Principles and Practice of Endocrinology and Metabolism



Cloning of Genes
Genomic Libraries and Gene Isolation
Gene Amplification by Polymerase Chain Reaction
Variations of Polymerase Chain Reaction
Approaches to the Quantitative Assessment of Gene Expression
Transcription Assays
Messenger RNA Assays
Protein Expression Assays
DNA-Protein Interaction Assays
Genetic Manipulations in Animals in Vivo
Transgenic Approaches
Conditional (Developmental) Interruption of Gene Expression
Prospects for the Future for Conditional Transgene Expression
Expressed Sequence Tags
DNA Arrays for the Profiling of Gene Expression
Oligonucleotide Arrays (Genomic and Expressed Sequence Tags)
Complementary DNA Arrays (Specific Tissues)
Strategies for Mapping Genes on Chromosomes
Genetic Linkage Maps and Quantitative Trait Loci
Restriction Enzymes and Chromosomal Mapping
Physical Maps
Separating Chromosomes
Somatic Cell Hybridization
Chromosome Walking (Positional Cloning)
Candidate Gene Approach
Future Prospects
Human Genome Project
Stem Cells
Somatic Cell Cloning in Vivo
Gene Knock-Out Libraries
Gene Therapy: Vectors and Problems
Chapter References

The beginnings of molecular biology as a distinct discipline occurred in the late 1940s and early 1950s with the recognition that polynucleotides were the repository of genetic information in the form of DNA and the transmitters of genetic information in the form of messenger RNA (mRNA), and that transfer RNAs are fundamental for the assembly of amino acids into proteins. Detailed descriptions of the historical developments of this modern era of molecular biology are provided in several books.1,2,3 and 4 These were exciting times, as understanding progressed rapidly from the discovery by Avery and Brundage that DNA was a genetic substance; Chargaff established that DNA is composed of four different deoxyribonucleotides (dATP, dGRP, dTTP, dCTP); Watson and Crick elucidated the double-helical structure of DNA; Jacob and Monod identified mRNA as the intermediary in the transfer of information encoded in DNA to the assembly of amino acids into proteins; Holly discovered transfer RNAs; and Nirenberg et al. discovered the genetic code (i.e., each of the 21 amino acids is specified by a triplet of nucleotides, or codons, within the mRNA to be translated into a protein).
In the 1960s, several major discoveries paved the way for the development of recombinant DNA technology and genetic engineering. Two of the major breakthroughs that made this possible were the discoveries of reverse transcriptase5 and restriction endonucleases,6,7 and 8 and techniques for determining the precise sequence of nucleotides in DNA.9,10 Reverse transcriptase, which is found encoded in the RNA of certain tumor viruses, is the means by which the virus makes DNA copies of its RNA templates. It allows molecular biologists to copy mRNA into complementary DNA (cDNA), which is an essential step in the preparation of recombinant DNA for purposes of cloning.
Another fundamental discovery was that of restriction endonucleases, enzymes that cut DNA at specific sequences, typically of 4 to 10 base pairs. The application of specific restriction endonucleases allows for the cleavage of DNAs at precise locations, a property that is critical for the engineering of DNA segments.
A most critical and important discovery was the technologic methodology to determine the sequential order of nucleotides in DNA. Both chemical and enzymatic approaches were developed. Currently, the nucleotide sequences of DNAs are determined by sophisticated automated instruments using random enzymatic cleavages of DNAs labeled with fluorescent markers.
By fortunate coincidence, research into the mechanisms by which bacteria become resistant to certain antibiotics led to the discovery of bacterial plasmids, which are “viruses” that live within bacteria and lend genetic information to the bacteria to ensure their survival. Plasmids faithfully replicate within bacteria. Importantly, plasmid DNA is relatively simple in structure and is amenable to genetic engineering by excision of DNA sequences and insertion of foreign DNA sequences, which will replicate within bacteria without interference by the host bacterium. These plasmids have become useful vehicles in which to express and amplify foreign DNA sequences.
Complementary DNA Libraries. The cloning of a particular expressed gene begins with the preparation and cloning of cDNAs from mRNAs of a particular cell (Fig. 2-1; Table 2-1) (for a more comprehensive description, see reference 11 and reference 12). The cDNAs are prepared by priming the reverse transcription of mRNAs, using reverse transcriptase and short oligonucleotide fragments of oligodeoxyribothymidine, which preferentially bind to the 3′-polyadenylate, or poly(A), tract that is characteristic of cellular mRNAs. Alternatively, random oligonucleotides of different base compositions may be used. Double-stranded DNA is then prepared from the single-stranded cDNA by using DNA polymerase, and the cDNAs are inserted into bacterial plasmids that have been cleaved at a single site with a restriction endonuclease. To ensure a reasonably high efficiency of insertion of the foreign DNA into the plasmids, cohesive, or “sticky,” ends are first prepared by adding short DNA sequences to the ends of the foreign DNA and to the plasmids. Vectors that are commonly used are derivatives of the plasmid pBR322, which was engineered specifically for the purposes of cloning DNA fragments (see Fig. 2-1). Foreign DNA is inserted into a unique site that is prepared by endonuclease cleaving of a desired site within a polylinker, multiple cloning site engineered into the plasmid. This site is often located within the gene that codes for bacterial b-galactosidase. The backbone plasmid also carries a gene for resistance to ampicillin or tetracycline. Thus, bacteria containing the plasmids can be selected by their resistance to ampicillin or tetracycline; those specifically containing DNA inserts can be selected by their inability to express b-galactosidase and to cleave b-galactopyranoside (blue-white screening).

