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.
A number of different assays are used to determine and evaluate protein-protein interactions. Two in vitro assays are coimmunoprecipitation and polyhistidine-tagged glutathione sulfonyl transferase (GST) pull-down. Two in vivo assays are the yeast and mammalian two-site interaction assays.
Coimmunoprecipitation. The commonly used coimmunoprecipitation assay makes use of antisera to specific proteins. In circumstances in which two different proteins, A and B, associate with each other, an antiserum to protein A will immunoprecipitate not only protein A, but also protein B. Likewise, an antiserum to protein B will coimmunoprecipitate proteins B and A. In practice, the proteins under investigation are radiolabeled by synthesis in the presence of radioactive amino acids, either in cell-free transcriptiontranslation systems in vitro, or in cell culture systems in vivo. Coimmunoprecipitated proteins are detected by gel electrophoresis and autoradiography. Alternatively, the proteins so immunoprecipitated or coimmunoprecipitated can be assayed by Western immunoblot techniques.
Glutathione Sulfonyl Transferase Pull-Down. GST is an enzyme that has a high affinity for its substrate, glutathione. This property of high-affinity interactions has been exploited to develop a cloning vector plasmid encoding GST and containing a polylinker site that allows for the insertion of coding sequences for any protein of interest. Thus, if protein A is believed to interact with protein B, the coding sequence for either protein A or protein B can be inserted into the GST vector. The GST–protein A or B fusion protein is synthesized in large amounts by multiplication and expression of the plasmid vector in bacteria. The GST-fusion protein is then incubated with either labeled or unlabeled proteins in extracts of cells or nuclei. Proteins in the extracts bound to protein A or B in the GST-fusion protein are pulled down from the extracts by capturing the GST on glutathione-agarose beads. Proteins are released from the beads and analyzed by either gel electrophoresis and autoradiography (labeled proteins) or by Western immunoblot (unlabeled proteins). Similar methods using polyhistidine tag in place of GST are also used for pull-down experiments.
Far Western Protein Blots. A variation on the Western blotting technique is the Far Western blot. In this technique, a radio-labeled or fluorescence-labeled known protein (instead of an antibody) is applied to a membrane to which proteins from an electrophoretic gel have been transferred. If the known protein binds to any one or more proteins on the membrane, it is detected as a labeled band by autoradiograph or autofluorography. Relatively strong protein-protein interactions are required for this approach to succeed.
Yeast and Mammalian Cell Two-Site Interaction Traps. The two-site interaction traps are useful for demonstrating protein-protein interactions in vivo. The principle of the approach is that, when a specific protein-protein interaction occurs, it reconstitutes an active transcription factor which then activates the transcription of a reporter gene. The cells (yeast or mammalian) are programmed to constitutively express a strong DNA-binding domain, such as Gal-4, fused to the expression sequence of the selected protein, protein A (the bait). The cells are also programmed to express a transcriptional reporter (e.g., CAT or luciferase linked to a promoter) that has binding sites for Gal-4. Thus, protein A anchors to the DNA-binding site of the reporter promoter via the Gal-4 binding domain but does not activate transcription of the reporter gene, and no reporter function is expressed. Protein B, however, is expressed as a fusion protein with a strong transcriptional activator sequence (e.g., the transcriptional transactivation domain of Gal-4 or of VP16). This transcriptional activation domain–protein B fusion protein does not bind DNA, but when, or if, protein B physically interacts with (binds to) protein A, a fully active transcription factor is reconstituted, the promoter reporter gene is transcribed, and the reported function is expressed.
The yeast two-hybrid system can be used to clone proteins that interact with a bait protein such as protein A. In this instance, a cDNA protein expression library is prepared or obtained that has all of the cDNAs of a given tissue fused to a coding sequence for a transactivation domain (e.g., VP16). Further, the reporter consists of a survival factor essential for the growth of the yeast cell. Thus, when a cDNA encodes a protein B (fish) that interacts with the bait protein A, the yeast cell expresses the survival protein and survives, whereas the other yeast cells die.
To create transgenic mice, DNA is injected into the male pronucleus of one-cell mammalian embryos (fertilized ova) that are then allowed to develop by insertion into the reproductive tract of pseudopregnant foster mothers (Fig. 2-3A). The transgenic animals that develop from this procedure contain the foreign DNA integrated into one or more of the host chromosomes at an early stage of embryo development. As a consequence, the foreign DNA is generally transmitted to the germline, and, in a number of instances, the foreign genes are expressed. Because the foreign DNA is injected at the one-cell stage, a good chance exists that the DNA will be distributed among all the progeny cells as development proceeds. This situation provides an opportunity to analyze and compare the qualitative and quantitative efficiencies of expression of the genes among various organs. The technique is quite efficient; >50% of postinjection ova produce viable offspring, and, of these, ~10% efficiently carry the foreign genes. In the transgenic animals, the foreign genes can be passed on and expressed at high levels in subsequent generations of progeny.

FIGURE 2-3. Approaches for (A) the integration of a foreign gene into the germline of mice, and (B) disruption or knock-out of a specific gene. A, DNA containing a specific foreign gene is microinjected into the male pronucleus of fertilized ova obtained from the oviduct of a mouse. Ova are then implanted into the uterus of pseudopregnant surrogate mothers. Progeny are analyzed for the presence of foreign genes by hybridization with 32P-labeled DNA probe and DNA prepared from a piece of tail from a mouse, which has been immobilized on a nitrocellulose filter (tail blots). B, To create a knock-out of a gene, pluripotential embryonic stem (Es) cells are used in vitro to introduce an engineered plasmid DNA sequence that will recombine with a homologous gene that is targeted. The recombination excises a portion of the gene in the ES cells, rendering it inactive (no longer expressible). ES cells in which the homologous recombination occurred successfully are selected by a combined positive-negative drug selection. The engineered ES cells are injected into the blastocoele of 3.5-day blastocysts that are then implanted into the uterus of pseudopregnant mice. The offspring are both chimeric and germline for expression of the knock-out gene and must be cross-bred to homozygosity for the genotype of a complete knock-out of the gene that is targeted for disruption.

