 10 wk. It is preferable to
use male mice. One usually obtains 30–50% more cells from male than from
female mice.
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NK Cell Function in Aging 311
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Age-Related Alterations
to Natural Killer Cell Function
Erminia Mariani, Corona Alonso, and Rafael Solana
1. Introduction
Human natural killer (NK) cells represent a heterogeneous lymphoid popu-lation involved in the recognition and lysis of tumor and virally infected cells.
NK cells are defined by the expression of the IgG Fc receptor CD16 (FcγRIIIA)
and/or CD56. NK cells do not rearrange immunoglobulin (Ig) or T-cell recep-tor (TCR) genes. Therefore, neither Ig nor the TCR–CD3 complex is expressed
at the cell surface, except for the zeta (ζ) chain. Both markers, CD16 and CD56,
have been used to analyze age-associated changes in the number of NK cells.
The most characteristic function of NK cells is non-major histocompatibil-ity complex (MHC) restricted cytotoxicity of tumor cell lines and killing of
target cell lines, in particular the erythroleukemia cell line K562. The effect of
aging on NK cytotoxicity has been extensively studied. NK cytotoxicity is
based on the interaction between the NK effector cells and target cells, usually
cell lines that do not express HLA class I antigens on the surface. Several steps
can be differentiated in the cytotoxic assays. The first is conjugate formation,
in which NK cells binds the target cell. In a second phase of NK cell triggering,
activation signals are transmitted from the cell surface inside the cells. As a
result perforin granules are reoriented to the effector–target contact site, and
they are finally released and the target cell lysed.
The K562 cell line is the target cell most commonly used for measuring
lysis by resting NK cells from elderly people, whereas the NK-resistant Daudi
cell line is used for testing the cytotoxic capacity of activated NK cells. Both
Daudi or P815, a mastocytoma murine cell line that is also resistant to lysis by
312 Mariani, Alonso, and Solana
resting NK cells, are frequently used for testing the antibody-dependent cell
cytotoxicity (ADCC, Fc receptor-mediated lysis by NK cells).
The results on the different phenotypic and functional analysis of NK cells
have demonstrated age-associated changes both in the number and function of
NK cells. Thus, although there is a general consensus that NK cytotoxicity of
peripheral blood lymphocytes is well preserved not only in healthy elderly
people but also in centenarians, a significant expansion in the number of NK
cells is also found in healthy aging (1). This indicates that NK cell killing is
impaired when it is considered on a per cell basis, as it is shown when single-cell cytotoxic assays are performed (2,3). The ability of interleukin-2 (IL-2)-activated NK cells to lyse the normally NK-resistant Daudi cell line is also
significantly decreased in the elderly, when compared to young people  (4).
However, ADCC- and LAK-mediated killing remain comparable between
young and elderly subjects. The decreased NK cytotoxic capacity found in the
elderly is associated with defective signal transduction (5). On the other hand,
perforins are distributed in the cytoplasm of almost all the NK cells from the
elderly, as in NK cells from the young, the ability to utilize perforin in the
generation of cytolytic activity against tumor target cells is maintained in NK
cells from the elderly (6).
Recently Ogata et al. (7) showed that low NK cell function relates to the
development of severe infections in elderly subjects and that well preserved
NK cell cytotoxicity is relevant for survival. These findings strengthen the
interest in studying NK cytotoxicity in aging.
In the following sections we introduce the techniques and experimental
protocols usually employed for the analysis of NK cell phenotype and func-tion in the elderly. The standard immunofluorescence and cytotoxicity assay
as well as the techniques for determining NK signal transduction are
2. Materials
2.1. Analysis of NK Cell Phenotype and Perforin Content
by Immunofluorescence
1. Lymphocyte separation medium (LSM), density 1.077 g/mL (e.g., Histopaque-1077, Sigma, St. Louis, MO, USA).
2. Culture medium: RPMI 1640 with  N-2-hydroxyethylpiperazine-N’-2-ethane-sulfonic acid (HEPES) buffer supplemented with 10% fetal calf serum (FCS),
antibiotics (penicillin: 100 U/mL plus streptomycin 100 µg/mL or alternatively
gentamicin 200 µg/mL) and 2–4 mM L-glutamine (all reagents from Gibco Life,
Paisley, UK).
3. Phosphate-buffered saline (PBS) (Gibco Life).
4. Trypan blue solution: 0.2% w/v in PBS/3 mM sodium azide (Sigma).
NK Cell Function in Aging 313
5. Petri dishes (Corning Costar Europe, Bad Loevedorp, The Netherlands).
6. Nylon wool (Robbins, Sunnyvale, CA, USA).
7. Monoclonal antibodies (MAbs): anti-CD16 (3G8, Immunotech, Marseille Cedex,
France; Leu11, Becton Dickinson, San Josè, CA, USA), anti-CD56 (NKH-1,
Coulter Corporation, Miami, FL, USA; Leu19 Becton Dickinson, San Josè, CA,
USA), anti-CD94 (HP3B1, Coulter Corporation, Miami, FL, USA). These MAbs
are available as PE or FITC labeled. Double fluorescence with anti-CD3 (Anti-Leu 4, Becton Dickinson, San Josè, CA, USA) is recommended.
8. For intracellular labeling of perforins the following additional material is
a. 2% Paraformaldehyde (Fluka Chemie AG, Buchs, Switzerland) solution in
b. 0.2% Tween-20 (Sigma, St. Louis, MO, USA) solution in PBS.
c. PBS with 2% FCS and 0.1% sodium azide (Sigma, St. Louis, MO, USA)
d. MAb anti-human perforin and Ig isotype control (Pharmingen, San Diego,
e. Fluorescein isothiocyanate (FITC)- or phycoerythrin (PE)-conjugated
goat anti-mouse Ig (GAM-FITC or GAM-PE) (Pharmingen, San Diego,
f. Normal mouse serum (Dako, Glostrup, Denmark).
9. Flow cytometer (FacsCount, Becton Dickinson, San Josè, CA, USA).
2.2. NK Cell Enrichment and Purification
1. LSM, density 1.077 g/mL (Histopaque–1077, Sigma, St. Louis, MO, USA).
2. Culture medium prepared as indicated in Subheading 2.1.
3. PBS (Gibco Life, Paisley, UK).
4. Laminar flow hoods.
5. Petri dishes (Corning Costar Europe, Bad Loevedorp, The Netherlands).
6. Nylon wool (Robbins, Sunnyvale, CA, USA).
7. MAbs: anti-CD3 (Anti-Leu 4, Becton-Dickinson, San Josè, CA, USA) anti-CD19
(J4.119,  Immunotech, Marseille Cedex, France; Leu12 Becton Dickinson, San
Josè, CA, USA), and anti-CD16 (3G8, Immunotech, Marseille Cedex, France;
Leu11, Becton Dickinson, San Josè, CA, USA).
8. GAM coupled magnetic beads (Dynal AS, Skoyen, Norway).
9. Magnetic particle concentrator (MPC1, Dynal AS, Skoyen, Norway).
10. MACS cell separation reagents and equipment (Miltenyi Biotec GmbH, Bergisch
Gladbach, Germany) can also be used to purify NK cells.
a. Separation reagents: the NK isolation kit consists of: reagent A, a cocktail of
hapten-conjugated monoclonal CD3, CD4, CD19, CD33 antibodies; reagent
B, colloidal superparamagnetic MACS microbeads conjugated to an anti-hap-ten antibody and depletion columns.
b. Equipment: VarioMACS magnetic cell separator.
314 Mariani, Alonso, and Solana
2.3. Cytotoxic Assays: NK Cytotoxicity, Lymphokine-Activated
Killing, ADCC, and Redirected Lysis
1. 51Sodium chromate (51Cr) solution in normal saline with a specific activity of
400–1200 Ci/g (14.8–44.4 TBq) (NEN, Bad Homburg, Germany)
2. Triton X-100 (Sigma, St. Louis, MO, USA).
3. 96-Well round- or V-bottom microtiter plates (Corning Costar Europe, Bad
Loevedorp, The Netherlands).
4. γ-Counter (Ultrogamma LKB, Upsala, Sweden, or Gamma counter, Beckman,
Palo Alto, CA, USA).
5. Cultured target cells: The most commonly used target cells for measuring NK
lysis are K562 cells, an HLA negative erythroleukemia derived cell line. The
HLA class I negative EBV transformed cells C1R or 721.221 are also extensively
used at present as NK target cells. Other cell lines such as Daudi (derived of a
Burkitt lymphoma) or P815 (a murine mastocytoma), that are resistant to NK
killing, are used to determine LAK activity, ADCC, or redirected lysis.
2.4. Signal Transduction in Aging
1. Inositol-free RPMI 1640 (Gibco Life, Paisley, UK) supplemented with 10% FCS.
2. myo-[3H]Inositol (Amersham International, Buckinghamshire, UK) (specific
activity 3.15 Tbq/mmol).
3. Susceptible target cells: K562.
4. Nonsusceptible target cells: Daudi as negative control.
5. MAb anti-CD16 and irrelevant MAb (Becton Dickinson, San Josè, CA, USA)
that will be used as negative control.
6. Extraction solution: methanol/chloroform/HCl 37% (80:160:1 v/v) (Fluka
Chemie AG, Buchs, Switzerland).
7. Washing solution: 0.7 mL of methanol/chloroform/1  N HCl (235:15:245 v/v)
(Fluka Chemie AG, Buchs, Switzerland).
8. 1 M KOH (Fluka Chemie AG, Buchs, Switzerland).
9. Bidistilled water.
10. Amprep ion-exchange minicolumns (Amersham International, Buckinghamshire,
11. Elution buffers: 0.1 M KHCO3, 0.17 M KHCO3, 0.25 M KHCO3 (Fluka Chemie
AG, Buchs, Switzerland).
12. Solubilizing solution: chloroform/methanol (2:1) (Fluka Chemie AG, Buchs,
13. 1% K oxalate (Sigma, St. Louis, MO, USA).
14. Thin-layer chromatography (TLC) plates.
15. Developing solution: chloroform/methanol/water/saturated ammonia (45:35:8:2
v/v) (Fluka Chemie AG, Buchs, Switzerland).
16. Extraction solution: 0.6 N HCl/methanol (60:40 v/v) (Fluka Chemie AG, Buchs,
17. Enhancer (NEN, Bad Homburg, Germany).
NK Cell Function in Aging 315
18. Iodine (Sigma, St. Louis, MO, USA).
19. Pico-Fluor 40 scintillation cocktail (Packard, Meriden, CT, USA).
20. Beta-counter (Beckman, Palo Alto, CA, USA).
3. Methods
3.1. Flow Cytometric Analysis of NK Cells
1. Isolate peripheral blood mononuclear cells (PBMCs) by centrifugation (900g/
20 min) of peripheral blood, diluted 1:1 with PBS, over the lymphocyte separa-tion medium. Remove PBMCs from the interface, dilute with PBS, and centri-fuge at 400g/10 min. Decant and repeat wash steps two additional times.
2. Perform white cell count. Dilute the cell suspension to a final concentration of
5 × 106 PBMCs/mL in culture medium.
3. For direct immunofluorescence 2 × 105 PBMC are incubated with each labeled
MAb in the dark in V-bottom plates for 30 min at 4°C, washed twice with PBS
plus 2% FCS and 0.1% sodium azide and resuspended in 1% paraformaldehyde
for the analysis. Mixtures of two PE or FITC differently labeled MAbs (i.e., FITC
anti-CD3 in combination to PE anti-CD56) can be incubated simultaneously with
the same cells for double fluorescence. Isotype fluorescence labeled Ig control
MAbs should be used as negative controls.
4. For indirect immunofluorescence (IIF) incubate 2 × 105 PBMCs as for the direct
method but using unlabeled antibodies. After washing with PBS, incubate cells
again with FITC- or PE-labeled goat anti-mouse Ig for 30 min at 4°C. After wash-ing fix the cells in 1% paraformaldehyde for the analysis. For negative control,
incubate cells only with the secondary labeled antibody or with a primary anti-body of the same isotype but with irrelevant specificity followed by the second-ary antibody (Note 1). When a double immunofluorescence technique is used,
perform an incubation with an optimal dilution (1:10 usually) of normal mouse
serum after the secondary antibody incubation, to reduce nonspecific binding
before the last incubation with a directly labeled MAb. The combination of IIF
with labeled anti-CD56 MAbs is useful to study the presence of other differentia-tion or activation antigens on NK cells (Note 2).
5. Techniques of IIF on permeabilized NK cells can be used to evaluate the perforin
content of NK cells. Resuspend 106 PBMCs in 1 mL of cold 2% paraformalde-hyde solution in PBS, incubate for 1 h at 4°C, and centrifuge for 10 min at 250g at
4°C. Resuspend the fixed cells in 1 mL of 0.2% Tween-20 in PBS at room tem-perature and incubate for 15 min at 37°C. Add 3 mL of PBS–FCS–Azide and
centrifuge the suspension for 10 min at 250g at room temperature. Incubate the
cell pellet with an optimal dilution of an anti-human perforin MAb for 30 min at
4°C; develop positive cells using a 1:20 diluted GAM Ig conjugated with FITC for
30 min at 4°C. The cell pellet is then washed with PBS–FCS–Azide and analyzed.
6. Flow cytometry: The simultaneous measurement of “forward scatter” that mea-sures cell size and “side scatter” that indicates heterogeneity of the cell structure
(“granularity or complexity”) allows the identification and gating of the different
316 Mariani, Alonso, and Solana
subpopulations of white blood cells (Note 3). Optimal gating for the identifica-tion of lymphocyte populations requires polygonal computer-generated windows
(Note 4). FITC green fluorescence or PE red fluorescence can be simultaneously
analyzed. For each sample, 104 gated cells are usually analyzed.
3.2. Cytotoxic Assays for NK Cells: Preparation of Effector Cells
Different sources of effector cells can be used as effectors of NK cell cyto-toxicity. In age-related studies plastic- and nylon-wool-depleted PBMCs are
commonly used to analyze NK killing of K562 target cells whereas IL-2-acti-vated cells are used to study LAK killing of the NK-resistant Daudi cell line.
1. PBMC enriched by depletion of plastic and NW adherent cells. Incubate PBMCs
at 5  × 106 PBMCs/mL in a Petri dish for 60–90 min at 37°C in 5% CO2, for
monocyte depletion. Recover nonadherent cells, wash twice with PBS, and dilute
in 2 mL of medium. The cells are then incubated in prewashed nylon wool for
45–60 min at 37°C in 5% CO2, for B cell depletion. Recover the nonadherent
(T/NK cells) by washing the column with 20 mL of culture medium and resus-pend. This cell population contains 20–30% of CD56+ NK cells.
2. Activation of NK cells with IL-2. Culture the cells obtained as indicated in
step 1 in 5% CO2 at 37°C for 7–10 d in the presence of 500 U/mL of IL-2,
without further activation requirements. This population is considered LAK
cells and is constituted by a mixture of activated NK and T-cells able to lyse
NK-resistant cell lines.
3. Purification of NK cells by immunomagnetic separation. Incubate the nonadherent
cells (T/NK cells) obtained as indicated in step 1 with anti-CD3 MAb for 30 min
at 4°C and add 0.7–0.9 × 108 of GAM-coupled magnetic beads. Incubate for 30 min
at 4°C under gentle shaking for T-cell depletion. Remove the cells rosetting with
the beads by using a magnetic particle concentrator (MPC1) and collect the super-natant with the CD3-negative cells. Alternatively, whole PBMCs (5 × 106/mL) can
be directly incubated with CD3, CD4, CD19, and CD14 MAb for 30 min at 4°C,
and then with GAM-beads for 45 min at 4°C. Cells and beads should be gently
mixed throughout. After the incubation with beads, use a magnet to separate beads
with attached CD3/CD4 T-cells, CD14 monocytes, and CD19 B cells from NK
cells. Centrifuge and wash with complete medium the recovered free cells. This
cell population should be 75–95% CD56+ and CD16+, <10% CD3+ and <5% CD14+,
and CD19+ as routinely established by flow cytometry.