FIGURE 2-1. An approach used in construction and molecular cloning of recombinant DNA. A, Preparation of double-stranded DNA from an mRNA template. The enzyme reverse transcriptase is used to reversetranscribe a single-stranded DNA copy complementary to the mRNA primed with an oligonucleotide of polydeoxythymidylic acid hybridized to the poly(A) tract at the 3′ end of mRNA. A complementary copy of the DNA strand is then prepared with DNA polymerase. Ends of double-stranded DNA are made flush by cleavage with the enzyme S1 nuclease, and homopolymer extensions of deoxycytidine are synthesized on 3′ ends of DNA with the enzyme terminal transferase. Oligo(dC) homopolymer extensions form sticky ends for purposes of insertion of DNA into a linearized bacterial plasmid on which complementary oligo(dG) homopolymer extensions have been synthesized. B, Insertion of foreign DNA into a bacterial plasmid for molecular cloning. A bacterial plasmid, typically pBR322, that has been specifically engineered for purposes of cloning DNA is linearized by cleavage with restriction endonuclease Pst I. Poly(dG) homopolymer extensions are synthesized onto 3′ ends of plasmid DNA. Foreign DNA with complementary poly(dC) homopolymer extensions is hybridized to and inserted into the plasmid. Recombinant plasmid DNA is transfected into susceptible host strains of bacteria, in which plasmid replicates apart from bacterial chromosomal DNA. Bacteria are then grown on a plate containing tetracycline. Colonies that are resistant to tetracycline are tested for sensitivity to ampicillin. Because native plasmids contain genes encoding resistance to both tetracycline and ampicillin and the gene encoding resistance to ampicillin is inactivated by insertion of a foreign DNA at the Pst I site, bacterial colonies harboring plasmids with DNA inserts are resistant to tetracycline and sensitive to ampicillin. Subsequent screening of tetracycline-resistant, ampicillin-sensitive clones containing specific DNA-inserted sequences is carried out by either DNA hybridization with labeled DNA probes or by other techniques such as hybridization arrest and cell-free translation.