Transgenic approaches can also be used to prevent the development of the lineage of a particular cell phenotype or to impair the expression of a selected gene. A cell lineage can be ablated by targeting a microinjected DNA containing a subunit of the diphtheria toxin to a particular cell type, using a promoter sequence specifically expressed in that cell type. The diphtheria toxin subunit inhibits protein synthesis when expressed in a cell, thereby killing the cell. The expression of a particular gene can be impaired by similar cell promoter– specific targeting of a DNA expression vector to a cell that produces an antisense mRNA to the mRNA expressed by the gene of interest. The antisense mRNA hybridizes to nuclear transcripts and processed mRNAs; this results in their degradation by double-stranded RNA–specific nucleases, thereby effectively attenuating the functional expression of the gene. The efficacy of the impairment of the mRNA can be enhanced by incorporating a ribozyme hammerhead sequence in the expressed antisense mRNA so as physically to cleave the mRNA to which it hybridizes. Another approach to producing a particular gene loss of function is to direct expression of a dominant negative protein (e.g., a receptor made deficient in intracellular signaling by an appropriate mutation, or a mutant transcription factor deficient in transactivation functions but sufficient for DNA binding). These dominant negative proteins compete for the essential functions of the wild-type proteins, resulting in a net loss of function.
Another approach, termed targeted transgenesis, combines targeted homologous recombination in embryonic stem (ES) cells with gain-of-function transgenic approaches.11,12,17 This method allows for targeted integration of a single-copy transgene to a single desired locus in the genome and thereby avoids problems of random and multiple-copy integrations, which may compromise faithful expression of the transgene in the conventional approach.
A major advance beyond the gain-of-function transgenic mouse technique has been the development of methods for producing loss of function by targeted disruption or replacement of genes. This approach uses the techniques of homologous recombination in cultured pluripotential ES cells, which are then injected into mouse blastocysts and implanted into the uteri of pseudopregnant mice (Fig. 2-3B). The targeting vector contains a core replacement sequence consisting of an expressed-cell lethal-drug resistance gene (selectable marker) (e.g., neomycin [Pgk-neo]) flanked by sequences homologous to the targeted cellular gene, and a second selectable marker gene (e.g., thymidine kinase [pgk-tk]). The ES cells are transfected with the gene-specific targeting vector. Cells that take up vector DNA and in which homologous recombination occurs are selected by their resistance to neomycin (positive selection). To select against random integration, a susceptibility to killing by thymidine kinase (negative selection) is used; only homologous recombination in which the thymidine kinase gene has been lost will confer survival benefit. Because the ES cells are injected into multicellular 3.5-day blastocysts, many of the offspring are mosaics, but some are germline heterozygous for the recombined gene. F1 generation mice are then bred to homozygosity so as to manifest the phenotype of the gene knock-out. Using this approach of targeted gene disruption, literally thousands of knock-out mice have been created. Many of these knock-out mice are models for human genetic disorders (e.g., those of endocrine systems such as pancreatic agenesis [homeodomain protein IDX-1], familial hypocalciuric hypercalcemia [calcium receptor], intrauterine growth retardation [insulinlike growth factor-II receptor], salt-sensitive hypertension [atrial natriuretic peptide], and obesity [a3-adrenergic receptor]).
Although targeted transgenesis using chosen site integration and targeted disruption of genes has proven helpful in analyses of the functions of genes, conditionally to induce expression of transgenes or conditionally to inactivate a specific gene is useful. Early on, randomly integrated vectors for the expression of transgenes used the metallothionein promoter that is readily inducible by the administration of heavy metals to transgenic mice. Now techniques have been developed to conditionally inactivate targeted genes in a defined spatial and temporal pattern. Several approaches to achieve conditional gene inactivation have been developed. Two of these approaches are (a) the Cre recombinase–loxP system (Fig. 2-4)18 and (b) the tetracycline-inducible transactivator vector (tTA) system (Fig. 2-5).19 Occasionally, both of these systems have been used effectively to knock out (Cre-loxP) or to attenuate (reverse tTA) the expression of specific genes. Both the Cre-loxP and reverse tTA systems require the creation of two independent strains of transgenic mice, which are then crossed to produce double transgenic mice.