4. Isolation with the MACS system. Resuspend 107 PBMCs in 80 µL of buffer (PBS
with 2 mM EDTA and 0.5% bovine serum albumin [BSA]), add 20 µL of reagent
A, and incubate for 15 min at 6°–12°C. Wash the cell suspension carefully, resus-pend again with 80 µL of buffer, add 20 µL of reagent B, and incubate as before.
Wash cell suspension and resuspend the pellet with 500 µL of buffer. Apply cell
suspension to a prefilled depletion column placed in the magnetic field of the
VarioMACS and collect effluent cells representing the purified NK cell fraction.
NK Cell Function in Aging 317
3.3. Cytotoxic Assays for NK Cells: Preparation of Target Cells
Several cell lines are used as target cells for measuring NK lysis. The most
commonly used are K562 cells, as well as C1R or 721.221. Daudi cell line, an
NK-resistant cell line, is used to study cytotoxicity by IL-2-activated NK cells.
Furthermore, P815, an NK-resistant mastocytoma murine cell line which
express high concentrations of Fc receptors, can be used to analyze redirected
lysis, for example, lysis of the target in the presence of antibodies against NK
triggering structures. Both Daudi and P815 cell lines can be used to analyze
ADCC by using NK cells and IgG antibodies specific for the target cells. Grow
these cell lines in RPMI 1640 + 10% FCS, 2 mM glutamine, 100 U/mL of
penicillin, and 10 µg/mL of streptomycin and use them during the logarithmic
growth phase (Note 5).
1. Radioactive labeling of target cells with 51sodium chromate: Use 51Cr solution in
normal saline to label target cells and no later than 15 d after the reference day.
For target cell labeling, wash 1–2  × 106 once with fresh medium, remove the
supernatant, and incubate pellet with 100 µCi 51Cr in differing volumes, accord-ing to the decay table (Note 6). Incubate the cells at 37°C in 5% CO2 for 1 h with
occasional shaking at 10–15-min intervals. During following procedures, keep
the target cells on ice to reduce spontaneous isotope release. Wash the target tumor
cells 3× at 4°C in cold RPMI 1640 + 10% FCS and resuspend in medium at a
concentration of 5 × 104 cells/mL (Note 7).
2. NK cell cytotoxicity assays: The effector/target cell ratio will depend on the
nature of the effector population and its level of cytotoxic activity. A starting
effector/target ratio of 100:1 or 50:1 will be required for assaying freshly isolated
NK cells, whereas for purified NK cells or activated LAK this may be at most
20:1 or 10:1. In addition to NK and LAK assays ADCC can be determined when
an NK-resistant cell line is used and anti-target antibodies are added to the assay.
Redirected lysis is measured when P815 are used as targets and MAbs against
NK triggering structures (i.e., CD16, CD69) are added to the assay. Perform the
cytotoxicity assays in V-bottom 96-well microtiter plates with a final volume of
200 µL. Seed each effector/target cell ratio (E/T) in triplicate. For spontaneous
release control, samples of target cells are resuspended in medium alone. Maxi-mum or total release of 51Cr from the target cells is obtained by mixing 5 × 103
labeled target cells with 100 µL of 2% Triton X-100. Use at least six replicate
wells to evaluate spontaneous and maximum release. Seed serial dilutions of
mononuclear effector in 100 µL of culture medium and 100 µLof 51Cr-labeled
tumor target cells with an E/T ranging from 100:1 to 12:1 (Note 8). Centrifuge
the plates at 4°C for 3 min and incubate at 37°C in 5% CO2 for 4 h. Harvest and
transfer to a tube 100 µL of culture supernatant at the end of the incubation time.
Determine 51Cr release in a γ-counter. Calculate specific 51Cr release according
to the formula:
318 Mariani, Alonso, and Solana
[(test release — spontaneous release) /
(maximum release – spontaneous release)] × 100.
3.4. Analysis of Phosphoinositide Turnover
in NK Cells from Elderly People
1. Labeling of inositol phospholipids: Resuspend purified NK lymphocytes at a con-centration of 1 × 106 cells in a final volume of 1 mL of inositol-free RPMI 1640
supplemented with 10% heat-inactivated and dialyzed FCS. NK lymphocytes are
metabolically labeled with  myo-[3H] inositol (5  µCi/106 cells/mL) for 18 h at
37°C in 5% CO2. Wash the labeled cells 2× for 10 min at 300g using cold inosi-tol-free RPMI 1640 with 10% FCS and resuspend the pellet in the same medium
at 2.5 × 106 cells/mL. Stimulate the purified NK cell samples (2.5 × 105/100 µL)
in Eppendorf tubes with either a similar number of target cells (E/T ratio 1:1) or
with an appropriate amount of MAb for various time intervals (from 1 to 30 min)
at 37°C in a water bath. Incubate control samples for the same time both in the
absence of stimuli and with nonsusceptible targets and/or irrelevant MAbs. Stop
incubation by adding methanol/chloroform/HCl 37% (80:160:1 v/v), keep the
samples at 4°C, and extract phospholipids by vortex-mixing the cells in the above
extraction solution. After centrifugation (100,000g for 5 min) an upper aqueous
phase containing inositol phosphates and a lower chloroform phase containing
phospholipids are obtained. Collect the upper phase, dry it in vacuo, and store at
–80°C until further use. Wash the lower phase with 0.7 mL of methanol/chloro-form/1 N HCl (235:15:245 v/v) by vigorous mixing. After centrifugation
(100,000g for 5 min) collect the new lower phase and dry it in vacuo.
2. Analysis of inositol phosphate (IP) fractions (IP, IP2, IP3). To analyze inositol
phosphates, treat the dried samples of the inorganic upper phase with 1 M KOH,
resuspend these samples in 4 mL of bidistilled water, and load them onto Amprep
ion-exchange minicolumns. IP and IP2 are eluted together from the column with
5 mL of 0.1 M KHCO3, while the remaining IP3 is eluted using a gradient based on
0.17 M of KHCO3 and 0.25 M KHCO3 buffers at a flow rate of 1 mL/min. Collect
samples in three fractions of 5 mL and measure 3H-labeled inositol phosphates by
liquid scintillation counting using a β-counter. Compare the eluted peaks to reten-tion times for standards prepared from [3H]PI, [3H]PIP, and [3H]PIP2. The scintil-lation counting is set to obtain a counting error lower than 5%.
3. Analysis of phosphatidylinositol phosphate fractions (PIP, PIP2, PIP3). Solubi-lize the lower phase containing phosphatidylinositols in chloroform/methanol
(2:1) and spot it onto 1% K oxalate-sprayed TLC plates to separate [3H] labeled
phosphatidylinositols. Develop TLC plates with chloroform/methanol/water/satu-rated ammonia (45:35:8:2 v/v), spray with Enhancer, and fluorograph at –80°C.
Autoradiograph TLC plates before exposure to iodine to visualize internal lipid
standards obtained from Sigma (St. Louis, MO, USA). Scrape off the single
“spot” of PI, PIP, and PIP2; extract with 1.5 mL of 0.6 N HCl/methanol (60:40 v/v)
for 48 h with gentle stirring, and count with a liquid scintillation counter using
NK Cell Function in Aging 319
9mL of Pico-Fluor 40 scintillation cocktail. The scintillation counting is set to
obtain a counting error lower than 5%.
4. Notes
1. The use of normal mouse serum is also recommended to reduce nonspecific bind-ing when an indirect immunofluorescence reaction is combined with a direct
immunofluorescence reaction on the same cell sample.
2. To study the expression of differentiation markers on NK cells the use of double
fluorescence, using PE labeled anti-CD56 or anti CD16 MAbs, is recommended.
3. Cytofluorimeters are instruments that measure cell size and phenotype by the
presence of bound fluorochrome-labeled antibodies and are widely used for
analysis of cells labeled as described earlier. A single-cell suspension labeled
with FITC- or PE-conjugated antibodies is forced through the nozzle of the
machine under pressure. The light scattered and reflected by the cells and fluo-rescence emitted by excited fluorochromes bound to the cell membrane are
detected, analyzed, processed, and stored by a computer. Forward light scatter or
“forward scatter” (FS) measures cell size while perpendicular light scatter or “side
scatter” (SS) indicates heterogeneity of the cell structure (“granularity or com-plexity”).
4. Most NK cells show large granular lymphocyte morphology and therefore pro-duce forward and side scatter signals higher than other lymphocyte subsets. Thus
we do not have to use a too-restricted lymphocyte gate. This point should be
borne in mind when setting gates for analyzing NK cells in whole PBMCs.
5. In the lysis assays the target cells should be collected in the logarithmic phase of
growing, as 51Cr labeling is poorer when quiescent cells are used.
6. The optimal labeling is obtained when 51Cr is added to a target cell pellet as dried
as possible to avoid dilution by culture medium.
7. One cpm per target cell can be considered an adequate total labeling. The sponta-neous release should be as low as possible, but is considered optimal if it is less
than 10% of the maximum release although about 30% can also be exceptionally
8. Assaying the lytic activity of peripheral blood lymphocytes with the standard test
may present some limitations when it is possible to collect only minimal blood
samples, as it occurs in elderly people. In these situations, the possibility of using
a low number of effector cells would greatly facilitate the performance of cyto-toxicity assays. In this microcytotoxicity assay, use a 10-fold lower number of
cytolytic effector and target cells with a maintained E/T ratio (3).
R. S. and C. A. are supported by grants from Spanish Ministry of Health (FIS95/
1242, FIS 98/1052) and Junta de Andalucía (Spain). E. M. is supported by grants
from MURST (60% fund) and University of Bologna. This work was carried out
under the aegis of the European Union Concerted Action on the Molecular Biology
of Immunosenescence (EUCAMBIS; Biomed I contract CT94–1209).
320 Mariani, Alonso, and Solana
1. Borrego, F., Alonso, C., Galiani, M., Carracedo, J., Ramirez, R., Ostos, B., Pena, J.,
and Solana, R. (1999) NK phenotypic markers and IL2 response in NK cells from
elderly people. Exp. Gerontol. 34, 253–265.
2. Mariani, E., Roda, P., Mariani, A. R., Vitale, M., Degrassi, A., Papa, S., and Facchini,
A. (1990) Age-associated changes in CD8+ and CD16+ cell reactivity: clonal analy-sis. Clin. Exp. Immunol. 81, 479–484.
3. Mariani, E., Monaco, M. C., Sgobbi, S., de Zwart, J. F., Mariani, A. R., and Facchini,
A. (1994) Standardization of a micro-cytotoxicity assay for human natural killer
cell lytic activity. J. Immunol. Methods 172, 173–178.
4. Kutza, J. and Murasko, D. M. (1996) Age-associated decline in IL-2 and IL-12
induction of LAK cell activity of human PBMC samples. Mech. Ageing Dev. 90,
5. Mariani, E., Mariani, A. R., Meneghetti, A., Tarozzi, A., Cocco, L., and Facchini,
A. (1998) Age-dependent decreases of NK cell phosphoinositide turnover during
spontaneous but not Fc mediated cytotlytic activity. Int. Immunol. 10, 981–989.
6. Mariani, E., Sgobbi, S., Meneghetti, A., Tadolini, M., Tarozzi, A., Sinoppi, M.,
Cattini, L., and Facchini, A. (1996) Perforins in human cytolytic cells: the effect of
age. Mech. Ageing Dev. 92, 195–209.
7. Ogata, K., Yokose, N., Tamura, H., An, E., Nakamura, K., Dan, K., and Nomura, T.
(1997) Natural killer cells in the late decades of human life.  Clin. Immunol.
Immunopathol. 84, 269–275.
Immunogenetics and Life-Span 321
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Immunogenetics and Life-Span
Derek Middleton, Martin D. Curran, and Fionnuala Williams
1. Introduction
The major histocompatibility complex (MHC) is located in the region 9p21–
6pter on the short arm of chromosome 6 and encompasses approx 4000
kilobases of genomic DNA. Contained within this complex are numerous genes
with immune-related functions: notably the class I and class II human leuko-cyte antigens (HLA), tumor necrosis factor A and B, the complement genes,
and genes that orchestrate the transport (TAP) and processing (LMP) of anti-gens for presentation. The HLA class I (HLA-A, HLA-B, HLA-C) and HLA
class II (HLA-DR, HLA-DQ, HLA-DP) cell surface glycoproteins present
antigenic peptides to CD8+ and CD4+ T cells, respectively, and play a central
role in mediating the immune response. The HLA class I and class II genes
display extreme degrees of polymorphism, making the MHC region the most
polymorphic and densely populated area of the human genome. It is now well
established that certain HLA specificities are strongly associated with numer-ous diseases, especially those with an autoimmune dimension, and confer
resistance/susceptibility to certain infectious diseases. The possibility that HLA
identity may have a genetic role in longevity in humans has long been sus-pected, spurred on by the importance of the HLA antigens in the immune response
and data from studies in mice indicating that genes in the MHC region are associ-ated with a significant effect on life-span. The results to date from these
studies are confusing and contradictory, with no consistent association found
as yet (1). Critical examination of the studies highlights the use of serology,
known to lack the resolution capable of defining many HLA specificities, and
the small number of cases >90 yr included in the studies. The advantages
322 Middleton, Curran, and Williams
(greater specificity and accuracy) of using molecular methods instead of sero-logical typing to define the HLA system have been well shown in transplanta-tion and disease association (2,3). Such approaches should allow future studies,
examining whether certain HLA alleles influence longevity, to reach a mean-ingful conclusion.
Many molecular methods are available to define the HLA alleles. Described
in this chapter is the sequence-specific oligonucleotide probe (SSOP) method.
This method is also directly applicable to defining other polymorphic loci
within the MHC or elsewhere in the genome. The basis of this method is the
specific amplification of the HLA locus by the polymerase chain reaction
(PCR) and the subsequent probing of this product by SSOPs. Most of the vast
polymorphism of the HLA system results from conversion events whereby
small nucleotide sections of one allele (usually no more than 100 bases long)
are transferred to another allele. Thus many of the sequences tend to be shared
by alleles and are not allele specific. Therefore probes are used that are sequence
specific. To differentiate the alleles, a battery of probes is required and it is the
pattern of reactivity of these probes that distinguishes the HLA alleles.
The detection system used in this laboratory consists of labeling the probes
with digoxigenin (DIG) and detecting the hybridization of these probes to a
complementary sequence present in the PCR-amplified HLA allele of an indi-vidual by adding an anti-DIG antibody conjugated with alkaline phosphatase
(ALP). The ALP then uses disodium 3-(4-methoxyspiro {1, 2-dioxetane –3, 2′-5′-chloro tricyclo [] decan}-4-yl) phenyl phosphate (CSPD) as its chemi-luminescent substrate and the light emitted is detected by autoradiography.
To define all alleles at any specific locus at the same time would require a
large number of probes as, although each allele group has a specific probe
pattern, the combined probe pattern of two alleles present in a heterozygous
individual can be identical to the combined probe pattern of another heterozy-gous individual with two different alleles. In addition the system would con-stantly need up-dating to take account of newly discovered alleles. In this
laboratory we use a two-tier SSOP system. The first level of resolution is
equivalent to very good serology, that is, the allele group is defined, for
example, HLA-A*02. Thereafter, depending on the initial type, a second PCR
specific for a group of alleles is performed and a further set of probes used to
give definition to the allele level. Thus the number of probes required is kept to
a minimum and, except for exceptional circumstances, only the high-resolu-tion system needs alteration to take account of new alleles. The primers used
for each locus are listed in Table 1. The primers for HLA-A and -B loci give a
locus-specific product covering exons 2 and 3 and the primer for HLA-DR
gives a product from exon 2. This product is not specific for the HLA-DRB1
locus and amplifies alleles of other HLA-DR loci (e.g., HLA-DRB3 locus).
Immunogenetics and Life-Span 323
Thus it is necessary to include a further amplification for alleles of HLA-DRB1*03, -DRB1*11, -DRB1*13, and -DRB1*14. This is referred to as the
HLA-DRB3/11/6 group. The reason for two 3′ end primers for HLA-B is
because HLA-B*7301 differs in intron 3 from all other known alleles at this
locus and the extra primer is required to amplify this allele.