TABLE 2-1. Approaches for the Selection of Cloned Complementary DNAs (cDNAs)

Hybridization Screening. The recombinant plasmids containing DNA sequences that are complementary to the specific mRNAs of interest are identified by hybridizing recombinant plasmids to the initial mRNA preparations used in the cloning. The hybrid-selected mRNA is subsequently eluted and translated in a cell-free system appropriate for the protein under study. Alternatively, specific inhibition of the translation of an mRNA can be used to identify the DNA of interest: DNA that is complementary to the mRNA being translated will bind the RNA, thus precluding translation and reducing the amount of the protein being synthesized.
The initial techniques of hybridization selection and hybridization arrest, in which cell-free translation is used as the assay system, are now supplanted by hybridization of the bacterial colonies with synthetic oligonucleotide probes that are labeled with phosphorus-32 (32P). Mixtures of oligonucleotides in the range of 14 to 17 bases are prepared that are complementary to the nucleotide sequences predicted from the known amino-acid sequences of segments of the protein encoded by mRNA. Because of the degeneracy in the genetic code (there are 61 amino-acid codons and 20 amino acids), mixtures of from 24 to 48 oligonucleotides ordinarily represent all possible sequences complementary to a particular 14- to 17-base region of mRNA.
Expression Screening. Later-generation cDNA libraries have been prepared in bacterial phages (l gt-11) or hybrids between plasmids and phages (phagemids), which have been engineered to allow the bacteria infected with the recombinant phages to translate mRNAs expressed from the cDNAs, and thereby to produce the protein products encoded by the cDNAs. The desired sequence of interest can be selected at the protein level by screening the library of bacterial clones with an antiserum directed to the protein. When the desired product is a DNA-binding protein, the library can be screened with a labeled DNA duplex containing copies of the target sequence to which the protein binds.
Yeast Two-Site Interaction Trap. The cloning of cDNAs encoding proteins that interact with other known proteins can be accomplished using the yeast two-site interaction trap, which functions much as a bait and fish system. The bait is a cDNA encoding a known protein that is engineered to bind to an enhancer in the promoter of a gene that encodes a factor essential for the survival of a yeast cell. The sequences (fish) in the cDNA library are engineered with a strong transcriptional transactivation domain, such as that from the herpes simplex virus and yeast transcription factors VP16 or Gal-4, respectively. The occurrence of proteinprotein interactions between the bait and one of the fish activates the expression of the yeast survival gene, which thereby allows for the selection and cloning of the yeast cell that harbors the described cDNA sequence from the cDNA library.
Rapid Amplification of Complementary DNA Ends. Most often cDNAs isolated by one or more of the approaches described above lack the complete sequence and are deficient in the 5′ ends. The 5′ sequences are determined by using the rapid amplification of cDNA ends (RACE) technique.
Southern Blots and Hybridization Screening. The techniques used in the cloning of genomic DNA are similar to those used for cloning cDNA, except that the genomic sequences are longer than the cDNA sequences and different cloning vectors are required. The common vectors are derivatives of the bacteriophage l that can accommodate DNA fragments of 10 to 20 kilobases (kb). Certain hybrids of bacteriophages and plasmids, called cosmids, can accommodate inserts of DNA of up to 40 to 50 kb. Even larger segments of DNA up to 1 to 2 megabases (Mb) can be cloned and propagated in yeast and are called yeast artificial chromosomes (YACs). In the cloning of genomic DNA, restriction fragments are prepared by partial digestion of unsheared DNA with a restriction endonuclease that cleaves the DNA into many fragments. DNA fragments of proper size are prepared by fractionation on agarose gels and are ligated to the bacteriophage DNA. The fragments of DNA containing the desired sequences can be detected by hybridization of a membrane blot prepared from the gel with a 32P-labeled cDNA, a Southern blot. The recombinant DNA is mixed with bacteriophage proteins, which results in the production of viable phage particles. The recombinant bacteriophages are grown on agar plates covered with growing bacteria. Then the bacteria are infected by a phage particle, which lyses the bacteria to form visible plaques. Specific phage colonies are transferred by nitrocellulose filters and are hybridized by cDNA probes labeled with 32P, similar to a Southern blot. Libraries of genomic DNA fragments and tissue-specific cDNAs from various animal species cloned in plasmids and bacteriophages are available from a number of commercial laboratories. The development of yeast chromosomal libraries that harbor large segments (several megabases) of chromosomal DNA has markedly accelerated the generation of gene linkage maps.
Enhancer Traps. One approach to identifying novel genes imbedded in the genome is to randomly insert a transcriptional reporter gene into chromosomal DNA that has been cleaved into 1- to 2-kb fragments by digestion with a restriction endonuclease. The family of ligated hybrid fragments is then cloned into plasmids that are individually introduced (transfected) into host cell lines (e.g., NIH or BHK fibroblasts). After the transfected cell lines are incubated with the cloned DNA fragments for 1 to 2 days, extracts are prepared from the cells and assayed for expression of the transcriptional reporter gene. Typical transcriptional reporter genes used are firefly luciferase, bacterial chloramphenicol acetyl transferase, or bacterial alkaline phosphatase. When, by chance, a transcriptional enhancer is encountered, as determined by the activation of the reporter gene, the particular cloned DNA fragment is sequenced and searched for transcribed exonic and/or intronic sequences of genes, many of which typically reside 100 to 1000 base pairs from the enhancer sequence. The transcribed sequences of genes usually, but not always, reside 3′ (downstream) from enhancer sequences.