FIGURE 2-4. Schema of the Cre-loxP approach to conditionally knock out a specifically targeted gene in mice. A, The approach requires the creation of two separate strains of transgenic mice that are crossed to produce double transgenic mice to effect the conditional gene knock-outs. One mouse strain is created so as to replace the gene of interest by one that has been flanked by loxP recombination sequences (floxed), using targeted recombinational gene replacement in embryonic stem cells as illustrated in Figure 2-3B. The other mouse strain is a transgenic mouse in which the Cre recombinase enzyme expression vector is targeted to the tissue of interest using a tissue-specific promoter, such as the proinsulin gene promoter, to target and restrict expression to pancreatic B cells. B, A more detailed depiction of the strategy for preparation of the gene replacement by homologous recombination to generate mice with a floxed gene. This approach is similar to that described in Figure 2-3B to create knock-out mice.18

FIGURE 2-5. Diagram showing the approach to reversible conditional expression of a gene in mice, using a tetracycline-inducible gene expression system. A, As in the Cre-loxP system (see Fig. 2-4A), the tetracyclineinducible gene system requires the creation of two independent strains of transgenic mice. One strain of mice targets the expression of a specially engineered transcription factor (rtTA) to the tissue of interest, using a tissue-specific promoter (TSP). B, the rtTA transcription factor consists of a modification of the bacterial tetracycline-responsive repressor that has been genetically engineered so as to convert it into a transcriptional transactivator when exposed to tetracycline or one of its analogs. The other mouse strain is one in which a gene of interest is introduced, usually driven by a ubiquitous promoter such as a viral promoter (CMV, RSV) or an actin promoter. The gene of interest could be one encoding an antisense RNA to a messenger RNA of a protein that is to be knocked out. The creation of double transgenic mice then allows for the expression of the gene of interest in a specific tissue under the control of the induced tetracycline. (See text for more detailed description.57) (tet op, tetracycline resistance operon; P, promoter; As, antisense; TPE, tissue promoter element.)

The Cre-loxP approach is based on the Cre-loxP recombination system of bacteriophage P1 (see Fig. 2-4). This system is capable of mediating loxP sitespecific recombination in embryonal stem cells and in transgenic mice. Conditional targeting requires the generation of two mouse strains. One transgenic strain expresses the Cre recombinase under control of a promoter that is cell-type specific or developmental stage specific. The other strain is prepared by using ES cells to effect a replacement of the targeted gene with an exact copy that is flanked by loxP sequences required for recombination by the Cre recombinase. The recombined gene is said to be floxed. The presence of the loxP sites does not interfere with the functional expression of the gene and will be normally expressed in all of its usual tissues not coexpressing the Cre recombinase. In those tissues in which the Cre is expressed by virtue of its tissue-specific promoter, the target gene will be deleted by homologous recombination. Thus, the Cre-loxP system acts like a timer in which the events that are to take place are predetermined by the prior reprogramming of the genes: the target gene will be ablated during development where and when the promoter chosen to drive the expression of Cre is activated. Thus, a disadvantage of the Cre-loxP system is the lack of control over when the gene knock-out will take place, because it is preprogrammed in the system. Newer genetically engineered Cre derivatives allow for pharmacologic activation of the recombinant event. A potential advantage of the Cre-loxP system is that one can theoretically generate extensive collections of mice expressing the Cre recombinase specifically and individually in many different tissues so that these mice could be made commercially available to investigators.
The Cre-loxP system leads to the irreversible targeted disruption of a particular gene at the time that the promoter encoding the Cre recombinase is activated during development. Having available a system that can be reversibly activated at any time would be desirable. A system that holds promise in this regard is the tetracycline-inducible transactivator vector (forward or reverse tTA), which, in response to tetracycline, switches on a specific gene bearing a promoter containing the tetracycline-responsive operon (see Fig. 2-5). This system allows any recombinant gene marked by the presence of the tet operon to be turned on or off at will simply by the administration of a potent tetracycline analog to the transgenic mice. The vectors were engineered from the sequences of the E. coli bacterial tetracycline resistance operon (tet op), in which a repressor sits on the operon, keeping the resistance gene off. When tetracycline binds to the repressor, it is deactivated, falls off of the operon, and turns on the gene. First, the repressor was converted into an activator by fusing the DNA-binding domain to the potent activator sequence (VP16) of the herpes simplex virus. In this system, tetracycline turned off the activator (tet-off) and thereby caused failure of expression of target genes containing the tet operon binding sites for the repressor turned into an activator. This tTA system required the continued presence of tetracycline to keep the gene off and withdrawal of the tetracycline to turn on the gene, raising problems of long and variable clearance times for the drugs. Turning the gene on by administration of tetracycline (tet-on) would be preferable. Therefore, the tTA vector was reengineered to reverse the action of tetracycline: in the current vector system, the binding of tetracycline to the reverse tTA enhances its binding to the tet operon. Theoretically, as the reverse tTA system now works, any gene can be reversibly turned on by the administration of tetracycline or one if its more potent analogs in the double transgenic mouse, which consists of a cross between a mouse that has the reverse tTA targeted to express in a specific tissue and a mouse that has a ubiquitously expressed transgene for any gene X under the control of the tet operon. The equivalent of gene knock-outs can be accomplished by constructing gene X in a context to express an antisense RNA containing a ribozyme sequence. When induced by tetracycline, antisense-ribozyme RNA binds to the mRNA expressed by gene X, cleaves it, and thereby functionally inactivates the gene.
The availability of the Cre-loxP and the forward and reverse tTA systems now makes it feasible to combine their key features in the creation of triple transgenic mice so that a targeted recombinational disruption of a gene can be accomplished by the administration of tetracycline. The Cre recombinase could be placed under the control of a tissue-specific promoter containing the tet operon uniquely responsive to the presence of tTA and targeted to a specific tissue by standard pronuclear injection targeted transgenesis. A second transgenic mouse is created with a ubiquitously expressed promoter during the expression of the reverse tTA. In the third mouse, the gene desired to be deleted would be replaced with an appropriately floxed gene. The latter mouse would be prepared by implantation of recombinantly engineered ES cells into blastocysts. The administration of tetracycline to the triple transgenic mouse would induce the Cre recombinase in a tissue-specific manner, thus allowing temporal and spatial control of gene knock-outs.
A very informative database of expressed sequence tags (ESTs) is being generated and placed in GenBank. Expressed sequence tags are prepared by random, single-pass sequencing of mRNAs from a repertoire of different tissues, mostly embryonic tissues (e.g., brain, eye, placenta, liver). Currently, the EST database contains ~50% of the estimated expressed genes in humans and mammals (70,000–80,000). The EST database will become extremely valuable when the sequences of the human, rat, and mouse genomes are completed.
Two variants of DNA-array chip design exist.20,21 The first consists of cDNA (sequences unknown) immobilized to a solid surface such as glass and exposed to a set of labeled probes of known sequences, either separately or in a mixture of the probes. The second is an array of oligonucleotide probes (sequences known, based on either known genes in GenBank or ESTs) that are synthesized either in situ or by conventional synthesis followed by on-chip immobilization (Fig. 2-6). The array is exposed to labeled sample DNA (unknown sequence) and hybridized, and complementary sequences are determined.