Table 1
HLA-A, -B, -C, -DR Primers Used for SSOP Typing
Sequence Band
Primers 5′ → 3′ size
HLA-A 94 (Intron 1) → 116
13 (Intron 3) → 274 (exon 3)
HLA-B 36 (Intron 1) → 57
68 (Intron 3) → 37
68 (Intron 3) → 37
HLA-B27 5 BINI-57M See above 970
3 BIN3-37M See above
42 (Intron 1) → 61 937
35 (Intron 3) → 12
HLA-DRB (Intron 1) 15 (Exon 2) → 24
279 (Exon 2) → 260
HLA-DRB 17 (Exon 2) → 38
3/11/6 group 3/11/6 GF GTTTCTTGGAGTACTCTACGTC 263
279 (Exon 2) → 260
*A ( ) in primer indicates that at this position two nucleotides are inserted when the primer is being
made. The primer is referred to as being degenerate.
324 Middleton, Curran, and Williams
In this laboratory we do not use tetramethylammonium chloride (TMAC)
owing to its toxic properties and the fact that in our experience it does not
necessarily mean the use of one wash temperature. It would appear that to fol-low this practice a laboratory would require a large number of water baths. In
this laboratory one individual normally performs 12 hybridizations at the same
time. Thus if a laboratory is defining alleles at three loci (HLA-A, -B, -DR),
probes can be selected for use at the same time according to their wash tem-perature. This eliminates the requirement for a large number of water baths.
In the methods described each probe is hybridized to two different mem-branes in the same hybridization bottle and the reagents are prepared accord-ingly. The SSOP method is thus very suitable for typing large numbers of
samples — we test at the same time 192 samples (96 on each membrane) which
includes controls. However, the volume of reagents can be scaled down and if
a laboratory is not performing tests on large numbers of samples, only one lot
of membranes need be hybridized.
2. Materials
1. Buffer 1 (4×): 0.4 M Maleic acid, 0.6 M NaCl, pH 7.5. Add 300 mL of 4 M NaCl
and 400 mL of 2 M maleic acid followed by 200 mL of 4 M NaOH to approx 800 mL
of double-distilled H2O (ddH2O). Add 27 g of NaOH pellets. (Note: A white
precipitate forms when all reagents are added — this will disappear as the pH
approaches 7.0.) Cool to room temperature and adjust pH to 7.5 by adding 4 M
NaOH by drops. Adjust volume to 2 L with ddH2O and sterilize by autoclaving.
2. Buffer 2: 2% Blocking reagent in buffer 1. Combine 768 mL of 5% blocking
reagent (in buffer 1), 288 mL of 4× buffer 1, and 864 mL of ddH2O. Leave 5%
blocking reagent at room temperature for 10 min before use.
3. Buffer 3: 0.1 M Tris-HC1, 0.1 M NaCl, 0.05 M MgCl2, pH 9.5. Add approx 1400 mL
of ddH2O to 200 mL of 1 M Tris-HCl, pH 9.5, and 50 mL of 4 M NaCl. Add
100 mL of filter sterilized 1 M MgCl2 and mix. Adjust pH to 9.5 and make up to
2 L with ddH2O. Do not autoclave as precipitates tend to form. Store at room
temperature for up to 1 wk.
4. Buffer hybridization: 192 mL of 2% blocking reagent, 144 mL of 6× saline
sodium phosphate EDTA (SSPE), 48 mL of 5× Denhardts, 48 mL of 0.1%
N-laurylsarcosine, 0.96 mL of 0.02% sodium dodecyl sulfate (SDS) and make up
to 480 mL with ddH2O.
5. Buffer washing: 0.3% Tween-20 in buffer 1. Add 14.4 mL of Tween-20 to 1200 mL
of 4× buffer 1 and make up to 4800 mL by adding ddH2O.
6. Blocking reagent: 5% in buffer 1 (Boehringer, Lewes, England, cat. no. 1096176).
Prepare 2 L of 1× buffer 1 by combining 500 mL of 4× buffer 1 with 1500 mL of
ddH2O. Add 100 g of blocking reagent in parts, with vigorous mixing using a
magnetic stirrer, to approx 1600 mL of 1× buffer 1. As blocking reagent is sup-plied in 50 g tubs, there is no need to weigh out. Heat to 65°C until blocking
Immunogenetics and Life-Span 325
reagent is dissolved. Allow to cool to room temperature and make up to 2 L with
buffer 1. Sterlize by autoclaving and store at 4°C.
7. Cresol red: 10 mg/mL, sodium salt (Sigma, St. Louis, MO, USA, cat. no. C9877).
Add 200 mg to some ddH2O taken from measured 20 mL dH2O in a sterile bottle.
Resuspend in remaining volume. Filter sterilize and dispense into 1-mL aliquots
and freeze at –20°C.
8. CSPD (Boehringer, cat. no. 1655884). Vortex-mix and centrifuge CSPD in a
microcentrifuge for 1 min before use. Dilute CSPD stock solution (25 mM,
.6 mg/mL) 1:100 in buffer 3.
9. Denhardts (50×): 1% Polyvinyl pyrrolidone (PVP), 1% Ficoll, 1% bovine serum
albumin (BSA). Prepare 200 mL of 2% PVP and 2% Ficoll by adding 4 g of each
to 180 mL ddH2O. (Prepare this solution in a fume cupboard. PVP is harmful if
inhaled.) Dissolve with gentle mixing and make up to 200 mL with ddH2O. Ster-ilize by autoclaving and cool to room temperature. Add 4 g of BSA to 200 mL of
the above solution slowly with gentle mixing. When the BSA has dissolved make
up to 400 mL with ddH2O, mixing well. Filter solution through 0.45 µm filter and
aliquot. Do not autoclave. Store at –20°C. Leave to thaw at +4°C the evening
before it is to be used.
10. Anti-DIG–ALP conjugate (Boehringer, cat. no. 1093274). Just prior to use
remove anti-DIG–ALP stock conjugate (0.75 U/µL) from the refrigerator, vor-tex-mix for 15 s and centrifuge for 1 min in a microcentrifuge. Make a 1:10,000
dilution of the conjugate in buffer 2 (i.e., 192 µL of anti-DIG–ALP conjugate in
1920 mL of buffer 2).
11. 0.5  M EDTA, pH 8.0. Add 186.1 g of EDTA Na22H2O in parts to 800 mL of
ddH2O. Adjust the pH to 8.0 using 4 M NaOH. Make up to 1 L with ddH2O and
sterilize by autoclaving.
12. Ethidium bromide (10 mg/mL, Sigma, cat. no. E-1510).
13. Gel loading buffer (GLB): Add 8 g of sucrose (slowly) to 10 mL of ddH2O and
mix by inversion until dissolved. Then add 1 mL of 1 M Tris, pH 7.6; 2 mL of 0.5 M
EDTA; 1 mL of 10% SDS; 0.02 g of cresol red. Make up to 20 mL with ddH2O.
Do not autoclave.
14. ddH2O. Double-distilled H2O or equivalent. Note that ddH2O used to set up PCR
is of ultra-high-purity quality.
15. MgCl2. Supplied with Ta q enzyme.
16. dNTPs (Pharmacia Biotech, St. Albans, England, cat. no. 27-2094).
17. NH4 buffer. Supplied with Ta q enzyme.
18. N-Laurolysarcosine (1%). Barrier face mask should be worn when weighing
N-laurylsarcosine. Dissolve 10 g of  N-lauryl sarcosine in approx 800 mL of
ddH2O. Adjust volume to 1 L with ddH2O and autoclave.
19. Nylon membrane (Boehringer, cat. no. 1417 240).
20. PCR plates, 96-well (Advanced Biotechnologies, Epsom, England, cat. no.
21. Size marker  Φ×174/HaeIII (0.1 mg/mL, Promega, Southampton, England,
cat. no. G1761). Add 450 µL of ddH2O to vial of the size marker. Add 20 µL of
326 Middleton, Curran, and Williams
GLB to 12 µL (1.2 µg) of the size marker and 8 µL of Tris-EDTA (TE) buffer.
Store at 4°C.
22. 10% SDS. This reagent is extremely harmful if inhaled. Wear a mask when work-ing with SDS powder. Also wear gloves. Wash skin thoroughly if in contact with
SDS. Wipe down work area after use. Preferably add SDS to ddH2O in fume
cupboard. SDS sometimes comes out of solution but will go back on heating.
Add 100 g of SDS in parts to approx 800 mL of ddH2O. As SDS is supplied in
100 g tubs there is no need to measure. Apply heat (up to 68°C) if necessary
to assist dissolution. Allow to cool to room temperature and adjust the volume to
1 L. Do not autoclave.
23. 20× SSPE: 3 M NaCl, 0.2 M NaH2PO4, 0.02 M EDTA, pH 7.4. Add 350.6 g of
NaCl followed by 48 g of NaH2PO4 to approx 1600 mL of ddH2O. Then add
80 mL of 0.5 M EDTA, pH 8.0. Adjust the pH to 7.4 using 4 M NaOH. Adjust
volume to 2 L and sterlize by autoclaving.
24. Saran Wrap® (Genetic Research Instrumentation, Felsted, England, SW1).
25. 2× SSPE/0.1% SDS: Combine 240 mL of 20× SSPE and 24 mL of 10% SDS.
Make up to 2400 mL with ddH2O.
26. 5× SSPE/0.1% SDS: Combine 600 mL of 20× SSPE and 24 mL of 10% SDS.
Make up to 2400 mL with ddH2O.
27. Sodium saline citrate (2×, pH 7.0) (SSC): 0.3  M NaCl + 0.03  M trisodium
28. Ta q enzyme (Bioline, London, England, cat. no. M958013).
29. Thermofast Plate (Advanced Biotechnologies, cat. no. AB-0600).
30. 1 M Tris, pH 7.6. Add 242.28 g of Tris base to 1400 mL of ddH2O. Adjust the pH
to 7.6 by adding 100 mL of concentrated HCl (take care — wear a mask and
goggles and where possible perform this in a fume cupboard). Allow the solution
to cool to room temperature before making final adjustments to the pH. Make up
to 2 L with ddH2O and sterilize by autoclaving. If the 1 M solution has a yellow
color, discard it and obtain better quality Tris. More than 100 mL of conc HCl
may be required.
31. Tris borate EDTA (10×) (TBE): Add 216 g of Tris, 110 g of orthoboric acid, and
80 mL of 0.5 M EDTA to 1400 mL of ddH2O. Adjust volume to 2 L with ddH2O
and sterilize by autoclaving.
32. TE buffer (10 mM Tris, 1 mM EDTA, pH 7.6). Combine 10 mL 1 M of Tris, pH
7.6, with 2 mL of 0.5  M EDTA and make up to 1 L with ddH2O. Sterilize by
autoclaving and aliquot.
2.1. Equipment
1. Enzyme boxes (Boehringer, cat. no. 800058).
2. Gel sealer and casting tray (Merck, Poole, England, cat. no. 306/7252/12).
3. Robbins hybridization incubator (Robbins Scientific, Sunnyvale, CA, USA, cat.
no. 1040-60-2).
4. Robbins Hydra dot blotting machine (Robbins Scientific, cat. no. 1029-60-1).
5. Robbins water bath (Robbins Scientific, cat. no. 1051-20-2).
Immunogenetics and Life-Span 327
3. Methods
3.1. PCR (using 96 well plates)
1. Heat DNA samples to be tested to 60°C for 5–10 min, vortex-mix, and centrifuge
for 5 s in microcentrifuge (see Note 1).
2. Prepare 10 mL of mastermix for appropriate locus (Table 2). Use dH2O of ultra
high purity quality. Dispense 100 µL slowly into tubes of the 96-well plate. Take
care to avoid splashes and air bubbles at the bottom of the tubes. When all tubes
have been filled, cover the 96-well plate with a sterile microtiter tray lid (see
Note 2).
3. Add 1  µL of DNA sample to each well from position 1A→1H, 2A→2H, etc.
Only one row at a time should be uncovered by the lid. Leave two wells with
master mix only, to act as negative controls, and leave appropriate number of
wells for control DNA (see Note 11). When a complete row of DNA samples
have been added, place a strip of eight caps over these samples and press down
gently. When DNA samples have been added to all tubes and caps are in place,
use a cap sealing tool to ensure that all caps are pushed firmly into place.
4. Centrifuge the plate for 1 min at 500g, place in PCR machine, and run appropri-ate cycle program (Table 3). After amplification, if the PCR samples are not to be
processed immediately, store at –20°C (see Note 3).
3.2. Electrophoresis of PCR Samples
1. Add 4.5 g of agarose to 300 mL of 1× TBE, boil, and allow solution to cool to 65°C.
It is important to stir agarose while cooling to prevent “lumps” from forming.
2. While agarose is cooling prepare 96-well-gel template by placing casting tray in
the gel sealer. Take care to ensure gel sealer is not over tightened; otherwise cast-ing tray may separate when agarose is added.
3. Place sealed casting tray on top of a leveling table and adjust the feet of the
leveling table until the bubble in the “spirit level” is centred.
4. Once agarose has cooled to 65°C add 15 µL of ethidium bromide (10 mg/mL)
and mix gently. (Ethidium bromide is mutagenic)
5. Pour the molten agarose solution into the level casting tray. Immediately push
any air bubbles to edges of the template using a pipet tip.
6. Insert four 24-slot combs into the gel, with equal spacing between combs. Allow
gel to set for approx 1 h at room temperature.
7. Add 1000 mL of 1× TBE to an electrophoresis tank. Carefully remove combs
from gel. Remove gel from the gel sealer. Place gel in tank containing 1× TBE
buffer. Ensure gel is covered by buffer to a depth of 2–3 mm.
8. Add 4 µL of each PCR product to a 96-well Thermofast plate. Ensure product is
in each well. Add 8 µL of GLB to each well. Centrifuge plate for 1 min to ensure
9. Load 10 µL of size marker into first well of each of the four rows.
10. Using an octapipet carefully load 10 µL of sample into each well of the gel. Care
must be taken to ensure the octapipet is oriented properly when the samples are
added to the gel.
328 Middleton, Curran, and Williams
Table 2
PCR Master Mixes
Locus Stock conc Master mix
HLA-A generic
ddH2O8220 µL
Cresol red 10 mg/mL 100 µL
NH4 buffer 10× 1000 µL
MgCl2 50 mM 300 µL
dNTPs 20 mM each 100 µL
Each primer (×2) 25 µM 120 µL
Ta q 5 U/µL40 µL
HLA-B generic
ddH2O 12345 µL
Cresol red 10 mg/mL 150 µL
NH4 buffer 10× 1500 µL
MgCl2 50 mM 450 µL
dNTPs 20 mM each 150 µL
Each primer (×3) 25 µM 120 µL
Ta q 5 U/µL45 µL
ddH2O4155 µL
Cresol red 10 mg/mL 50 µL
NH4 buffer 10× 500 µL
MgCl2 50 mM 150 µL
dNTPs 20 mM each 50 µL
Each primer ×225 µM 40 µL
Ta q 5 U/µL15 µL
11. Place the lid of the electrophoresis system on to the electrophoresis tank, connect
the electrodes to the power pack, and electrophorese the samples at 250 V, 250
mAmp for 20 min.
12. Once electrophoresis is complete, remove the gel from the tank and photograph
under UV light. Check size of PCR product against size marker to ensure correct
product has been amplified (Ta ble 1) (see Notes 4 and 5).
3.3. Dot blotting of membranes.
We use the Robbins Hydra dot blotting machine which enables us to make
as many replicate membranes as required from the PCR product. Other labora-tories use other equipment and some will dot blot by hand. If the preparation of
Immunogenetics and Life-Span 329
Table 2 (cont.)