Rapid Amplification of Genomic DNA Ends. The principle of rapid amplification of genomic DNA ends (RAGE) is similar to that of RACE previously described and allows for the identification of unknown DNA sequences in genomic DNA. Oligonucleotide primers (amplimers) are annealed to the test genomic DNA sample and extended on the genomic DNA template with DNA polymerase, and a second set of oligonucleotide primers is ligated to the extended ends. The extended DNA fragments are then amplified by polymerase chain reaction (see next section), isolated by electrophoresis on agarose gels, and sequenced.
The development of the polymerase chain reaction PCR, a technique for the rapid amplification of specific DNA sequences, constituted a major technological breakthrough.13,14,15 and 16 This procedure relies on the unique properties of a thermally stable DNA polymerase (Taq polymerase) to allow for sequential annealing of small oligonucleotide primers that bracket a DNA sequence of interest; the result is successive synthesis of the DNA strands. Specific DNA sequences as short as 50 and as long as several thousand base pairs can be amplified over a million-fold in just a few hours by using an automated thermal cycler. The technique is so sensitive that DNA (genomic DNA or cDNA reverse-transcribed from RNA) from a single cell can be so amplified. Indeed, a sample containing only a single target DNA molecule can be amplified. The applications of this technique are diverse. Not only is it possible to amplify and to clone rare sequences for detailed studies, but also the technique has applications in the fields of medical diagnosis and forensics. Scarce viruses can be detected in a drop of serum or urine or a single white blood cell. Genotyping can be done from a blood or semen stain, saliva, or a single hair. Paradoxically, a major drawback of PCR is its exquisite sensitivity, which leaves open the possibility of false-positive results because of minute contaminations of the samples being tested. Thus, extreme precautions must be taken to avoid the introduction of contaminants.
PCR is carried out using DNA polymerase and oligonucleotide primers complementary to the two 3′ borders of the duplex segment to be amplified. The objective of PCR is to copy the sequence of each strand between the regions at which the oligonucleotide primers anneal. Thus, after the primers are annealed to a denatured DNA containing the segment to be amplified, the primers are extended using DNA polymerase and the four deoxynucleotide triphosphates. Each primer is extended toward the other primer. The result is a double-stranded DNA (which itself is then denatured and annealed again with primer, and the DNA polymerase reaction is repeated). This cycle of steps (denaturation, annealing, and synthesis) may be repeated 60 times. At each cycle, the amount of duplex DNA segment doubles, because both new and old DNA molecules anneal to the primers and are copied. In principle (and virtually in practice), 2n copies (where n = number of cycles) of the duplex segment bordered by the primers are produced.
The heat-stable polymerase isolated from thermophilic bacteria (Thermophilus aquaticus), Taq polymerase, allows multiple cycles to be carried out after a single addition of enzyme. The DNA, an excess of primer molecules, the deoxynucleotide triphosphates, and the polymerase are mixed together at the start. Cycle 1 is initiated by heating to a temperature adequate to assure DNA denaturation, followed by cooling to a temperature appropriate for primer annealing to the now-single strands of the template DNA. Thereafter, the temperature is adjusted for DNA synthesis (elongation) to occur. The subsequent cycles are initiated by again heating to the denaturation temperature. Thus, cycling can be automated by using a computer-controlled variable-temperature heating block.
In addition to permitting automation, the use of the DNA polymerase of T. aquaticus has another advantage. The enzyme is most active between 70° and 75°C. Base pairing between the oligonucleotide primers and the DNA is more specific at this temperature than at 37°C, the optimal functioning temperature of Escherichia coli DNA polymerase. Consequently, the primers are less likely to anneal nonspecifically to unwanted DNA segments, especially when the entire genome is present in the target DNA.
Simple modifications of the PCR conditions can expand the opportunities of the PCR. For example, synthesizing oligonucleotide primers that recognize domains (motifs) shared by cDNAs and their respective protein products, and choosing less stringent annealing conditions for the primers, permit new sequences of yet unknown DNAs to be generated with PCR, ultimately resulting in the discovery of new cDNAs belonging to the same family. For example, the pancreatic B-cell transcription factor IDX-1 was identified by PCR using oligonucleotide primers that would anneal to sequences shared by the homeodomain transcription factor family.
PCR primers can be modified in their sequence and thus are not completely complementary to the template DNA. The amplified PCR product then carries the sequence of the primer and not the original DNA sequence. This strategy can be used to insert mutations site-specifically into known DNA sequences.
Nuclear Run-On Assays. Several assays are available that provide an index of relative rates of gene transcription (Fig. 2-2). A simple, straightforward assay is the nuclear run-on assay in which nuclei are isolated from tissue culture cells and nascent RNA chains are allowed to continue to polymerize in the presence of radiolabeled deoxyribonucleotides in vitro. This assay has the advantage that it surveys the density of nascent transcripts made from the endogenous genes of cells and, on average, is a good measure of gene transcription rates in response to the existing environmental conditions in which the cultured cells are maintained. Newly synthesized RNA is applied (hybridized) to a nylon membrane on which a cDNA target complementary to the desired RNA has been adsorbed. Radiolabeled RNA hybridized to the cDNA is determined in a radiation counter.