FIGURE 2-6. Sample preparation and hybridization for oligonucleotide assay. A complementary DNA (cDNA) is transcribed in vitro to RNA, and then reverse-transcribed to cRNA. This material is fragmented and tagged with a fluorescent tag molecule (F). The fragments are hybridized to an array of oligonucleotides representing portions of DNA sequences of interest. After washing, hybridization of the cRNA probe is detected by localization of the fluorescent signals. (PCR, polymerase chain reaction.)

In cDNA chips, immobilized targets of single-stranded cDNAs prepared from a specific tissue are hybridized to single-stranded DNA fluorescent probes produced from total mRNAs to evaluate the expression levels of target genes.
The oligonucleotide gene chip (1.28 × 1.28 cm2) consists of a solid-phase template (glass wafer) to which high-density arrays of oligonucleotides (distance between oligonucleotides of 100 Å) are attached, with each probe in a predefined position in the array. Each gene EST is represented by 20 pairs of 25 base oligonucleotides from different parts of the gene (5′ end, middle, and 3′ end).
The specificity of the detection method is controlled by the presence of single-base mismatch probes. Pairs of perfect and single-base mismatch probes corresponding to each target gene are synthesized on adjacent areas on the arrays. This is done to identify and subtract nonspecific background signals. The gene chip is sensitive enough to detect one to five transcripts per cell and is much more sensitive than the Northern blot technique.
Poly (A) mRNA is isolated from cells or tissue of interest, and synthesis of double-stranded cDNA is accomplished by reverse transcription of cDNA, followed by synthesis of double-stranded cDNA using DNA polymerase I. In vitro transcription of double-stranded cDNA to cRNA is accomplished using biotin-16-UTP and biotin-11-CTP for labeling and a T7 RNA polymerase as enzyme. This cRNA is used for hybridization with the gene chip. The gene chip is stained with R-physoerythrin streptavidin to detect biotin-labeled nucleotides, and different wash cycles are performed. Thereafter the gene chip is scanned digitally and analyzed by special software. (A grid is automatically placed over the scanned image of the probe array chip to demarcate the probe cells.) After grid alignment, the average intensity of each probe cell is calculated by the software, which then analyzes the patterns and generates a report.
The applications of the gene chip include:

Simultaneous analysis of temporal changes in gene expression of all known genes and ESTs.

Sequencing of DNA.

Large-scale detection of mutations and polymorphisms in specific genes (i.e., BRCA1, HIV-1, cystic fibrosis CFTR, b-thalassemia).