Locus Stock conc Master mix
HLA-C generic
ddH2O8300 µL
Cresol red 10 mg/mL 100 µL
NH4 buffer 10× 1000 µL
MgCl2 50 mM 300 µL
dNTPs 20 mM each 100 µL
Each primer ×225 µM 80 µL
Ta q 5 U/µL40 µL
HLA-DRB generic
ddH2O8280 µL
Cresol red 10 mg/mL 100 µL
NH4 buffer 10× 1000 µL
MgCl2 50 mM 300 µL
dNTPs 20 mM each 100 µL
Each primer (×2) 25 µM 100 µL
Ta q 50 U/µL20 µL
ddH2O8300 µL
Cresol red 10 mg/mL 100 µL
NH4 buffer 10× 1000 µL
MgCl2 50 mM 300 µL
dNTPs 20 mM each 100 µL
Each primer ×225 µM 80 µL
Ta q 5 U/µL40 µL
the required number of membranes proves difficult an alternative method is to
dehybridize used membranes as follows.
1. Dehybridize a maximum of three membranes in 300 mL of each solution with shaking.
2. Rinse membranes in ddH2O for 5 min at room temperature.
3. Wash membranes in 0.4 M NaOH/0.1% SDS at 45°C for 30 min.
4. Wash membranes in 2× SSC for 30 min at room temperature.
5. Check dehybridization is complete by exposing membranes overnight to X-ray
film and developing in usual manner.
6. Store membrane flat at +4°C in a sealed plastic bag if not using immediately.
3.4. Denaturation and Fixing of Blots
1. After PCR product has been dispensed onto the membranes, allow to air-dry for
at least 20 min.
330 Middleton, Curran, and Williams
2. Carefully place membranes DNA face up onto two sheets thick (3MM) Whatman
paper soaked in 0.4 M NaOH. Leave for 10 min. When placing membranes onto
Whatman paper, take care to ensure that membrane is not dragged over denatur-ation pad, all of the membrane soaks up the 0.4 M NaOH, and there are no air
bubbles beneath the membrane.
3. Transfer each membrane onto Whatman paper (3MM) soaked in 10× SSPE. Leave
for 5 min.
4. Gently wash in 2× SSPE and allow to air-dry for at least 25 min.
5. Wrap membranes in Saran Wrap and place (DNA face down) on UV transillumi-nator for 4 min. Ensure that all the UV lights are fully on during the procedure;
do not switch transilluminator off between each step. Place a glass plate on top of
membranes to hold them flat during this procedure. Store membranes wrapped in
tin foil at +4°C if not using immediately.
Table 3
PCR Amplification Conditions
No. of
Locus Hold Cycle cycles Hold Hold
HLA-A 96°C/5 min 96°C/1 min
generic 60°C/30 s 35 72°C/5 min 15°C/forever
72°C/1 min
HLA-B 96°C/5 min 96°C/30 s
generic + 65°C/30 s 72°C/5 min 15°C/forever
HLA-B27 72°C/45 s 32
HLA-C 96°C/5 min 96°C/1 min
generic 66°C/30 s 30 72°C/5 min 15°C/forever
72°C/1 min
HLA-DRB 96°C/5 min 96°C/1 min
generic 55°C/1 min 30 72°C/5 min 15°C/forever
72°C/1 min
HLA-DR 3/11/6 96°C/5 min 96°C/1 min
64°C/1 min 10 72°C/5 min 15°C/forever
72°C/1 min
96°C/1 min
56°C/1 min 20
72°C/1 min
Immunogenetics and Life-Span 331
3.5. 3′ End Labeling of HLA Oligonucleotides
The labeling reagents are obtained in a kit from Boehringer (cat. no.
1362372) (see Notes 6 and 7).
1. Remove all reagents from freezer (except Terminal Transferase; this should be
removed just before use) and allow to thaw. Vortex-mix reagents briefly, and cen-trifuge in a microcentrifuge for 5 s.
2. Combine the following: 4 µL of reaction buffer (5×), 4 µL of CoCl2 (25 mM),
1 µL of DIG–ddUTP (1 mM), 1 µL of Terminal Transferase (50 U), and 100 pmol
of probe. Make up to 20 µL with ddH2O. Vortex-mix samples briefly, centrifuge
in a microcentrifuge for 5 s, and incubate at 37°C for 30 min in a water bath.
3. Centrifuge for 5 s in a microcentrifuge and place on ice for 5 min. Add 80 µL of
ddH2O, vortex mix briefly, and centrifuge in a microcentrifuge for 5 s. Aliquot in
volumes related to the amount of probe used (Tables 4–8) and store at –20°C.
3.6. Prehybridization, Hybridization and SSPE Stringency Washes
Each probe is simultaneously hybridized to two different membranes each
containing 96 DNA samples.
1. Hand roll membranes lengthwise to form a cylinder. Place two membranes in a
hybridization bottle. One membrane should have the DNA side of the membrane
facing the glass, while the second membrane should have the DNA side facing
inwards in the bottle.
2. Add 20 mL of freshly prepared, hybridization buffer. Screw cap on tightly and
clamp to the rotisserie of a Robbins incubator (preset at 45°C). Rotate the bottles
for 1 h.
3. Just before the incubation is complete thaw appropriate aliquots of DIG-labeled
oligonucleotide probe, vortex-mix briefly, and centrifuge for 5 s in a
4. Add the appropriate number of picomoles of probe (Tables 4–8) to 20 mL of
prewarmed (45°C) hybridization buffer and mix by inversion.
5. Remove the hybridization bottle from the incubator and pour off the hybridiza-tion buffer into a disposable collection container. Add 20 mL of hybridization
buffer containing DIG-labeled probe and incubate bottle for 1 h at 45°C.
6. Remove the bottle from the incubator and pour off the fluid into a disposable
collection container.
7. Add 100 mL of 2× SSPE/0.1% SDS. Recap the bottle and place inside a Robbins
incubator (preset to 25°C) and incubate for 10 min. Make sure temperature does
not rise above this.
8. Discard the fluid and repeat step 7.
9. Remove the bottle from the incubator. Uncap the bottle and using forceps care-fully remove the membranes from the bottle, prior to discarding fluid, directly into
a small plastic tray containing 200 mL of 5× SSPE/0.1% SDS, which has been
heated to the appropriate temperature (Tables 4–8) (see Notes 8 and 9). Place one
332 Middleton, Curran, and Williams
membrane DNA face down and the other membrane DNA face up into the wash-ing solution. Incubate with shaking for 40 min. Check temperature reading and record
any variation on the hybridization record sheet. If the temperature varies more
than 2°C above or below the required temperature abandon this hybridization.
10. Remove the membranes from the tray, blot dry, but do not allow the membrane to
dry out. Wrap the membrane in Saran Wrap, and store in tin foil at 4°C, until
ready to perform chemiluminescent detection.
Table 4
Probes Used for HLA-A Typing
Sequence temp Picomoles Nucleotide
Probe 5′  → 3′ (°C) used position
Exon 2
A (56R) GAGAGGCCTGAGTAT 46 40 163–177
B (62LQ) TGGGACCTGCAGACA 48 50 178–192
C (62G) GACGGGGAGACACGG 52 20 181–195
O (62RN) GACCGGAACACACGG 52 20 181–195
D (62EG) GAGGAGACAGGGAAA 46 40 184–198
Y (A276) GGCCCACTCACAGACT 52 50 204–219
E (731) TCACAGATTGACCGA 45 40 211–225
X (A290) CTGACCGAGTGGACCT 51 40 218–233
R (A26) TGACCGAGCGAACCTG 54 40 219–234
F (77S) GAGAGCCTGCGGATC 50 20 226–240
Exon 3
Z (A347) CTCACACCATCCAGA 45 70 5–19
T (95V) CACACCGTCCAGAGG 48 40 7–21
P (114EH) TATGAACAGCACGCC 46 30 67–81
G (131R) CGCTCTTGGACCGCG 52 40 121–136
H (142TK) ACCACCAAGCACAAG 46 40 154–168
I (149T) TGGGAGACGGCCCAT 50 40 169–183
J (150V) GAGGCGGTCCATGCG 60 20 172–186
K (151R) GCGGCCCGTGTGGCG 60 20 175–189
1 (A525) TGAGGCGGAGCAGTTG 54 40 183–198
N (156Q) GAGCAGCAGAGAGCC 52 20 190–204
Q (156W) GAGCAGTGGAGAGCC 50 10 190–204
L (161D) CTGGATGGCACGTGC 50 20 208–222
V (A551) TGGAGGGCACGTGCGT 56 40 209–224
M (163R) GAGGGCCGGTGCGTG 54 20 211–225
S (A355) GGCGAGTGCGTGGAGTGGC 68 10 214–232
U (A357) GGCGAGTGCGTGGACGGGC 68 10 214–232
Immunogenetics and Life-Span 333
Table 5
Probes Used for HLA-B Typing
Sequence temp Picomoles Nucleotide
Probe 5′   → 3′ (°C) used position
Exon 2
31 (B89) GGTATTTCGACACCGCC 56 40 17–33
32 (B156) GGACGGCACCCAGTT 52 40 84–98
33 (B168) GTTCGTGCGGTTCGA 50 40 96–110
09 (BL09) GAGTCCGAGAGAGGAGCC 57 6 123–140
01 (BL01) GAGGAAGGAGCCGCGGGC 64 20 129–146
02 (BL02) GAGGACGGAGCCCCGGGC 64 40 129–146
07 (BL07) GAGGATGGCGCCCCGGGC 64 60 129–146
34 (B249) TTGGGACGGGGAGAC 50 40 177–191
24 (BL24) GGGAGACACAGATCTCCA 55 40 185–202
05 (BL05) ACACAGATCTTCAAGACC 55 14 190–207
10 (BL10) GATCTACAAGGCCCAGGC 58 10 195–212
12 (BL12) ATCTGCAAGGCCAAGGCA 56 20 196–213
18 (BL18) ACTGACCGAGTGAGCCTG 58 20 217–234
35 (B73) ACTGACCGAGTGGGCCTG 63 40 217–234
20 (BL20)* AGCGGAGCGCGGTGCGCA 64 40 233–250
21 (BL21) CGGAACCTGCGCGGCTAC 62 40 235–252
22 (BL22) CGGACCCTGCTCCGCTAC 61 30 235–252
23 (BL23) CGGATCGCGCTCCGCTAC 62 40 235–252
Exon 3
36 (B348) TCACACCATCCAGAGG 49 50 6–21
37 (B354) CATCCAGGTGATGTAT 46 40 12–27
28 (BL28) CCAGTGGATGTATGGCTG 56 40 15–32
38 (B361) AGGATGTTTGGCTGC 48 40 19–33
26 (BL26) CTGCGACCTGGGGCCCGA 65 40 30–47
30 (BL30) GGCATAACCAGTTAGCCT 54 50 65–82
39 (B409) TATGACCAGGACGCCT 55 40 67–82
40 (B427) GACGGCAAAGATTACA 46 40 85–100
41 (B499) ACCCAGCTCAAGTGG 47 40 157–171
42 (B505) CGCAAGTTGGAGGC 46 40 163–176
43 (B532) GAGCAGCTGAGAGCCT 52 40 190–205
44 (B539) GAGAACCTACCTGGA 46 40 197–211
45 (B553a) GAGGGCCTGTGCGT 48 40 211–224
46 (B553b) GAGGGCACGTGCGT 48 40 211–224
47 (B566) TGGAGTCGCTCCGC 48 40 224–237
48 (B597) GAAGGACACGCTGGA 52 40 255–269
49 (B599) AGGACAAGCTGGAGCG 52 40 257–272
aComplementary to coding sequence.
334 Middleton, Curran, and Williams
3.7. Chemiluminescence
All steps are performed at room temperature with shaking using a platform
shaker. Use separate enzyme storage boxes for different buffer solutions and
keep light-tight. Use one enzyme box for a maximum of three membranes at
the same time (see Note 10).
1. Add 240 mL of anti-DIG–ALP conjugate in buffer 2 to the enzyme box. Place
membranes into the boxes DNA side down. Incubate for 15 min on shaker.
Table 6
Probes Used for HLA–C Typing
Sequence temp Picomoles Nucleotide
Probe 5′    → 3′ (°C) used position
Exon 2
15 (C2D6) AGCCCCGGGCGCCGT 56 35 137–151
3 (C2G2) AGTGAACCTGCGGAAACTG 59 25 225–243
2 (C2G1) TGAGCCTGCGGAACCTG 56 30 227–243
17 (C2H303) CCAGAGCGAGGCCAGT 54 25 258–2
(Intron 2)
Exon 3
21 (C3A14) CTCCAGTGGATGTTTGGC 56 25 13–30
4 (C3A1) TCCAGTGGATGTGTGGC 54 25 14–30
19 (C3A4) CAGAGGATGTTTGGCTGC 56 20 16–33
12 (C3A7023) AGGATGTCTGGCTGCGA 54 25 19–35
7(C3A212) TGTACGGCTGCGACCTG 56 20 23–39
22 (C3C15) GGCATGACCAGTTAGCC 54 25 65–81
20 (C3D58) GCCCTGAATGAGGACCT 55 30 103–119
6 (C3E1203) TCCTGGACTGCCGCGG 56 25 124–139
5 (C3E12) GGACCGCTGCGGACAC 56 25 128–143
11 (C3G17712) CGCAAGTTGGAGGCGG 54 25 163–178
13 (C3G716) GGCCCGTGCGGCGGA 56 25 177–191
23 (C3G8013) GCCCGTACGGCGGAG 54 25 178–192
8(C3G2612) TGAGGCGGAGCAGTGGA 57 25 183–199
14 (C3G16) GCGGCGGAGCAGCAGA 57 25 184–199
9 (C3H2) GGAGGGCGAGTGCGTG 57 25 210–225
16 (C3H3) GGAGGGCCTGTGCGTG 56 25 210–225
10 (C3J17) GCTCCGCGGATACCTG 54 25 231–246
Immunogenetics and Life-Span 335
2. Transfer membranes to 300 mL of washing buffer and incubate for 15 min on
shaker. Discard washing buffer and replace with fresh washing buffer and incu-bate for a further 15 min.
3. Transfer membrane to 300 mL of buffer 3 and incubate for 5 min on shaker.
4. Remove from buffer 3, and place two membranes back to back in a plastic bag.
Add 20 mL of CSPD (1:100 dilution) and reseal the bag. Place the bag on a
platform shaker, cover with tin foil, and shake for 5 min at room temperature.
5. Pour off CSPD fluid into a 20-mL plastic tube for reuse (up to 5×). Store at –20°C
if using on more than 1 d, but note that CSPD should only be frozen once. Care-fully remove the membrane from the bag, blot off excess liquid, and wrap in
Saran Wrap.
Table 7
Probes Used for HLA-DR Typing
Sequence temp Picomoles Nucleotide
Probe 5′    → 3′ (°C) used position
Exon 2
09 (1007) GAAGCAGGATAAGTTTGA 50 10 24–41
03 (1008N) GAGGAGGTTAAGTTTGAG 54 2 25–42
07 (1004) GAGCAGGTTAAACATGAG 56 4 25–42
08 (1006) TGGCAGGGTAAGTATAAG 50 10 25–42
06 (1003) GTACTCTACGTCTGAGTG 56 4 27–44
02 (1002) AGCCTAAGAGGGAGTGTC 56 30 29–46
18 (DR18) CTACGGGTGAGTGTTAT 48 40 32–48
10 (2810) GCGAGTGTGGAACCTGAT 56 10 66–83
01 (2801) CGGTTGCTGGAAAGATGC 60 6 73–90
25 (DR25) CGGTTCCTGGACAGATA 52 40 73–89
11 (DRB12) CAGGAGGAGCTCCTGCGC 58 4 100–117
22 (DR22) CGGCCTAGCGCCGAGTA 5850 163–179
05 (5703) GCCTGATGAGGAGTACTG 54 20 165–182
15 (DRB14/1) GGCCTGCTGCGGAGCACT 64 4 164–181
14 (7031) CTGGAAGACAAGCGGGCCG 60 30 202–220
16 (DRB13) TGGAAGACGAGCGGGCCG 64 3 203–220
24 (DR24) AGCGGAGGCGGGCCGAG 62 40 206–222
17 (7012)* ACCGCGGCCCGCCTCTGC 66 30 207–224
23 (7005)* ACCGCGGCCCGCTTCTGC 66 40 207–224
12 (DRB8) GCGGGCCCTGGTGGACAC 64 20 213–230
04 (7004) GGCCGGGTGGACAACTAC 62 1 217–234
aComplementary to coding sequence.