FIGURE 2-2. Approaches to the quantitative assessment of gene expression. Shown are the various types of assays that can be used to examine regulation of gene expression at various levels. (mRNA, messenger RNA; RNase, ribonuclease; RT-PCR, reverse transcription polymerase chain reaction.)

Cell-Free In Vitro Systems. Rates of RNA synthesis can also be determined in broken cell or cell-free lysates to assess the relative strengths of different promoters. To restrict the newly synthesized radiolabeled RNA to a single size and, thus, to enable more ready detection by electrophoresis, a DNA template is used that does not contain guanine bases, called a G-free cassette. RNA synthesis is carried out in the absence of the guanine nucleotide. After synthesis of a specified length of RNA at the end of which guanine bases are encountered, RNA synthesis is terminated.
Transfection of Promoter-Reporters in In Vivo Cell Culture. Many of the currently used assays of gene transcription employ promoter sequences fused to genes encoding proteins that can be quantitated by bioassays (e.g., bacterial chloramphenicol acetyl transferase, firefly luciferase, alkaline phosphatase, or green fluorescent protein). The hybrid DNAs, so called promoter-reporter DNAs, are introduced into tissue culture cells by one of several chemical methods (i.e., DNA adsorbed to calcium phosphate precipitates, diethylaminoethyl (DEAE)-dextran incorporated into liposomes, or human artificial chromosomes [Table 2-2]); or physical methods (i.e., electroporation, direct microinjection of DNA, or ballistic injection using a gene gun [Table 2-3]). After introduction of the reporter DNA into the cells, the transfected cells are incubated for a specified time under the desired experimental conditions, the cells are harvested, and extracts are prepared for assays of the reporter-specific enzymatic activity. By these transfection methods, cell-type specificity for the expression of genepromoter sequences can be determined by comparing promoter-reporter efficiencies in cells of different phenotypes. In addition, important transcriptional control sequences in the promoter can be mapped by DNA mutagenesis studies.