Gene mapping by determining the order of overlapping clones.
Expensive equipment for generating and analyzing the data using genechips is required. When the cloning of all genes is completed (Human Genome Project), the gene chip will allow monitoring of the expression of all known genes in various situations.
A genetic linkage map shows the relative locations of specific DNA markers along the chromosome.22,23,24,25,26 and 27 Any inherited physical or molecular characteristic that differs among individuals and is easily detectable in the laboratory is a potential genetic marker. Markers can be expressed DNA regions or DNA segments that have no known coding function, but whose inheritance pattern can be followed. DNA sequence differences (polymorphisms; i.e., nucleotide differences) are especially useful markers because they are plentiful and easy to characterize precisely. Markers must be polymorphic to be useful in mapping. Alternative DNA polymorphisms exist among individuals, even among members of a single family, so that they are detectable among different families. Polymorphisms are variations in DNA sequence in the genome that occur every 300 to 500 base pairs. Variations within protein-encoding exon sequences can lead to observable phenotypic changes (e.g., differences in eye color, blood type, and disease susceptibility). Most variations occur within introns and have little or no effect on the phenotype (unless they alter exonic splicing patterns), yet these polymorphisms in DNA sequence are detectable and can be used as markers. Examples of these types of markers are: (a) restriction fragment length polymorphisms (RFLPs), which reflect sequence variations in DNA sites that are cleaved by specific DNA restriction enzymes; and (b) variable number of tandem repeat sequences (VNTRs), which are short repeated sequences that vary in the number of repeated units and, therefore, in length. The human genetic linkage map is constructed by observing how frequently any two polymorphic markers are inherited together.
Two genetic markers that are in close proximity tend to be passed together from mother to child. During gametogenesis, homologous recombination events take place in the metaphase of the first meiotic step (meiotic recombination crossing-over). This may result in the separation of two markers that originally resided on the same chromosome. The closer the markers are to each other, the more tightly linked they are and the less likely that a recombination event will fall between and separate them. Recombination frequency provides an estimate of the distance between two markers.
On the genetic map, distances between markers are measured in terms of centimorgans (cM), named after the American geneticist Thomas Hunt Morgan. Two markers are said to be 1 cM apart if they are separated by recombination 1% of the time. A genetic distance of 1 cM is roughly equal to a physical distance of 1 million base pairs of DNA (1 Mb). The current resolution of most human genetic map regions is approximately 10 Mb.
An inherited disease can be located on the map by following the inheritance of a DNA marker present in affected individuals but absent in unaffected individuals, although the molecular basis of a disease or a trait may be unknown. Linkage studies have been used to identify the exact chromosomal location of several important genes associated with diseases, including cystic fibrosis, sickle cell disease, Tay-Sachs disease, fragile X syndrome, and myotonic dystrophy.
The restriction endonucleases, which have been isolated from various bacteria, recognize short DNA sequences and cut DNA molecules at those specific sites. A natural biofunction of restriction endonucleases is to protect bacteria from viral infection or foreign DNA by destroying the alien DNA. Some restriction enzymes cut DNA very infrequently, generating a small number of very large fragments, whereas other restriction enzymes cut DNA more frequently, yielding many smaller fragments. Because hundreds of different restriction enzymes have been characterized, DNA can be cut into many differentsized fragments.
Different types of physical maps vary in their degree of resolution. The lowest resolution physical map is the chromosomal (cytogenetic) map, which is based on the distinctive banding pattern observed by light microscopy of stained chromosomes. A cDNA map shows the locations of expressed DNA (exons) on the chromosomal map. The more detailed cosmid contiguous DNA block (contig) map depicts the order of overlapping DNA fragments spanning the genome (see the section High-Resolution Physical Mapping). A macrorestriction map describes the order and distance between restriction enzyme cleavage sites. The highest resolution physical map will be the complete elucidation of the DNA base-pair sequence of each chromosome in the human genome.
Chromosomal Map. In a chromosomal map, genes or other identifiable DNA fragments are assigned to their respective chromosomes, with distances measured in base pairs. These markers can be physically associated with particular bands (identified by cytogenetic staining) primarily by in situ hybridization, a technique that involves tagging the DNA marker with an observable label. The location of the labeled probe can be detected after it binds to its complementary DNA strand in an intact chromosome.
As with genetic linkage mapping, chromosomal mapping can be used to locate genetic markers defined by traits observable only in whole organisms. Because chromosomal maps are based on estimates of physical distance, they are considered to be physical maps. The number of base pairs within a band can only be estimated.
Fluorescence In Situ Hybridization.28,29 A fluorescently labeled DNA probe locates a DNA sequence detected on a specific chromosome. The fluorescence in situ hybridization (FISH) method allows for the orientation of DNA sequences that lie as close as 2 to 5 Mb. Modifications to the in situ hybridization methods, using chromosomes at a stage in cell division (interphase) when they are less compact, increase map resolution by an additional 100,000 base pairs.
A cDNA map shows the positions of expressed DNA regions (exons) relative to particular chromosomal regions or bands. (Expressed DNA regions are those transcribed into mRNA.) The cDNA is synthesized in the laboratory using the mRNA molecule as the template. This cDNA can be used to map the genomic region of the respective molecule. A cDNA map can provide the chromosomal location for genes whose functions are currently unknown (ESTs). For hunters of disease genes, the map can also suggest a set of candidate genes to test when the approximate location of a disease gene has been mapped by genetic linkage analysis.
Two current approaches to high-resolution mapping are termed top-down (producing a macrorestriction map) and bottom-up (resulting in a contig map). With either strategy, the maps represent ordered sets of DNA fragments that are generated by cutting genomic DNA with restriction enzymes. The fragments are then amplified by cloning or by PCR methods. Electrophoretic techniques are used to separate the fragments (according to size) into different bands, which are visualized by staining or by hybridization with DNA probes of interest. The use of purified chromosomes, separated either by fluorescence-activated flow sorting from human cell lines or in hybrid cell lines, allows a single chromosome to be mapped.
A number of strategies can be used to reconstruct the original order of the DNA fragments in the genome. Many approaches make use of the ability of single strands of DNA and/or RNA to hybridize to form double-stranded segments. The extent of sequence homology between the two strands can be inferred from the length of the double-stranded segment. Fingerprinting uses restriction enzyme cleavage map data to determine which fragments have a specific sequence (finger-print) in common and, therefore, overlap. Another approach uses linking clones as probes for hybridization to chromosomal DNA cut with the same restriction enzyme.
In top-down mapping, a single chromosome is cut (using rare-cutter restriction enzymes) into large pieces, which are ordered and subdivided; the smaller pieces are then mapped further. The resulting macrorestriction maps depict the order of and distance between locations at which rare-cutter restriction sites are found in the chromosome. This approach yields maps with more continuity and fewer gaps between fragments than contig maps, but map resolution is lower and the map may not be useful in finding particular genes. In addition, this strategy generally does not produce long stretches of mapped sites. Currently, this approach allows DNA pieces to be located in regions measuring ~100 kb to 1 Mb.
The development of pulsed-field gel (PFG) electrophoretic methods has improved the mapping and cloning of large DNA molecules. Whereas conventional gel electrophoretic methods separate pieces of DNA <40 kb in size, PFG separates molecules up to 10 Mb, allowing application of both conventional and new mapping methods to large genomic regions.
The bottom-up (contig) approach involves cutting the chromosome into small pieces, each of which is cloned and ordered. The ordered fragments form contigs, or contiguous DNA blocks. Currently, the resulting library of clones varies in size from 10 kb to 1 Mb. An advantage of this approach is the accessibility of these stable clones to other researchers. Contig construction can be verified by FISH, which localizes cosmids to specific regions within chromosomal bands.
Contig maps consist of a linked library of small overlapping clones of DNA representing a complete chromosomal segment. Although useful for finding genes localized in a small area, contig maps are difficult to extend over large stretches of a chromosome because all regions are not clonable. DNA probe techniques can be used to fill in the gaps, but they are time consuming.
Technological improvements now make possible the cloning of large DNA pieces using artificially constructed chromosome vectors that carry human DNA fragments as large as 1 Mb. These vectors are maintained in yeast cells (i.e., YACs).30,31 Before YACs were developed, the largest cosmids carried inserts of only 20 to 40 kb. YAC methodology drastically reduces the number of clones to be ordered; many YACs span entire human genes. A more detailed map of a large YAC insert can be produced by subcloning (a process in which fragments of the original insert are cloned into smaller insert vectors). Because some YAC regions are unstable, large-capacity bacterial vectors have also been developed (bacterial artificial chromosome, BAC).