336 Middleton, Curran, and Williams
6. Tape two membranes to the one X-ray film and place a second film on top.
Expose the top film for 5 min and check the intensity of the dots. Depending on
these results process the second film accordingly. It may be necessary to
reexpose the membrane to a third or fourth film for a further period of time,
depending on dot intensity.
7. Record the probe reaction for each sample and analyze according to the known
patterns (Tables 9–13) using a computer programme (see Notes 11 and 12).
4. Notes
1. We do not routinely determine the concentration of DNA in each isolation. When
isolating DNA the amount of TE buffer added to the pellet of DNA is judged by
eye. However, we assess approx 10% of samples to ensure that the DNA is at an
appropriate concentration. For our methods we normally have the DNA concen-tration at approx 0.5 µg/µL.
2. When setting up a PCR wear a separate laboratory coat, wear gloves and change
them frequently, and perform all work in pre-PCR room using dedicated equip-ment. Pipettes should not be removed from the pre-PCR room. Pipets are labeled
according to reagents and must be used only for these reagents. The use of tips
with filters is advisable. When preparing the master mix thaw out following re-agents (MgCl2, dNTPs, PCR buffer, and appropriate primers). Vortex-mix each
reagent briefly and centrifuge in a microcentrifuge for 5 s and place in an ice
bucket (PCR buffer and MgCl2 should be centrifuged for 2 min). Ta q polymerase
should always be added last, after vortex-mixing and centrifuging, and just prior
to dispensing the master mix. The aliquoted master mixes should not be left on
Table 8
Probes Used for HLA-DR 3/11/6 Group
Sequence temp Picomoles Nucleotide
Probe 5′ → 3′ (°C) used position
Exon 2
1 (DR19) CGGTACCTGGACAGAT 5040 73–88
2 (5703) GCCTGATGAGGAGTACTG 54 20 165–182
3 (DRB14/1) GGCCTGCTGCGGAGCACT 64 4 164–181
4 (7031) CTGGAAGACAAGCGGGCCG 60 30 202–220
6 (DR24) AGCGGAGGCGGGCCGAG 62 40 206–222
7 (7012) ACCGCGGCCCGCCTCTGC 66 30 207–224
8 (7005) ACCGCGGCCCGCTTCTGC 65 40 207–224
10 (7004) GGCCGGGTGGACAACTAC 62 1 217–234
11 (5701) GCCTGATGCCGAGTACTG 58 40 165–182
Immunogenetics and Life-Span 337
Table 9
HLA-A SSOP Patterns
Probes A B C D E F G H I J K L M N O P Q R S T U V W X Y Z 1
HLA-A alleles
0101 + + + + + + +
0102 + + + + + +
2.1a +++ ++++
0202 + + + + + + +
0203 + + + + + + + + +
0204/17 + + + + + +
0205/08 + + + + + +
0206/10/21 + + + + + +
0211 + + + + + + + (+)
0212/13 + + + + + + + +
0214 + + + + +
0216 + + + + + + +
0219 + + + + + + +
0222 + + + + + + + +
0301/03N + + + + + +
0302 + + + + + +
1101/02/03 + + + + +
1104 + + + + +
2301 + + + + +
2402/03/05 + + + + +
2404 + + + + + +
2406 + + + + +
2407 + + + +
2408 + + + + +
2409N + + + + +
2410 + + + + +
2413 + + + +
2414 + + + + + +
2501 + + + + + + + +
2502 + + + + + + +
the bench too long (max 15 min —  Ta q loses activity once diluted in buffer).
Switch on the PCR machine for at least 10 min prior to use to allow the machine
to heat up. PCR machine should be situated in the post-PCR room.
3. After setting up a PCR, wash work areas with sodium hypochlorite (containing
2% chlorine). Soak all racks used to hold samples in sodium hypochlorite for
approx 30 min, and rinse thoroughly in water. Pipets should be wiped with sodium
hypochlorite, followed by ddH2O. Wipe microcentrifuge, vortex-mix, freezer
338 Middleton, Curran, and Williams
Table 9 (cont.)
Probes A B C D E F G H I J K L M N O P Q R S T U V W X Y Z 1
2601/02 + + + + + + + +
2603/05/06 + + + + + + +
2604 + + + + + + +
2607 + + + + + + + +
2608 + + + + + + + +
68011/012/02 + + + + + + +
6803 + + + + + + + +
6804 + + + + + + (+) +
6805 + + + + + + +
6901 + + + + + +
2901/02/03 + + + + + +
3001 + + + + + +
3002 + + + + + +
3003 + + + + +
3004 + + + + + + +
31012 + + + + + (+) +
3201 + + + + + + +
3202 + + + + + + +
3301/03 + + + + + (+) +
3401 + + + + + + +
3402 + + + + + + +
3601 + + + + + + +
4301 + + + + + + + +
6601 + + + + + + +
6602 + + + + + + +
6603 + + + + + + + +
7401/02/03 + + + + + + +
8001 + (+) + + + +
The alleles listed are those identified in HLA-A sequences from ASHI, April 1997.
a2.1 represents the alleles 0201, 0207, 0215 N, 0218, 0220.
A (+) indicates probe is positive in practice but would not appear to be from sequence.
handle, etc., with sodium hypochlorite. Expose the working area, including pipets
etc., to UV light for 60 min.
4. When performing a PCR on 96 samples there may be one or two samples that are
not amplified. Therefore we always run a gel to ensure that we have product. This
enables the SSOP method to be well controlled. (If an amplification fails in the
Sequence Specific Primer method this would lead to an incorrect result.) On some
occasions the product is deemed weak and this sample will always be repeated.
Good amplification always gives a clean and clear-cut SSOP hybridization
whereas almost all the problematic typing results we have encountered were due
Immunogenetics and Life-Span 339
Table 10
HLA-B SSOP Patterns
HLA-B 7 0 0 0 0 0 0 1 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 3 3 4 4 4
allele . 7 7 8 8 8 8 3 3 4 8 7 7 7 7 5 5 5 5 5 5 7 7 8 8 9 9 0 0 0
2 00000000 00 . 0 . 0 . 1120000 . . 0 . 0
38123413 122 8384590121223125
Probe 2 8
01 + + + +
02 + ++++ +++
05 ++++ +++++
07 + +
09 + + +++++ + +++ + ++++
10 +
12 +++ +
20 + + ++ +
21 + + ++ + ++ +++++++ +++++
22 + + +
23 + + +
24 + + + + ++++
26 + + +++
28 +
30 + +
31 + + ++
32 +
36 +
41 +
42 +
43 + ++++++ ++ ++++ + +++
44 ++++
45 + +++ ++ +
46 + + ++ ++ + ++++
48 + + ++
49 + + + +
340 Middleton, Curran, and Williams
Table 10 (cont.)
HLA-B 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 5 5 5 5 5 5 5 5 5 5 5
allele 0 0 1 1 2 4 4 4 4 4 4 4 5 6 7 7 8 8 9 0 1 1 1 2 3 4 5 5 5 5
000000 . 0000100 000000 . 00 0 . 000
78121214568011 121211157 11346
Probe 3 8
01 + + ++ ++++ ++ ++ ++
02 + + ++++
05 + + + + + +
07 + +
09 + + ++ +++
10 + + + + +
18 + +
20 ++++ ++
21 + + +++ ++ +++ + +++++
22 +
23 + + + ++++
24 + + ++++ +++ ++++++ +
26 + + + + + +
27 + + + +
30 + + + +
33 +
38 + + +
39 +++ ++
40 + +++
42 + +
43 + +++++++ +++ ++++++++
45 + + ++++++ ++++++++
46 +++ + +++++
47 ++++++++
48 +++
49 + +
Immunogenetics and Life-Span 341
Table 10 (cont.)
HLA-B 5 5 5 5 5 5 5 5 6 7 7 8 8 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1
allele 6 6 7 7 7 8 8 9 7 3 8 1 2 5 5 5 5 5 5 5 5 5 5 5 5 5 5 5 5 5
. 0000000 0 00 . 000000 . . 11122222
1 3124121 1 113 2356895434602347
Probe 3 1 5 7
02 + + + +
05 + + + +
07 +++ + + +++ +++++ ++
09 + + + + + + ++ ++ +
10 + + + + +
12 +
21 + + ++++++ ++++++++ + ++ +
23 ++++++ + + ++
24 + +++ + + ++
26 + + + +
27 +
28 +
30 + + +
34 +++++
35 +
37 +++
38 + + +
39 +
40 + +
42 +
43 + + +++ +++ ++++ ++ + ++ +
44 +
45 + + +++++ + + ++++++++++++++++
46 + +
47 + +
49 +
342 Middleton, Curran, and Williams
Table 10 (cont.)
allele 5
Designation of groups
01 7.2 0702 0704 0705 0706 0707
02 14 1401 1402
05 + 18 1801 1802 1803
07 4001 40011 40012
09 + 40.2 4002 4003 4004 4006 4009
10 44.1 44031 44032 4407
12 51.1 51011 51012 51021 51022 5103 5104 5106
18 39.2 39011 39013 3903 3904 3905 39061 39062 3907 3909 3910 3911 3912
20 39.3 39021 39022 3908
21 + 35.3 3501 3502 3503 3504 3505 3506 3507 35091 35092 3510 3511 3513 3521
22 35.4 3512 3516 3517
23 27.2 2703 2704 27052 27053 2706 2707 2709 2710 2711
24 15.3 1501 1504 1507 1512 1519 1526 N 1530 1532 1533 1534 1535
26 15.4 1511 1515 1528 1531*
27 15.5 1510 1518 1537
28 55.1 5501 5502 5505
30 56.1 5601 5602
31 67 67011 67012
32 78 7801 78021 78022
33 52 52011 52012
43 +
45 +
The alleles listed are those identified in HLA-B sequences from ASHI, April 1997.
aNB 1531 full sequence data not available for probes B532 and B553A.
Immunogenetics and Life-Span 343
Table 11
HLA-C SSOP Patterns
Probes 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23
01 + + +
02021 + + + + +
02022/024 + + + + + +
02023 + + + + +
0302 + + + +
0303 + + + +
0304 + + +
04 + + +
0501 + + +
0602 + + + + +
7.1a ++ + + +
0702 + + + + + +
0703 + + + + +
0704 + + + + +
0707 + + + + +
0801/03 + + + +
0802 + + +
1202 + + + + + +
1203 + + + + +
1204 + + + +
1301 + + + + + + +
14 + + + +
1502/03 + + +
1504 + + +
1505 + +
1601 + + + + +
1602 + + + + +
17 + + +++
18 + + + +
The alleles listed are those identified in HLA-C sequences from ASHI, April 1997.
a7.1 represents 0701, 0705, 0706.
to poor amplification. Interpretation of weak hybridization signals can give an
incorrect result.
5. When determining the conditions necessary for this technique one important
aspect is to ensure that the amplification works for both alleles. On some occa-sions when we were determining the amplification conditions we found that a
product could be obtained whereby only one allele could be detected. Therefore,
344 Middleton, Curran, and Williams
Table 12
HLA-DR SSOP Patterns
Probe 01 02 03 04 05 06 07 08 09 10 11 12 13 14 15 16 17 18 22 23 24 25 26
HLA-DR allele
1 1.1 + + +
01022 + +
0103 + + +
2 15.2 + + +
1503/1607 + +
16.1 + + +
1604 + + + +
1608 + + + +
3 3.1 + + + +
3.2 + + +
0304 + + +
0308 + + + + +
4 4.2 + + + + + +
4.3 ++ ++++
4.5 + + + + +
4.6 + + + + + +
4.7 + + + + + +
4.8 + + + + + + +
0409 + + +++++
0412 + + + + + + +
0415 + + + + + +
0418 + + + + + +
0422 + + + + + +
11 11.2 + + + + +
11.4 + + + +
11.16 + + + + + +
1105/30 + + +
1107 + + + + +
1109 + + + + +
1117 + + + + +
1122 + + + +
1123/25 + + + + +
1126 + + + + +
12 12.1 + + +
1204 + + + +
1205 + +
Immunogenetics and Life-Span 345
Table 12 (cont.)
Probe 01 02 03 04 05 06 07 08 09 10 11 12 13 14 15 16 17 18 22 23 24 25 26
HLA-DR allele
13 13.2 + + + + +
13.3 + + +
13.5 + + + +
13.6 + + + +
13.8 + + + +
13031/32 + + + + +
1304 + + + + +
1310 + + + + +
1313 + + + + +
1315/27 + + + +
1317 + + + +
1318 + + + + +
1319 + + +
1326 + + +
14 14.1 + + + + +
14.3 + + + +
14.4 + + + +
14.5 + + + +
14.7 + + + + +
1404/28 + + + + +
1410 + + + + +
1411 + + + + +
1413 + + + + +
1415 + + + +
1416 + + + + +
1418 + + + +
1419 + + + +
1420 + + +
1421 + + + + +
1422/25 + + + +
1423 + + + +
1424 + + +
1426 + + + +
7 0701 + + + +
8 8.2 + + + + +
8.3 + + + +
0808 + + + + +
9 09012 + + + +
10 1001 + +
The alleles listed are those identified in HLA-DR sequences from ASHI, April 1997.
346 Middleton, Curran, and Williams
Table 12 (cont.)
Designation of groups:
1.1 0101 01021 0104
15.2 15011 15012 15021 15022 1504 1505 1506
16.1 16011 16012 16021 16022 1603 1605
3.1 03011 03021 0303 0305 0306 0307
3.2 03012 03022 0311
4.2 0402 0414
4.3 04011 04012 0413 0416 0421
4.5 0403 0406 0407 0420
4.6 0411 0417 0424
4.7 0404 0408 0419 0423
4.8 04051 04052 0410
11.2 1102 1103 1111 1114 1121
11.4 11011 11012 11013 11041 11042 1106 11081 11082 1110 1112
1113 1115 1118 1119 1124 1127 1129
11.16 1116 1120
12.1 1201 12021 12022 12031 12032
13.2 1301 1302 1316 1320 1328 1329
13.3 1307 1311 1314 1325
13.5 1305 1306 1309
13.6 1312 1321 1330
13.8 1308 1322 1323 1324
14.1 1401 1407
14.3 1403 1412 1427
14.4 1402 1406 1429
14.5 1405 1408 1414
14.7 1409 1417
8.2 0801 08031 08032 0805 0806 0810 0812 0814 0816
8.3 08021 08022 08041 08042 0807 0809 0811 0813 0815
if a laboratory is setting up a technique from scratch it should ensure that there is
no differential amplification by testing various combinations of alleles.
6. Many of the probes used in this laboratory are DIG-labeled during their manufac-ture, adding the DIG moiety to 5′ amino oligonucleotides by incubating with a
DIG ester under mild alkali conditions.
7. A commercial method similar to that described is now available (Lifecodes Cor-poration, Stamford, CT, USA) whereby probes are supplied already labeled with
alkaline phosphatase. This removes the DIG labeling of probes and the use of
anti-DIG antibody.
8. When we implement new probes to the system we will initially use a wash tem-perature that is equivalent to the melting temperature of the probes. This is equal
Immunogenetics and Life-Span 347
Table 13
HLA-DR 3/11/6 Subgroup SSOP Patterns
DR3 3.1 + + +
3.2 + +
3.3 + +
0308 + + +
DR11 11.1 +
11.2 + +
11.3 + +
1107 + +
1117 + +
1126 + +
DR13 13.1 + +
13.2/1424 +
1303 +
1304 +
1310 + +
1313 +
1318/14.3 + +
1327 + + +
DR14 14.1 + +
14.2 + +
14.4 +
14.5 + +
14.6 + +
14.7 +
1413 +
1416 + +
in °C to 2× (number of A + T bases) +4 × (number of G + C bases). Thereafter we
adjust the wash temperature by 1°C either up or down according to the probe
reaction at the melting temperature. We also start with 20 pmol of probe and
adjust accordingly.
9. When the probe conditions, that is, number of picomoles and wash temperature
have been determined it is worthwhile to keep a record on the performance of the
probes, that is, whether the probe is not giving an adequate signal with its posi-tive control or whether it is crossreacting with controls with which it should be
negative. On occasions the conditions for the probes need to be altered. This in a
way is similar to HLA sera whereby after long-term storage the specificities iden-
348 Middleton, Curran, and Williams
Table 13 (cont.)