TABLE 2-2. Chemical Methods for Introducing Genes into Mammalian Cells

TABLE 2-3. Physical Methods for Introducing Genes into Mammalian Cells

Transfection of Transcription Factor Expression Vectors. An extension of the promoter-reporter transfection approach is to cotransfect recombinant expression plasmids encoding transcription factors that bind to control sequences in the promoter DNA and activate transcription of the reporter. By this approach, critical functional components of transcription factors and critical bases in DNA control sequences can be examined experimentally.
Transgenic In Vivo Mouse Models. A method developed for examining specificity of tissue expression and efficiency of expression of promoter-reporter genes is their introduction into mice in vivo, using transgenic technology (see the section Genetic Manipulations in Animals In Vivo). Recombinant promoter-reporter genes are injected into the pronucleus of fertilized mouse ova and implanted into surrogate females. The tissues of transgenic neonatal mice are examined for the tissue distribution and relative strength of the expression of the reporter function. Commonly used reporter functions are the genes encoding either b-galactosidase or green fluorescent protein.
Northern Blot Hybridization. RNA blotting (Northern blotting) is analogous to DNA blotting (Southern blotting). RNA is separated according to size by electrophoresis through agarose gels. Generally, the electrophoresis is performed under conditions that denature the RNA so that the effects of RNA secondary structure on the electrophoretic mobility of the RNAs can be minimized. Alkaline conditions are unsuitable; therefore, agents such as glyoxal, formaldehyde, or urea are used. The size-separated RNA is transferred by blotting to an immobilizing membrane without disturbing the RNA distribution along the gel. A labeled DNA is then used as a probe to find the position on the blot of RNA molecules corresponding to the probe. The immobilized RNA is incubated with DNA under conditions allowing annealing of the DNA to the RNA on the immobilized matrix. After washing away excess and unspecifically annealed DNA, the matrix is exposed to an x-ray film to detect the position of the probe. RNA blotting allows the estimation of the size of the RNA that is being detected. In addition, the intensity of the band on an x-ray film indicates the abundance of the RNA in the cell or tissue from which the RNA was extracted.
Solution Hybridization Ribonuclease Protection. To obtain more precise information on the amount of a specific RNA species in a certain cell or tissue, a single-stranded radioactive probe is generated that is complementary to a portion of the RNA being studied. An excess amount of this single-stranded probe is then mixed in solution with the total RNA of the cells or tissue being investigated. Digestion with ribonuclease of all single-stranded nucleic acids present after hybridization leaves the double-stranded species, consisting of the labeled probe annealed to its complementary RNA, in the solution. The contents of the solution are then size-separated on an electrophoretic gel, which is exposed to an x-ray film. Knowing the amount of input labeled single-stranded probe allows a quantification of the specific RNA present in the total RNA of the cells or tissue.
In Situ Hybridization. In situ hybridization with labeled single-stranded probes onto tissues is, in principle, similar to the ribonuclease protection assay. Detection and determination of the location of a certain species of RNA within a tissue is possible.
Reverse Transcription Polymerase Chain Reaction. Reverse transcription polymerase chain reaction (RT-PCR) can be used to quantitate the abundance of a specific RNA. This method is particularly practical when small amounts of tissue or cells are available to be analyzed. The RNA is reverse-transcribed to DNA with reverse transcriptase. The cDNA population is then subjected to PCR amplification with specific primers that recognize the cDNA in question. By choosing the number of PCR cycles within the linear range of product generated after each cycle (i.e., enough primers, nucleotides, and DNA polymerase in the reaction mixture for none of them to be the limiting factor of the