Flow-sorting uses flow cytometry to separate, according to size, chromosomes isolated from cells during cell division, when they are condensed and stable.32,33 As the chromosomes flow singly past a laser beam, they are differentiated by analyzing the amount of DNA present, and individual chromosomes are directed to specific collection tubes.
In somatic cell hybridization, human cells and rodent tumor cells are fused (hybridized); over time, after the chromosomes mix, human chromosomes are preferentially lost from the hybrid cell until only one or a few remain. Those individual hybrid cells are then propagated and maintained as cell lines containing specific human chromosomes. Improvements to this technique have generated a number of hybrid cell lines, each with a specific single human chromosome.
Starting maps and sequences is relatively simple. Finishing them requires either development of new strategies or use of a combination of existing methods. After a sequence is determined, the task that remains is to fill in the many large gaps left by current mapping methods. One approach is a single chromosome microdissection, in which a piece of DNA is physically cut from a chromosomal region of particular interest and then broken up into smaller pieces and amplified by PCR or cloning. These fragments of DNA can then be mapped and sequenced by conventional methods.
Chromosome walking, one strategy for cloning genes and filling gaps, involves hybridizing a primer of known sequence to a clone from an unordered genomic library and synthesizing a short complementary strand (called walking on the chromosome). The complementary strand is then sequenced and its end used as the next primer for further walking. In this way, the adjacent (previously unknown) region is identified and sequenced. The chromosome is thus systematically sequenced from one end to the other. Because primers must be synthesized chemically, a disadvantage of this technique is the need to construct a large number of different primers for walking large distances. Chromosome walking is also used to locate specific genes by sequencing the chromosomal segments between markers that flank the gene of interest (i.e., positional cloning).
The degree of difficulty encountered in finding a disease gene of interest depends largely on what information is already known about the gene and especially on what kind of DNA alterations cause the disease. Spotting the disease gene is difficult when disease results from a single altered DNA base; sickle cell anemia is an example of such a case, as are probably most major human inherited diseases. When disease results from a large DNA rearrangement, the anomaly can be detected as alterations in the physical map of the region or even by direct microscopic examination of the chromosome. The location of these alterations pinpoints the site of the gene.
Another approach to identifying a disease-causing or diseaserelated gene is the candidate gene approach. Here, a gene known to be important in the development or function of a certain organ or organ system is analyzed for mutations in a cohort with the disease and compared with the gene in a healthy control group. This approach has been valuable in identifying single gene mutations in well-defined, relatively small pedigrees. With the possibility for large-scale genome-wide screening using DNA-array technology, this approach may be applied to large cohorts in a widespread manner in the future.
The Human Genome Project was begun in 1990 by the U.S. government and is coordinated by the U.S. Department of Energy and the National Institutes of Health. Although it was originally conceived as a 15-year program, rapid technological advances as well as collaboration between genome projects of several countries (United Kingdom, Germany, Japan, France) and privately funded endeavors to sequence the human genome have accelerated the project to an expected completion date of 2001, with 90% of the genome available in the spring of 2000.34 The Human Genome Project initiated by the U.S. government has set out to sequence the entire genome of one human individual (identity strictly secret), whereas the privately initiated effort is sequencing the entire genome of five different individuals. The latter approach is addressing the issue of some, although limited, variability in the genome of humans.
The goals of the Human Genome Project are to identify the estimated 80,000 expressed genes in human DNA; to determine the sequences of the ~3 billion chemical bases that make up the human DNA; to store the information in databases that will be made accessible to the public; to develop tools for data analysis; and to address the ethical, legal, and societal issues that may arise from the project. Furthermore, to allow comparison of genetic information among species and to address questions related to the interaction of humans with infectious pathogens, researchers are studying the genetic makeup of several nonhuman organisms, including Drosophila melanogaster, Plasmodium falciparum, and the laboratory mouse. The genomes of several organisms such as E. coli, Saccharomyces cerevisiae, Caenorhabditis elegans, Bacillus subtilis, Synechtosis species, A. fulgidus, P. aerophilum, Haemophilus influenzae, M. thermoautrophicum, M. jannaschii, A. aolicolus, Borrelia burgdorferi, Treponema pallidum, Mycoplasma pneumoniae, M. genitalum, Chlamydia trachomatis, Rickettsia prowazekii, Helicobacter pylori, and Mycobacterium tuberculosis are already known and accessible.
During progress of the Human Genome Project, several new goals have been formulated. For example, one goal is to identify regions of the human genome that differ from person to person. Although the majority of individuals’ DNA sequences are the same, estimates are that humans are only ~99% identical genetically. These DNA sequence variations can have a major impact on how people’s bodies respond to disease (i.e., to environmental insults, such as bacteria, viruses, and toxins; and to drugs and other therapies). Methods have been developed to rapidly detect different types of variation, particularly the most common type, called single-nucleotide polymorphisms (SNPs), which occur approximately once every 100 to 300 bases. SNP maps will ultimately help identify multiple genes associated with complex diseases such as cancer, diabetes, vascular disease, and some forms of mental illness. These associations are difficult to establish with conventional gene-hunting methods. because a single altered gene may make only a small contribution to disease risk.
The atlas of the human genome will revolutionize medical practice and biologic research into the twenty-first century and beyond. All human genes will be found, and accurate diagnostics will be developed for most inherited diseases. In addition, animal models for human disease research will be more easily developed, facilitating the understanding of gene function in health and disease.
Single genes associated with a number of diseases (e.g., cystic fibrosis, Duchenne muscular dystrophy, myotonic dystrophy, neurofibromatosis, diabetes mellitus, and retinoblastoma) have already been identified. Diseases caused by several genes or by a gene interacting with environmental factors can be studied more efficiently. Genetic susceptibilities have been implicated in many major disabling and fatal diseases, including heart disease, stroke, diabetes, and several kinds of cancer. The identification of these genes and their proteins will pave the way to more effective therapies and preventive measures.
The potential benefits of the Human Genome Project are manifold. Only some of the important potential applications are discussed here. Molecular medicine (a term lending more significance to medicine and the understanding of human disease and its potential cure at the molecular level) will gain increasing importance in medical practice. Improved diagnosis and earlier detection of genetic predisposition to disease are already commonplace for certain conditions (cystic fibrosis, trisomy, fragile X syndrome, myotonic dystrophy, and neurofibromatosis). The design of drugs that are tailored to the demands imposed by genetic predisposition and the choice of treatment based on genetic information will allow a more rational approach to treatment.
Microbial genomics, which yields the sequence and function of microbial genes, can potentially identify new energy sources (biofuels) and identify bacteria useful in environmental remediation, toxic waste reduction, and industrial processing. New tools for risk assessment for exposure to radiation and toxic agents can be made more reliably at the genetic level with information on the human genome. Furthermore, comparison of genomic sequences can increase the capabilities to study evolution through germline mutations in lineages; examine migration of different population groups based on female genetic inheritance; investigate mutations on the Y chromosome to trace lineage and migration of males; and compare breakpoints in the evolution of mutations with ages of populations and historical events.
DNA forensic analysis aimed at proper identification of individuals involved in crimes and catastrophic events, establishment of paternity and family relationships, and matching of organ donors is already commonplace with the limited possibilities available today. The expansion of genetic information with the Human Genome Project should allow for improving the reliability of such tests.
Agriculture, livestock breeding, and bioprocessing are additional areas affected directly by the genome project. Understanding plant and animal genomes can potentially allow the creation of stronger, more resistant plants and animals. Bioengineered seeds are already being used to grow disease-resistant crops, thereby reducing costs in agriculture. Some additional applications that are already being tested experimentally are the design of pesticides, development of edible vaccines incorporated into food products, development of new environmental cleanup uses for plants like tobacco, and generation of animals and plants producing molecules to be used for the treatment of human conditions (i.e., clotting factors, hemoglobin). Although human genome research itself does not pose any new ethical dilemma, the use of data arising from these studies presents challenges that need to be addressed before the data accumulate significantly. Some of the issues pertinent to the question are summarized in Table 2-4. To assist in policy development, the ethics component of the Human Genome Project is funding conferences and research projects to identify and consider relevant societal issues, as well as activities to promote public awareness of these topics.