Designation of groups:
3.1 03011 0304 0305 0306
3.2 03012 0311
3.3 03021 03022 0303 0307
11.1 11011 11012 11013 11041 11042 1106 11081 11082 1109 1110 1112 1113 1115
1118 1119 1124 1127 1129
11.2 1102 1103 1111 1114 1116 1120 1121
11.3 1123 1125
13.1 1301 1302 1308 1315 1316 1319 1320 1322 1323 1324 1328 1329
13.2 1305 1306 1307 1309 1311 1314 1325
1303 13031 13032
13.4 1312 1321 1326 1330
14.1 1401 1407
14.2 1402 1406 1409 1417 1420 1429
14.3 1403 1412 1427
14.4 1405 1408 1418
14.5 1414 1423
14.6 1419 1421
14.7 1422 1425 1426
Alleles not amplified by this system: 1105 1122 1130
1404 1410 1411 1415 1428
tified can change. If a probe appears to be giving strong false-positive reactions
we will initially increase the wash temperature by 1°C, or if giving weak false-positive reactions we will decrease the probe concentration by approx 20%. If a
probe appears to be giving false-negative results we will decrease the wash
temperature by 1°C. If a probe is giving weak reactions we will initially increase
the probe concentration by approx 20%. One way to monitor the performance
of the probes is to record the length of time needed for autoradiography expo-sure. If this varies to such an extent that it takes more than 30 min to achieve a
good signal the conditions of the probe should be altered.
10. It is normal practice in this laboratory for chemiluminescent detection to be per-formed on 24 membranes at the same time. All membranes are processed up to
the end of step 2. Thereafter membranes are processed in groups of six simulta-neously, leaving the remaining membranes in the washing buffer.
11. Enough controls should be included so that each probe will have two positive
reactions. In addition control DNA should be included as negative controls. These
contain alleles with sequences that are closely related to the sequence that
the probe detects and with which the probe might cross-hybridize. This is espe-cially important when initally determining the optimum conditions for the probe
Immunogenetics and Life-Span 349
to work. To maintain consistency between membranes we try to use the same
controls. If a laboratory finds it difficult to have a large enough supply of the
same control DNA it may consider cloning control DNA by long-range amplifi-cation (4). (Owing to lack of sequence information this is practical only for HLA-A and -B at present). This gives material to use in as many tests as needed. This is
especially important when the control DNA has been obtained from an outside
12. In this laboratory we always have two independent readings of the membrane.
We do not believe in recording a result according to the strength of the reaction
(e.g., 1, 2, 4, 6, 8, as in serology). The result should be positive or negative. If in
doubt it should be repeated. In the future it would be beneficial to all laboratories
if a scanning mechanism was available for reading the membranes as mistakes
are possible in the transmissions of results. We believe it is important that the
probe patterns are not analyzed by eye. It would be far too easy to see the obvious
allele(s) when examining the probe patterns rather than those that are obscure. To
overcome this laboratories should have a computer program.
1. Rea, I. M. and Middleton, D. (1994) Is the phenotypic combination A1 B8 Cw7
DR3 a marker for male longevity? J. Am. Geriatr. Soc. 42, 978–983.
2. Opelz, G., Mytilineos, J., Scherer, S., Dunckley, H., Trejaut, J., Chapman, J.,
Middleton, D., Savage, D., Fischer, O., Bignon, J., Bensa, J., Albert, W., and Noreen,
H. (1991) Survival of DNA HLA-DR typed and matched cadaver kidney trans-plants. Lancet 338, 461–463.
3. Charron, D. and Fauchet, R., eds (1997) HLA. Genetic Diversity of HLA Functional
and Medical Applications. Vol. 1 Workshop, EDK, Paris, France.
4. Curran, M. D., Williams, F., Earle, J. A. P., Rima, B. K., Van Dam, M. G., Bunce,
M., and Middleton, D. (1996) Long range PCR amplification as an alternative strat-egy for characterizing novel HLA-B alleles. Eur. J. Immunogenetics 23, 297–309.
Dietary Restriction 353
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
Dietary Restriction and Life-Span Extension
Byung Pal Yu
1. Introduction
The popularity of the dietary restriction (DR) paradigm (often used inter-changeably with calorie restriction) among gerontologists is primarily based
on the research finding of the last two decades. Originally discovered by
McCay’s group in the 1930s, this paradigm showed that animals placed on the
DR regimen, meaning reduced nutrition without malnutrition, had robust life
extensions (1). This scientific breakthrough showed that nutritional status can
bring about distinctive metabolic adjustments.
Among the most obvious phenotypic changes seen with the implementation
of DR are reduced body weight or size, slow growth, leanness due to reduced
adipose mass, young appearance, and agility. The hallmark of these age-related
changes, however, is a robustly extended life-span, accompanied by low mor-bidity and mortality rates (2). The life prolonging action of DR is unparalleled
with any other laboratory paradigm in its ability to induce such a broad spec-trum of physiological and pathological anti-aging effects (3). The unequivocal
experimental data confirmed by many laboratories around the world have estab-lished dietary intervention as the most effective and dependable tool or proce-dure available tool today for gerontologists in their exploration of the aging
As an experimental tool, DR has at least three important characteristics that
make it the gold standard of aging research (1,4,5). They are as follows:
1. Simplicity of execution: Studies using this paradigm are simple to execute and the
procedures are easy to implement. Unlike other nutritional manipulations in
which either individual dietary components or ingredients are adjusted either
quantitatively or qualitatively, DR can be administered by simply reducing the
amount of the food given, thereby resulting in the calorie restriction. This simple-
354 Yu
to-implement intervention and calorie reduction (not anything else) are the most
important assets of the DR paradigm.
2. Reproducibility: Good laboratory procedures must effectively reproduce consis-tent results, regardless of the location of the experiment or the responsible inves-tigator. To date, no reproducibility failure has been reported in the ability of DR
to extend life in laboratory animals, when conducted under proper experimental
conditions and conscientiously followed procedures.
3. Diverse effect: The DR paradigm has a broad application. Not only has it been
used in the study of laboratory rats and mice, but it also has been proven effective
when applied to the study of lower organisms, such as in studies using fruit flies
and nematodes. In addition, several investigators have shown DR to work equally
well using both sexes of rodents, allowing DR to manipulate physiological sys-tems without concern regarding their differences. Another factor that often raises
questions relating to the use of DR is age at implementation. Studies indicate that
timing is not a problem with the DR paradigm. DR shows effective results when
imposed at any point during the life cycle, whether pre- or postpuberty, maturity,
or even during early senescence (1,4,6,7).
2. Materials
DR, as a dietary manipulation, requires no special tools or instrumentation.
However, for a more appropriate discussion, in lieu of the chemical reagents
and instruments normally found in the laboratory, descriptions of desirable (if
not required) facilities and features are needed, which researchers should con-sider to obtain optimal results. The main reason for these considerations is that
DR work requires the use of a long-term, chronic feeding paradigm. Readers
interested in understanding the various forms of the DR paradigm in detail
should consult the recent publication of Bertrand et al. (5).
1. The barrier facility: It should be stated at the onset of this discussion that DR
studies do not require specific barrier conditions. In fact, many aging studies
claim that experiments can be carried out successfully using conventional animal
facilities without the use of a specific pathogen-free facility (4–6,8,9). Research-ers should be aware, however, that aging studies using the DR paradigm often
involve life-span measurements, pathological monitoring, and survival analysis.
It is therefore this author’s view and recommendation that if only a conventional
facility is available, a separate location for long-term housing of animals is essen-tial, one that is as far as possible from other animal facilities. If logistically pos-sible, investigators should conduct long-term, chronic studies under specific
pathogen-free conditions, or at least in a self-contained facility that is isolated
from the institution’s usual animal facility. This precaution aids in the prevention
of cross-contamination by minimizing the heavy trafficking that results in mixed
gender, rodent species, etc. These protective measures cannot be overempha-sized for the assurance of high-quality animals and successful experimentation
Dietary Restriction 355
2. Specific pathogen-free animals: A discussion similar to that of animal facility
conditions required to guard against possible infections can be extended to the
health status of the animals used in studies, particularly when employing long-term, chronic DR. Although the beneficial effects of DR are known to occur in
almost all laboratory rodents, even when housed in conventional, nonbarrier
facilities, researchers would be better served to use animals having specific patho-gen-free status (1,9). Experience has taught that prudent animal selection should
be made from strains having well-documented records that show the animal’s
growth patterns, metabolic responses, and pathologic status. This precautionary
measure gives an added advantage when used in combination with a specific
pathogen-free barrier. This is especially true when holding and maintaining ani-mals for a lifelong study to investigate aging processes or survival effects, during
which time the animals have an increased susceptibility to infection.
3. Selection of animal cages: Although not essential, shoebox type, plastic-bottom
cages that are fitted with a wire rack are recommended. The reason for choosing
this type of cage over wire-bottom cages relates to food spillage, among other
considerations. Food spillage is common to the use of wire-bottom cages and can
create inaccurate food consumption data. On the other hand, plastic-bottom cages
with raised wire-meshed floor racks allow food spillage recovery, which would
result in more accurate data.
4. Requirements for a balanced diet: A word of caution is need for the selection of
the chow used in DR studies. As practiced in the laboratory, DR is imposed on
test animals by a reduction of daily food allotments (usually 40%). It is therefore
of prime importance that the reduced amounts of diet contain all the required
nutrients, particularly those given during the growth period, to avoid any defi-ciencies. To ensure nutritional sufficiency, it is prudent to choose a well-defined,
semisynthetic diet (see Ta ble 1) from a reputable test chow manufacturer. As part
Table 1
Dietary Compositiona
Component Percentage
Dextrin 45.99
RP 101 soy protein 21.00
Sucrose 15.00
Corn meal 6.00
Mineral mix 5.00
Vitamin mix 3.30
Fiber 3.00
DL-Methionine .35
Choline chloride .35
aThis semisynthetic diet mixture is available from Purina Test Diets
(Richmond, IN, USA).
356 Yu
of their quality control procedures, investigators should periodically send sample
chows to independent laboratories for complete analysis. In some cases, research-ers carrying out DR studies should use chows fortified with additional vitamins
proportional to the reduced food amounts, which would ensure that the DR ani-mals receive a vitamin intake equal to that of the nonrestricted, control animals.
One other important practical matter concerning chow selection is the choice
of using powder or pellet form. Although more time consuming to use, the pow-der form is usually preferred because it permits more accurate weight measure-ments and less spillage. Pellet form, on the other hand, is likely to be spilled by
the animals because they tend to grab the pellets outside the food cup, which
results in even more food spillage.
3. Methods
3.1. Preparatory Procedures
1. Acclimatize animals to their new environment by caging them singly for 2 wk.
Although age of animals depends on the study design, shipped animals are typi-cally 1 mo old. Note that the younger the animal, the more robust the DR effect.
Also, specifying a body weight range is advantageous.
2. Initiate collection of animal body weights 2 d after arrival.
3. Allow all animals free access to water and choice of chow during the 2-wk accli-matization period.
4. Monitor and record food consumption during the 2-wk acclimatization period to
determine DR food allotments.
5. Select animals randomly to initiate DR regimen.
6. If periodic sacrifices for cross-sectional studies are planned, it is wise to place
more rats in the nonrestricted (i.e., ad libitum group) owing to the required shorter
life-span of these animals. This will ensure sufficient rats are available at senes-cence, a prudent measure to compensate for the higher mortality from ad libitum
feeding. A guideline for choosing the number of animals needed for survival and
longevity studies is given in Subheading 3.4., step 1.
7. For DR animals, initiate the regimen at the end of the 2-wk acclimatization period
with the set amounts of reduced chow. For the ad libitum fed, control animals,
continue to monitor and record food consumption twice weekly. The amount of
food consumed will serve as the basis for the next dietary allotment.
3.2. Optimal DR Levels
1. Because the DR paradigm can be executed in varying degrees (i.e., 10–40%),
researchers should first decide on the extent of restriction, as dictated by the study
design. A common observation is that the efficacy of DR is inversely related to
the extent of DR; that is, the greater the restriction, the more robust the DR effect
and the longer the life extension (10). The common practice is to implement 40%
DR (1).
2. As explained in Subheading 3.1., step 7, the chow consumption of the ad libitum
group is used as the basis to allot daily food amounts for the DR group. Thus, it is
Dietary Restriction 357
important to allow true  ad libitum  feeding. However, one word of caution is
needed: Ensure the chow in food cups is fresh, as rodents tend to consume less if
the food is stale, which would preclude accurate food consumption measure-ments, especially for small rodents.
3. Continually collect, usually twice weekly, the average amount of food consumed
from the ad libitum, control animals.
4. During the animal’s rapid growth phase, adjust food allotment for the DR ani-mals, twice weekly if necessary.
3.3. Animal Monitoring
An animal’s morbidity and mortality are profoundly influenced by its nutri-tional status. It is therefore imperative that the colony of experimental animals
be closely monitored as follows:
1. Check animal behavior and health conditions daily, paying close attention to
spontaneous movements and eating and drinking habits.
2. Inspect animals with increased frequency as they age (more than twice daily, if
3. Perform necropsy on all animals killed, fixing tissue specimens in a formulin
solution for a histological evaluation for the probable cause of death.
3.4. Longevity Studies: Survival Data and Analysis
The hallmark of DR is life extension. It is therefore important to carefully
collect and analyze survival data based on animal mortality.
1. Prior to the start of the study, consider the number of animals needed to meet the
statistical requirements of your study based on power function analysis.
2. If possible, allow animals to die naturally, using death as the endpoint for data
collection criterion.
3. To construct survival data, register each death, noting age in days.
4. Use data collected to calculate median life-span, 10% survival, and maximum
5. Another method used to analyze longevity data is to measure mortality rate dou-bling time (MRDT) base on the Gompertzian equation. Some investigators con-sider MRDT a more reliable expression of the rate of aging or “biomarker” of
aging (11). Although DR intervention has been show to increase MRDT, this
method has some inherent shortcomings as the Gompertzian derivation is based
on mortality rate (i.e., death) as an index of the rate of aging, which it is not in
4. Notes
4.1. Extent and Time of DR Intervention
Today, the DR paradigm is the most powerful and reproducible intervention
available to gerontologists to retard aging processes and suppress the patho-
358 Yu
genesis of major diseases  (1–4,9). When considering DR implementation,
investigators face the following three practical questions in evaluating its
maximum effect: How much (or little) restriction is need to show a discernible
life-span extension? When is the best time to initiate DR? How long should an
animal remain on the restricted diet? Brief comments regarding these important
questions are listed below:
1. Extent of DR: Consider these facts when deciding the extent of the DR to be imposed.
a. The effectiveness of DR is inversely related to the amount of caloric intake
(i.e., the greater the DR, the stonger the DR effect, the longer the life-span).
b. Beyond 40% restriction, which has become a routine procedure for some stud-ies, increases the risk of premature death among young animals.
2. The optimal timing for DR implementation: Based on the results of all studies
using DR, it is safe to say that the younger the animal when DR is imposed, the
more robust the outcome. In one of our studies (2) we began DR at 6 mo of age
(i.e., after sexual maturity). A reanalysis of the survival data from this study by
Neafsey (7) concluded that, although to varying degrees, DR exerts a life-pro-longing action irrespective of its initiation time. Another study (6) showed that
mice benefited from the effects of DR by a substantial life extension (~15%),
even when DR was initiated at 12 mo of age. The consensus is that, for many
reasons, the earlier DR is implemented, the more marked the results.
3. Length of DR: The rule of thumb is that the longer DR is imposed, the longer the
life-span extension. The success of DR is also dependent on time of initiation
(e.g., at young age or in adulthood). To date, all indications are that DR is more
effective when initiated during an animal’s youth or growth phase and continued
through adulthood than when simply carried out over the same length of time and
initiated during adulthood.