reaction) and adding to the PCR reaction a defined amount of an artificial DNA template that is also recognized by the primer oli gonucleotides but yields a differentsized product, one can detect differences in abundance of cDNA (and hence RNA in the original sample) among two or more samples. Newer methods allow for an on-line monitoring of each PCR reaction of the product generated. This is achieved by using primer oligonucleotides that can be monitored during the PCR reaction cycles by external optical devices. Such on-line continuous monitoring allows the performance of PCR reactions without prior determination of the number of cycles required to keep the PCR reaction within the linear range of amplification. Continuous PCR monitoring provides immediate information on abundance of a given cDNA species in PCR reactions. Knowledge of the absolute amount of labeled oligonucleotide primer added to the PCR reaction at the start can be used to determine the exact amount of the PCR product generated.
Cell-Free Translation. A commonly used method to analyze proteins encoded by mRNA is to translate the mRNA in cell-free translation systems in vitro. By this method, proteins can be radioactively labeled to a high specific activity. The cell-free translation also provides the primary protein product, such as a proprotein or prohormone, encoded by the mRNA.
Pulse and Pulse-Chase Labeling. Studies of protein syntheses can also be carried out in vivo by incubation of cultured cells or tissues with radioactive amino acids (pulse labeling). Posttranslational processing (e.g., enzymatic cleavages of prohormones) can be assessed by first incubating the cells or tissues for a short time with radioactive amino acids and then incubating them for an additional period with unlabeled amino acids (pulse-chase labeling).
Western Immunoblot. Another approach to the analyses of particular cellular proteins is the Western immunoblot technique. Proteins in cell extracts are separated by electrophoresis on polyacrylamide or agarose gels and transferred to a nylon or nitrocellulose membrane, which is then treated with a solution containing specific antibodies to the protein of interest. The antibodies that are bound to the protein fixed to the membrane are detected by any one of several methods, such as secondary antibodies tagged with radioisotopes, fluorophores, or enzymes.
Immunocytochemistry. A refinement of the Western immunoblot technique is the detection of specific proteins within cells by immunocytochemistry (immunohistochemistry). Cultured cells or tissue sections are fixed on microscope slides and treated with solutions containing specific antibodies. The antibodies that are bound to the proteins within the cells are detected with fluorescently tagged secondary antibodies or by an avidin-biotin complex. Immunocytochemistry is a powerful technique when used for the simultaneous detection of two or even three different proteins with examination by confocal microscopy.
The binding of proteins such as transcription factors to DNA sequences is commonly done by two approaches: electrophoretic mobility shift assay (EMSA) and Southwestern blotting. Typically, EMSA consists of incubation of protein extracts with a radiolabeled DNA sequence or probe. The mixture is then analyzed by electrophoresis on a nondenaturing polyacrylamide gel, followed by autoradiography or autofluorography to evaluate the distribution of the radioactivity or fluorescence in the gel. Interactions of specific proteins with the DNA probe are manifested by a retardation of the electrophoretic migration of the labeled probe, or band shift. The EMSA technique can be extended to include antibodies to specific proteins in the incubation mixture. The interaction of a specific antibody with a protein bound to the DNA probe causes a further retardation of migration of the DNA-protein complex, leading to a super shift.



  1. Biology laboratory, with emphasis on the techniques for large scale manual are referred back to the original literature.

  2. He mixture is then analyzed by electrophoresis on its temperature all other machine with a independent consists of incubation of protein extracts with a radio labeled leading to a super shift.

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