TABLE 2-4. Implications and Issues Arising from the Human Genome Project

ES cells are pluripotent cells that give rise to all adult cell types.35 They can be derived from the blastocyst, a preimplantation-stage embryo,36 or from primordial germ cells, cells of the early embryo that eventually differentiate into sperm and oocytes.37 Mouse ES cells have been used for approximately a decade to generate genetically altered mice (knock-out, knock-in, Cre recombinants). Mouse ES cells are pluripotent in that they can differentiate into many cell types. ES cells, unlike fertilized eggs, cannot develop completely into individual organisms. ES cells that are placed into a uterus will never develop into an embryo. After desired manipulations, ES cells are injected into the cavity of a developing blastocyst, which ultimately develops into a chimera harboring all tissue cells from the original embryo as well as filiae of the artificially introduced ES cells. ES cells cannot develop into an embryo on their own; they must be placed into an artificial environment, one in which the host cells provide the placental tissues of the conceptus. The derivation of human ES cells from developing fetuses has opened exciting new possibilities for therapy as well as raising legal and ethical issues.36,38
To be able to generate tissue for potential transplantation or organ replacement, ES cells that are immunologically identical to the prospective host would need to be generated. This could possibly be achieved by somatic nuclear transfer, with a nucleus of a prospective host inserted into a recipient enucleated oocyte of either the same or a different species. From the embryo that developed, the inner cell mass of the blastocyst would need to be isolated and grown in vitro to yield ES cells, which then would need to be differentiated into the desired tissue. To date, not all of these steps have been mastered.
The use of ES cells therapeutically has dangers, however. Mouse ES cells are tumorigenic, growing into teratomas or teratocarcinomas when injected anywhere in the adult mouse. Human ES cells might behave similarly.
Renewable tissues (blood, intestinal epithelium, epidermis) are considered to harbor stem cells that can, by multiplication, yield cells which can then further differentiate into cells of the respective host tissue.
Cells in the nervous system have been found to have the capacity to generate new neurons and glial cells (astrocytes and oligodendrocytes), and, because of this, they are considered to be neuronal stem cells.39,40,41 These cells can be isolated from the wall of the lateral ventricle of the brain.42 These cells, which constitute ~0.1% to 1% of the cells in the ependymal lining, express immature neural markers consistent with a stem-cell function.
In vitro, these stem cells divide in response to epidermal growth factor and fibroblast growth factor-2.43 On dividing, they are thought to give rise to both neural and glial progeny. If these cells are transplanted into the intact brain, they integrate and differentiate into a range of neuronal and glial cells.44,45 Moreover, these cells can regenerate blood cells when transplanted into lethally radiated host mice.
Human embryonic stem cells, which can be cloned46 and grown for extended periods,47 reside in the adult brain (Table 2-5).48