4.2. DR as a Mechanistic Probe
Because DR is the subject of widespread scientific inquiry (9,10), even out-side the field of gerontology, some comments are merited for those who are not
familiar with the DR paradigm. Below are some of its advantages as well as
1. Advantages: DR is shown to extend life-span by preventing most physiological
dysfunction and pathological changes. This ability provides researchers with an
important tool by which to uncover the mechanisms that underlie age-related
deterioration at various biological system levels, including the molecular events
involved in gene activity. This amazing aspect of the effectiveness of DR is high-lighted by the evidence showing its ability to modulate almost all the age-related
changes observed in ad libitum fed animals.
DR has also been used successfully to test several theories and hypothesis of
aging (3). One interesting recent test using the DR paradigm to substantiate the
“oxidative stress theory of aging” demonstrated its ability to attenuate oxidative
Dietary Restriction 359
damage and enhance antioxidant defense systems. If DR turned out not to have
the ability to modulate oxidative stress, then the basis as a bona fide theory of
aging would be weakened.
2. Limitations: A caveat should be added that the DR paradigm has a limitation that
critics refer to as its nonspecificity (i.e., lacking a specific action), because of the
broad modulation spectrum of DR in almost all biological systems. The broad
efficacy of DR should be expected, however, because the aging process itself is
broad and multifaceted. This limitation is therefore clearly outweighed by its
advantages, especially as studies move toward more molecular approaches and
the DR paradigm shows its usefulness in its ability to modulate genomic activity.
This work was supported, in part, by a grant from the National Institute on Aging
(AGA–01188). The author thanks Ms. Corinne Price for manuscript preparation.
1. Yu, B. P., ed. (1994) Modulation of Aging Processes by Dietary Restriction. CRC
Press, Boca Raton, FL.
2. Yu, B. P., Masoro, E. J., and McMahan, C. A. (1985) Nutritional influences on
aging Fischer 344 rats: 1. Physical, metabolic and longevity characteristics.
J. Gerontol. 40, 657–670.
3. Yu, B. P. (1996) Aging and oxidative stress: modulation by dietary restriction. Free
Radic. Biol. Med. 21, 651–668.
4. Weindruch, R. and Walford, R. L. (1988) The Retardation of Aging and Disease by
Dietary Restriction. Charles C Thomas, Springfield, IL.
5. Bertrand, H. A., Herlihy, J. T., Ikeno, Y., and Yu, B. P. (1999) Methods of assessing
aging processes: dietary restriction, in Methods in Aging Research (Yu, B. P., ed.),
CRC Press, Boca Raton, FL.
6. Weindruch, R. and Walford, R. L. (1982) Dietary restriction in mice beginning at
1 year of age: effect of life span and spontaneous cancer incidence. Science 215,
7. Neafsey, P. J. (1990) Longevity hormesis: a review. Mech. Ageing Dev. 51, 1–31.
8. Lane, M. A., Baer, D. J., Rumpler, W. V., Weindruch, R, Ingram. D. K., Tilmont, E.
M., Culter, R. G., and Roth, G. (1996) Calorie restriction lowers body temperature
in rhesus monkeys, consistent with postulated anti-aging mechanisms in rodents.
Proc. Natl. Acad. Sci. USA 93, 4159–4164.
9. Lewis, S. M., Leard, B. L., Turturro, A., and Hart, L. W. (1999) Long-term housing
of rodents under specific-pathogen free barrier conditions, in  Methods in Aging
Research (Yu, B. P., ed.), CRC Press, Boca Raton, FL.
10. Weindruch, R., Walford, R. L., Fligiel, S., and Guthrie, D. (1986) The retardation of
aging in mice by dietary restriction: longevity, cancer, immunity, and lifetime energy
intake. J. Nutr. 116, 641–654.
11. Finch, C. E. Pike, M. C., and Witten, M. (1990) Slow mortality rate accelerations
during aging in some animals approximate that of humans. Science 249, 902–905.
Genetically Engineered Mice 361
From: Methods in Molecular Medicine, Vol. 38: Aging Methods and Protocols
Edited by: Y. A. Barnett and C. R. Barnett   © Humana Press Inc., Totowa, NJ
The Use of Genetically Engineered Mice
in Aging Research
Julie K. Andersen
1. Introduction
The primary model systems used for studying the role of regulated gene
expression in senescence and the effects that genetic variations have on this
process have been to date either mammalian cells in vitro or invertebrate sys-tems such as yeast, C. elegans, and Drosophila. Both types of model systems
have been very useful in elucidating potential genetic pathways involved in the
aging process and are appealing owing to a variety of factors, not the least of
which is the ease of genetic manipulation. As such, they constitute a good start-ing point for understanding the phenomenon of aging. However, such models
have their limitations; no tissue culture system can yet approach the subtleties
of complex cell–cell interactions existing in the whole, intact animal, and
although significant evolutionary conservation has been found between many
genes in invertebrates and humans, they are still farther removed from humans
than available vertebrate model systems.
In the last 20 yr, rapid advances have been made in techniques allowing the
introduction of selected mutations into mammalian systems in vivo. The abil-ity to create laboratory mouse strains containing targeted mutations has allowed
the creation of vertebrate models to study the aging process and has extended
our understanding of many age-related diseases in humans.
2. Making a Transgenic Mouse: The Basics
In the mouse, transgenic technology is a process by which genetic engineer-ing at the embryonic stage is performed to allow elevated expression of existing
genes or ectopic expression of novel genes in the adult animal. This process
involves microinjection of a cloned gene of interest into the pronuclei of a
362 Andersen
mouse embryo at the one-cell stage (1). The transgenic DNA thus introduced
will randomly integrate into the genomic DNA of the embryo, where it can be
transcribed and translated into functional protein. Following injection of the
foreign DNA, the embryos are implanted into pseudopregnant recipient
females. Because the transgenic DNA integrates into the mouse’s genomic
DNA at the one-cell stage, the integrated DNA will be contained in the DNA of
all cells of the mouse’s body including its germ line cells and will therefore
become a heritable component of its genetic make-up. Following their birth,
founder animals are tested for presence of the transgene via the polymerase
chain reaction (PCR) or Southern blot analysis, and positive founders bred out
to create transgenic lines in which patterns and levels of expression of the gene
can be observed as well as its physiological consequences. The extent of expres-sion of the transgene appears to be more related to its site of genomic integra-tion as opposed to copy number. Many independent lines must be examined to
separate out the effects that the integration site vs the transgene itself have on
the observed phenotype.
3. Further Considerations in Transgene Design
Normally, the injected foreign DNA used for transgenic production is
designed to contain the gene of interest expressed under the control of a regu-latory promoter element that designates at what time and in what cell types the
expression of the transgene will occur. The injected DNA can be either the
entire gene itself including its native promoter element and all of its introns
and exons and 5′ and 3′ regulatory DNA sequences, or it may consist of a heter-ologous promoter driving expression of the cDNA that is missing the introns
and regulatory sequences. It has been demonstrated in previous studies that
genomic constructs are expressed more efficiently than cDNAs. However,
owing to the large size of some genes, it is not always possible to use the entire
gene although there have now been several reports of successful integration of
transgenic yeast artificial chromosome (YAC) DNAs of several hundred
kilobases in size into the mouse genome (2–7). It has been suggested that the
presence of the first intron is necessary for high levels of expression of a cDNA
even if the intron is a heterologous one (8). The choice of the promoter (native
vs heterologous) is normally dependent on when, where, and to what extent
expression of the gene product is desired.
4. Making a “Knockout” Mouse: The Basics
Although expression of an antisense mRNA in a transgenic construct has
been used in a few instances to reduce levels of a desired gene product (9–14),
the success of antisense RNA expression is dependent on several factors includ-ing the level of expression of the endogenous RNA and its turnover rate as well
Genetically Engineered Mice 363
as the rate of turnover of the protein product produced from it. However, in the
last decade a second type of genetic engineering called gene targeting or gene
“knockout” has been developed that allows the elimination of expression of an
endogenous gene of interest by producing an insertional mutation in the
endogenous gene. Here, a cloned fragment from the gene (a minimum of 2 kb
in size) is altered in vitro usually via insertion of a neomycin resistance (neoR)
gene into a region of the gene vital for its expression, for example, an exon
(15,16). Thymidine kinase sequences from the herpes simplex virus (HSV-tk)
are next introduced at both ends of the linearized gene fragment and the altered
gene (termed the “targeting vector”) is introduced into pluripotent embryo-derived stem (ES) cells either via either direct injection or electroporation (for
a review of ES culture conditions, see ref. 17). The most well-studied and com-monly used ES cells are derived from the inbred 1295v mouse strain, although
the use of ES cells from both inbred C57B1/6 and hybrid C57B1/6  × CBA/
JNCrj backgrounds has also been described (18–20).
Following introduction of the altered gene fragment into the ES cell, homo-logous recombination occurs between it and the endogenous gene in the ES
cell DNA at the ends of the region of homology between the transgene and
genomic target sequence so that the copy of the gene containing the neoR inser-tional mutation replaces the normal copy of the endogenous gene (14,15). Use
of DNA in the targeting vector that is derived from the same background strain
as the ES cells used may improve the frequency of homologous recombination,
that is, syngeneic or isogenic DNA (21).
During homologous recombination, the distal HSV-tk sequences are elimi-nated. When the transgene is inserted into the ES cell DNA via random integra-tion (a more frequent event than homologous recombination in mammalian
cells unlike in yeast), the HSV-tk sequences are retained. Cells containing the
homologously integrated copy of the gene can therefore then be selected by
growth in media containing both neomycin and gancyclovir to select for cells
containing the neoR insertion but not the HSV-tk sequences (“positive–nega-tive selection”). Other methods to distinguish homologous recombination from
random integration include using targeting vectors containing a positive selec-tion marker lacking either its own promoter or polyadenylation site (22).
ES cells containing the neoR gene insertion in the gene of interest are next intro-duced into mouse blastocysts by either injection into the cavity of a host blas-tocyst (a 3.5-d embryo at the 32-cell stage,  23) or aggregation with a host
embryo at the morula stage (2.5-d embryo at the 8–16-cell stage,  23–26).
Because they remain pluripotent even following long periods of growth in tissue
culture, the ES cells will contribute to all tissues of the developing animal
including cells of the germ line. ES cells derived from mice with one coat color
may be introduced into a host embryo of a different coat color to distinguish
364 Andersen
which resulting animals are chimeras, that is, made up of tissues from both ES
and host cells. Following implantation into foster mothers, the resulting chi-meric offspring are bred, producing mice that, if the ES cells have become part
of the germ line, are heterozygous for the introduced mutation. Heterozygous
animals are bred to obtain homozygous mutant mice.
5. Generation of a Particular Genetic Mutation via Gene Targeting
Sometimes, a specific subtle mutation in the endogenous gene is desired
rather than generation of a neoR insertional mutation. This requires a two-step
double gene replacement. First the mutant is made containing the neoR marker,
and then the neoR marker is replaced with another targeting vector containing
the desired mutation. Following selection, the ES cell clones can be checked
by PCR or Southern blot analysis to ascertain that they contain the correct
6. Special Factors to Consider When Making Genetically
Engineered Mice for Aging Studies
6.1. Adult-Specific Expression
In aging studies, the researcher is generally interested in the effects of alter-ations in a particular gene product in a particular tissue in the adult animal.
Many cell-type specific promoters are available that will allow the researcher
to target the particular tissue of interest; however, they often allow expression
prior to adulthood in a manner that may confound the researcher’s analysis of
aging effects related to changes in the activity of the gene product. To eliminate
any developmental effects of gene alteration, inducible systems can be used.
Previously, the only types of inducible systems available were those that
involved the use of inducing agents that themselves could be toxic to cells or
elicit pleiotrophic changes that could confound the analysis of any resulting
phenotype, for example, heat shock, heavy metals, glucocorticoids, and inter-feron (27–29). These systems were also plagued by low levels of inducibility,
leakiness in the noninduced state, and the inability to achieve effective induc-tion in vivo. In the last few years, new inducible systems have been developed
that overcome many, if not all, of the drawbacks of the earlier systems and
allow both temporal and cell specificity of gene expression.
The tetracycline (Tet) system allows inducible expression of transgenes
without the requirement of agents that might themselves cause cellular stress
(30). In this system, a Tet-inducible reverse transcriptional transactivator (rTta)
protein consisting of a “reverse” Tet repressor (rTetR) fused to the herpes sim-plex virus VP16 transcription activation domain is constitutively expressed via
a cytomegalovirus immediate early promoter (pCMV) from what has been
termed the “regulatory” transgene. Expression of this transgene can be limited
Genetically Engineered Mice 365
in both cell lines and transgenic animals by replacing the CMV promoter nor-mally used with one that is cell specific. The gene of interest is placed into a
second vector called the “response” transgene at a multiple cloning site located
downstream of a Tet response element (TRE) consisting of seven tandem Tet
operator sequences fused to a minimal CMV promoter (pminCMV). The com-ponents of this system are now commercially available from Clontech. The
rTta protein will bind and activate the gene of interest only in the presence of
Tet or its more lipophilic derivatives such as doxycyclin (dox). Both highly
inducible and tightly controlled levels of transgene expression are observed
upon treatment with inducing agent with little basal expression. The Tet induc-ible system has been recently used to generate transgenic mice containing
tetracycline-inducible transgenes in a cell-type specific manner including under
the control of neuronal-specific promoters indicating that dox administered
orally crosses the blood–brain barrier (31,32). In addition, oral dox treatment
seems to have no detrimental effects on body tissues or general toxicity for the
animals (33). The Tet system can be used to drive expression of antisense RNAs to
control the activity of endogenous gene expression  (34). Like with other
transgenic systems, the site of integration of both activator and response
transgenes may have an effect of their levels of expression.
In addition to the Tet system, other systems are also available that appear to
be effective in inducing both cell-specific and inducible transgene expression.
The recently described ecydysone system, for example, also boasts high levels
of inducibility, low basal expression, rapid kinetics, a minimum of confound-ing inducing agent-elicited effects, no apparently toxicity to mammalian cells
in vitro or in vivo, and the components are also now commercially available
from Invitrogen (35). This system requires the expression of two separate com-ponents of a modified nuclear Drosophila ecdysone receptor (the Drosophila
ecdysone receptor itself and a mammalian retinoic X receptor both can be
expressed from cell-specific promoters); these receptors bind to one another
and activate expression of the transgene of interest from an ecdysone respon-sive promoter upon administration of the steroid compound, muristerone.
Induction of transgene expression using this system was reported by the authors
to be greater than that obtained with the Tet system and to have lower levels of
basal expression; however, there has been only one publication to date in which
the system has been demonstrated to work in vivo (35).
Besides the availiability of systems for tissue-specific and inducible expres-sion of transgenes, systems have also been developed for the inducible inacti-vation of gene expression only in certain organs or cell types. Such systems
overcome the problems associated with the possible lethality associated with
disrupting genes during the embryonic period so that the effects of gene loss
can be analyzed at later stages in the life of the organism. One such system is
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based on the Cre–lox recombinase system in bacteriophage P1 (29,36,37). The
Cre recombinase protein will catalyze site-specific recombination between two
short segments of phage recombinase recognition sequence DNA within the
genome called loxP sites, removing any intervening sequences between them.
The targeting vector can therefore be designed to contain lox sites flanking a
portion of the endogenous gene that the researcher wishes to delete. The lox
sequence-containing mice can then be mated with transgenic mice containing
the cre recombinase gene behind a particular cell-specific promoter resulting
in an animal in which the fragment of the gene of interest is excised only in the
tissues in which the cre recombinase is expressed. The sites of the original lox
insertions in the targeting vector must be carefully chosen as to not disrupt
normal endogenous gene function. A second similar system involves the use
of a recombinase from yeast called flp that can remove DNA flanked by
34-basepair sequences called frt (38,39).
These systems can also be used to incorporate subtle alterations into a gene
of interest by introducing a targeting vector containing both the mutation and
at a separate location the neoR gene flanked by lox sequences into ES cells.
Cre recombinase can then be transfected into selected drug-resistant cells to
remove the selectable marker leaving only the mutation within the ES cell
genome (40).
The cre recombinase protein can be expressed in a tissue-specific and
inducible fashion by use of the Tet system. In this way, the researcher is able to
produce mice that when bred with  lox sequence-containing lines produce
progeny in which loss of function of the gene of interest is both inducible and
cell-specific (41).
6.2. Genetic Background
Another important factor to consider in the construction of genetically engi-neered mice for aging studies is the choice of genetic background. Genetic
background may have a significant effect on the expression phenotype of a
particular gene as there may be allelic modifiers in one genetic background vs
another that alter the phenotype of the modified gene’s expression, for example,
by suppressing or enhancing activity of another related gene with redundant
function (42).
Life-span studies done by crossing various inbred strains of mice have deter-mined that genetic background has a significant effect on longevity (43–45). A
current problem in the aging field as well as many others is that many people
perform their phenotypic analyses in hybrid rather than inbred or outbred
strains so it is unclear whether observed phenotypic effects can be attributed to
the specific genetic modification or are due to background effects (46). A com
mon solution to this problem is to measure the phenotypic effect in a large
Genetically Engineered Mice 367
number of animals, thereby increasing the probability that the difference
between the phenotype in the genetically altered mice vs control animals is
statistically significant and decreasing sampling error. Another solution is to
backcross into an inbred strain prior to phenotypic analysis. The most com-monly used inbred strain in aging studies is C57B1, which is often the
background of choice for many studies owing to its extensive characterization;
the 129 strain from which the original ES cells are often derived have unique
behavioral and neuroanatomical anomalies, are difficult to breed, and are prone
to a number of diseases (46,47). However, if an allelic modifier gene is linked,
that is, in close physical proximity to the genetically altered gene, then back-crossing into the desired strain will not be sufficient to alleviate the effects of
the modifier unless it is done numerous times, which can be both incredibly
time consuming and expensive. In terms of phenotype in knockout mouse
strains, one possible solution is to see if transgenic gene replacement negates
the effects of loss of gene function (47).
Lastly, it is also important to be aware that genetic alteration, such as admin-istration of pharmacological agents, can have pleiotropic effects leading to a
plethora of compensatory changes that result in secondary phenotypical alter-ations unrelated to the effects of alteration of expression in the original gene
product. Therefore, whatever gene is altered or background strain is used, it is
important to be aware that considerable care must be taken in intepreting the
results of genetic manipulation experiments in mice.
7. Examples of Genetically Engineered Mice Constructed
to Test Molecular Theories of Aging
Transgenic mouse lines with extended lifespans have not yet been reported.
However, likely candidates for such analyses based on studies in lower organ-isms such as Drosophila would include such genetically engineered models as
superoxide/catalase double transgenic lines and those containing metabolic
genes found to be important in dietary restriction paradigms in rodents (48).
Although the use of genetic engineering has yet to produce any longer-lived
mouse strains, engineered lines have been used to assess the validity of some
current popular molecular theories of aging including the roles of replicative
senescence and DNA damage and repair on longevity, examples of which
are described in the following subheadings.
7.1. Replicative Senescence
Fibroblasts cultured in vivo undergo a limited number of cell divisions before
they enter a senescent phase where they can remain for long periods of time
without undergoing further rounds of mitosis. This phenomenon is known as
replicative senescence (49). It has been suggested that the loss of proliferative
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capacity of cells is due either to random accumulation of cellular damage or to
a genetic preprogrammed process or innate “biological clock.”
Replicative senescence of fibroblasts can be prevented by expression of large
T antigen (T Ag) which allows proliferation to continue indefinitely as long as
T Ag is expressed. If it is removed, cells enter a postmitotic state. To test at
what point fibroblasts become dependent on T Ag for continued cell division,
embryonic fibroblasts were prepared from transgenic mice expressing a tem-perature-sensitive form of T Ag under the control of the interferon promoter
(50). When grown in the presence of interferon at 33°C, fibroblasts derived
from these animals become immortal. Removal of interferon or switching cul-tures to 39°C resulted in reduction of functional T Ag and cessation of cell
division. When T Ag levels were reduced while the cells were still dividing,
they did not lose proliferative potential. T Ag appeared to be required to
maintain cell division only once the normal mitotic life-span had elapsed. In
addition, when cells that were immortalized by switching them to growth in the
presence of T Ag were switched back to conditions of no T Ag, they underwent
only the exact number of cell divisions that would have been remaining if they
had not ever been exposed to T Ag. This suggests that fibroblast replication is a
regulated phenomenon and that the biological clock that limits mitotic life-span continues to operate in the presence of functional T Ag.
7.2. DNA Damage and Repair
Mice containing a targeted deletion of the Pms2 DNA mismatch repair gene
show a 1000-fold elevation in mutation frequency in all tissues examined com-pared to control animals; however, the presence of increased rates of mutagen-esis did not affect life-span (51). This suggests that DNA damage and repair is
not a limiting factor in longevity.
8. Examples of Genetically Engineered Mice Constructed
to Dissect Cellular Events Involved in Age-Related Diseases
The use of genetically engineered mice has been invaluable in aiding in the
understanding of a myriad of age-related diseases. Transgenics and knockouts
have provided us with several valuable animal models that have not only
allowed the analysis of disease progression but also the testing of new drug
therapies for various disorders associated with aging in the human population.
Some selected examples are described briefly in the following sections.
8.1. Alzheimer’s Disease
In the case of Alzheimer’s disease, several recent transgenic studies have
contributed greatly to our knowledge of how mutations in both the amyloid
precursor protein (APP) and the presenilin genes may contribute to the accu-
Genetically Engineered Mice 369
mulation of amyloid plaques and neuritic tangles in the cerebral cortex and the
hippocampus that eventually leads to the neurodegeneration and resulting cog-nitive decline associated with this disorder.
APP overexpressing transgenic lines have been constructed in which either
a shorter form of the human APP cDNA containing the same mutation found in
a familial Swedish Alzheimer’s pedigree was expressed behind the prion pro-tein promoter (PrP) or a longer form of the cDNA containing a separate muta-tion was expressed from the platelet-derived growth factor (PDGF) promoter
(52,53). In both cases, mice were found to develop the selective amyloid
plaques and gliosis in the hippocampus and cortex characteristic of the disease
by 6 to 12 mo of age. Plaque development appeared to be dependent on which
background strain the APP cDNA was expressed in, suggesting the presence of
allelic modifier genes (52). In the Hsiao line, a loss in cognitive ability as exem-plified by a deficit in various learning and memory tests was also reported,
although the loss was somewhat variable (52).
Transgenic mice expressing mutant human presenilin-1 (PS-l) cDNA under
control of the PDGF promoter exhibited an increase in β-amyloid (Aβ) deposi-tion, a product of abberrant APP processing that is found to be elevated in the
brains of Alzheimer’s disease patients and is considered a hallmark of the dis-ease (54). Amyloid deposits or behavioral deficits have yet to be reported in
these animals. Double transgenics expressing both mutant APP and PS-i cDNA
have an accelerated rate of amyloid deposition (55).
An important issue to be addressed with the lines described above is the
temporal relationship between the formation of amyloid deposits and the onset
of memory deficits in these animals (56). If cognitive effects occur prior to Aj3
deposition, this would imply that it is the soluble form of Aβ as opposed to the
fibrillary form that is responsible for disease pathology and that amyloid
plaques may occur coincidentally along with cognitive decline. Interestingly,
there appears to be no correlation between the amount of Aβ deposition and
degree of dementia in humans with the disease, and in addition transgenic mice
expressing the Swedish APP mutation in the FVB background show cognitive
effects in the absence of plaque formation (52,57).
8.2. Immunosenescence
Genetically engineered mice have also been used to study the phenomenon
of immunosenescence. Older people have an increased susceptibility to infec-tion as a result of an age-related decrease in the responsiveness of the immune
system to attack by foreign antigens such as bacteria and viruses. This immu-nodeficiency appears to be primarily due to a decline in the response of T cells
to receptor stimulation which in turn can result in a decline in both T-cell pro-liferation and alterations in cytokine secretion  (58). There appears to be an
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age-related shift in T-cell populations so that the number of T cells that express
a naive phenotype, that is, those T cells that have never encountered an antigen,
decreases while the number of memory T cells that have specificity toward a
particular antigen with age increases. It has been suggested that it is this decline
in the proportion of naive T cells that can respond to novel antigen that may
account for the age-related decrease in immune function or immunosenes-cence. To test this hypothesis, the function of T cells from a transgenic mouse
line expressing naive T cell receptor (TCR) were examined in both young
and aged animals (58). T cells expressing the naive TCR had decreased pro-liferative and secretion capacities in the older animals compared to those
in younger mice, suggesting that age-related deficiencies in immune func-tion may not be solely attributable to the switch in the proportion of naive vs
memory T cells but may also include a decrease in the response of naive
T cells in older individuals.
8.3. Atherosclerosis
Atherosclerosis is the major cause of mortality among elderly populations
in most highly developed nations. Genetically engineered mouse models devel-oped in the last several years which mimic this condition have contributed much
toward our understanding of the genetic and environmental factors involved in
susceptibility to this disease. For example, mice have been generated that are
deficient in apolipoprotein E (apoE), the surface component of lipoprotein par-ticles that is necessary for their recognition by lipoprotein receptors and there-fore their clearance from the bloodstream. These mice demonstrate delayed
clearance of lipoproteins and develop pathology characteristic of atherosclero-sis in an age-dependent manner including fatty deposits in the blood vessels
and plaque formation at the same types of vascular sites (i.e., arterial branch
points) seen in humans with this disorder. This condition was exacerbated by
feeding the mice a high-cholesterol, high-fat “Western” style diet  (59,60).
Replacement of apoE in these mice specifically in macrophages and blood ves-sels, however, appeared to be sufficient to decrease at least some of the disease
pathology (61).
Other atherosclerotic models have been generated in mice via transdominant
expression of mutant forms of the apoE protein such as the Leiden and R142C
mutations which have been demonstrated to cause altered clearance of lipopro-teins in humans (62,63). When fed a high-cholesterol diet, animals from both
these transgenic lines developed cardiovascular lesions including fatty depos-its and formation of fibrous plaques. Transgenic mice containing the human
apolipoprotein E*2(Arg-158  → Cys) mutation on an apoE null background
develop an even more severe hyperlipoproteinemia than the apolipoprotein
E*3–Leiden transgenic mice (64).
Genetically Engineered Mice 371
Other atherosclerotic models developed to date include mice deficient in
low-density lipid (LDL) receptors (65). In LDL-deficient animals, a high-fat
diet was found to have a greater effect on lesion size in male vs female animals,
suggesting a role for hormones in the initiation or progression of the disease.
Transgenic mice expressing the human apoB protein, the only protein compo-nent of LDL and a ligand for removal of this type of lipoprotein from the circu-lation, develop fatty lesions but only when fed a high-fat diet (66).
All of these models have provided insight into genetic modifiers and par-ticular environmental elements involved in lesion formation and disease pro-gression and are allowing this phenomenon to be teased out at the molecular
level. They should be extremely valuable in the testing of various candidate
drugs to combat the disease whose effects would be more expensive to investi-gate in currently more widely used rabbit or primate model systems.
8.4. Adult-Onset Diabetes
Adult-onset or type II diabetes is yet another age-related disorder in which
the advent of genetically engineered mice as models of the disease has been
extremely informative in understanding the molecular process involved in its
development. A major target of alterations in such experiments has been the
glucose transporters, a family of molecules involved in mediating the uptake of
glucose into cells with various tissue-specific expressions, glucose affinities,
and insulin sensitivities.
Glucose transporter 4 (glut 4) is the major molecule involved in insulin-induced transport of glucose into skeletal muscle and fat tissue when blood
glucose levels are high, for both use it as a primary energy source via glycoly-sis by these tissues and store it in the form of glycogen for leaner times (67).
Both glut 4 transgenics in which glut 4 levels were increased in relevant tis-sues, for example, muscle and glut 4 knockout mice have been generated to
examine what effects alterations in the levels of this molecule have on insulin
sensitivity and whole body metabolism. In one line of transgenics expressing
glut 4 specifically in muscle under the control of the myosin light chain
enhancer and promoter, fast-twitch muscle tissue was found to have increased
basal and insulin-stimulated glucose uptake (68). These animals also displayed
more rapid clearance of glucose from the bloodstream, decreased glucose lev-els after an oral glucose feeding, and increased basal and insulin-stimulated
whole body glucose utilization. Mice in which muscle-specific expression of
glut 4 was driven by the aldolase A promoter made diabetic by streptozotocin
adminstration were found to clear blood glucose more rapidly after insulin
injection than controls, which suggests that in the presence of elevated glut 4
levels, insulin resistance may be overcome even under diabetic conditions (69).
However, in these studies the mice were unable to significantly decrease levels
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of glucose after a high glucose load, which suggests that although elevation of
glut 4 in muscle cells reverses insulin resistance it is not sufficient to maintain
homeostatic blood glucose levels. Mice deficient in the glut 4 locus suprisingly
were able to maintain normal glycemic control and therefore appear to be able
to compensate for the loss of the glut 4 in some manner, perhaps via elevation
of other glucose transporters (70). However, these animals do develop a reduc-tion in adipose tissue and an enlargement of the heart muscle and die at 5–7 mo
of age. These animals will doubtless be valuable in studying the role of glut 4
in various tissues by replacing the molecule selectively in the knockout lines
with cell-specific promoters to see if this prevents the observed loss in these
animals. It would also be interesting to examine the effects of deletion of glut 4
later in life by making a conditional knockout animal; in the current knockout,
glut 4 expression was eliminated prior to birth during late embryogenesis.
9. Summary
In conclusion, genetically engineered animal models have been and will
continue to be invaluable for exploring the various factors involved in the basic
aging process as well as extending our understanding of many of the diseases
found to be more prevalent in the older human population. Further develop-ment of such in vivo systems will allow scientists to dissect further the role
genetic and environmental factors play in aging and in age-related disease states
and to enhance our understanding of these processes.
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2 comments on “Understanding Aging

  1. Vitamin for Hansen’s disease, hansen’s disease is a chronic infectious disease that primarily affects the skin, the peripheral nerves, the mucosa of the upper respiratory tract, and the eyes. Leprosy can lead to progressive permanent damage of these structures, and the resulting devastating disfigurement and disability has led to the historical social stigma and isolation (leper colonies) of those affected by the disease. The literature relating diet to leprosy is abundant between 1900 and 1960, peaking around 1940. Dietary factors that appear to influence the etiopathogenesis of Hansen’s disease include: walmart vitamin A, vitamin B group, vitamin C, vitamin D, vitamin E, calcium, and zinc. We noted a frequent lack of detailed dietary data in much of the literature cited. This is particularly true when the thrust of the investigation is not dietary. The literature strongly suggests the beneficial influence of adequate diet on the outcome of Hansen’s disease and the deleterious effect of a deficient diet. In contrast with the paucity of reported hard data in the previous reviews concerned with the effect of nutrition and diet on leprosy, is the increasing volume of literature reviews and experimental studies showing the profound impact of nutrition and diet on the immune system of man and laboratory animals. That diet has a global, if poorly understood, effect on the immune system is being increasingly recognized. The difficult question that remains is how to use this information in the control and prevention of disease. now vitamins Therefore, we believe that more emphasis should be given to diet in the study of this important worldwide disease in light of the current understanding of biochemistry and immunology. Historically speaking, leprosy has existed since at least 4000 BC, and the disease was present and described in the ancient civilizations of China, India, and Egypt. The first known written reference to the disease on Egyptian papyrus dates from about 1550 BC. It is believed that leprosy was brought to Europe by the Romans and the Crusaders and that later the Europeans brought it to the Americas. For centuries, leprosy remained a poorly understood disease characterized by human suffering and social isolation. In 1873, G.A. Hansen discovered the bacterial cause of this infectious disease. The first medication breakthrough occurred in the 1940s with the development of the drug dapsone, and later it was discovered that the bacteria which caused leprosy was more effectively killed by using multiple medications

  2. Hello would you mind letting me know which hosting company you’re utilizing? I’ve loaded your blog in 3 different browsers and I must say this blog loads a lot faster then most. Can you recommend a good web hosting provider at a honest price? Kudos, I appreciate it!

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