TABLE 2-5. Human Stem Cells Isolated

Another example of pluripotent stem cells is mesenchymal stem cells (MSCs), which can be isolated from bone marrow.49 These cells can be expanded in culture and still differentiate into osteoblasts, osteocytes, chondrocytes, adipocytes, tendonassociated cells, and myotubules50; they have been transplanted into children with osteogenesis imperfecta and into women who have undergone high-dose myelotoxic chemotherapy,51,52 and used for Achilles tendon repair.53
Several ethical and biologic questions need to be addressed before ES cells can be used widely for human applications (Table 2-6).

TABLE 2-6. Biologic and Ethical Questions Pertinent to Use of Embryonic Stem (ES) Cells

Cloning by blastomere separation (called “twinning”) (Table 2-7) involves splitting a developing embryo soon after fertilization of the egg by a sperm to give rise to two or more embryos. The resulting organisms are identical twins (clones) containing DNA from both the mother and the father. This technique has been used for approximately one decade for generating transgenic or knock-out embryos and animals. Twinning in the sense of generating two or more genetically identical organisms has been practiced for thousands of years with plants.

TABLE 2-7. Animals Cloned by Somatic Nuclear Transfer

Cloning by somatic nucleus transfer involves the insertion of a nucleus of a cell from one single adult into an egg from which the original nucleus has been removed. This allows for the generation of an animal with the genetic information coming from one single adult (as opposed to cloning by blastomere separation). The technique of somatic nucleus transfer was attempted in the 1950s in frogs, and early attempts were conducted in mammals in the 1980s. In 1996, lambs were successfully cloned via somatic nucleus transfer from embryonic cells that had been cultured in vitro for several months. Dolly, the lamb that was cloned by transferring the nucleus of a cell from the udder of a 6-year-old sheep, gained broad attention. The theoretical possibility was created of generating an unlimited number of genetically identical individuals from one single parent. However, the clones generated through somatic nucleus transfer are not completely genetically identical to the donor animal. The mitochondrial DNA in the host oocyte originates from the mother. Dolly appears to be a healthy sheep and has reportedly given birth to a lamb named Bonnie per vias naturalis. In 1997, a second lamb named Polly was generated by an additional cellular manipulation. Nuclei from fetal fibroblasts were modified in vitro to incorporate the cDNA for human factor IX under control of the b-lactoglobin promoter. These “transgenic” fibroblasts were used as donor nuclei for somatic transfer. Polly, the lamb generated by this method, produces up to 40 g/L of human factor IX in its milk. Factor IX can now be isolated without much effort and used in therapeutic applications. Since these advances were reported, successful cloning of calves and mice has also been accomplished.
The majority of clones fail sometime during development, and some fail after birth. One report suggests that the immune defense system of animals cloned by somatic nucleus transfer may be impaired. Further, telomeres of cloned animals appear to be shorter, suggesting that the cells may have a reduced life span and the animals may manifest symptoms of premature aging. However, the offspring of Dolly appears to have telomeres of normal length. Some accomplishments of somatic nuclear transfer are summarized in Table 2-7.
A main area of investigation when the entire genome of humans as well as that of laboratory animals (i.e., mice and rats) is known and large-scale mutation screening of the entire genome can be performed rapidly will be the study of the function of genes identified in the context of diseases and the consequences of mutations in these genes.
For this purpose, collections of knock-out mice strains are made available to researchers for analysis in various assay systems. Furthermore, tissue-specific ablation of certain genes with the Cre-loxP methodology are being provided to researchers. The effects of gene ablation in defined physiologic situations and the effects of drugs in an animal lacking the function of a defined gene can then be studied more efficiently.
The concept of gene therapy is the transfer of genetic information to a host organism or a specific tissue within an organism. The product of the delivered gene either may be missing or of insufficient quantity in the host (replacement therapy) or may be of pharmacologic/therapeutic value (e.g., immune modulation, vaccination). Agents carrying the DNA to target cells are called vectors. The requirements for an ideal vector are that it should accommodate an unlimited amount of inserted DNA, lack the ability of autonomous replication of its own DNA, be easily manufactured, and be available in a concentrated form. For most applications, it should have the ability to target specific cell types or limit its gene expression in the long term or in a controlled fashion. It should not be toxic or immunogenic. Such vectors do not exist, and none of the DNA delivery systems so far available for in vivo gene transfer is perfect. Thus, the advancement of gene therapy lies in the development and improvement of new gene vector systems.
Different ways of introducing genes into mammalian cells and tissues have been devised, the simplest being the inoculation of naked DNA by means of microinjection, electroporation, or bioballistics (known as the gene gun technique55). More elaborate and efficient ways include the use of self-assembling complexes of lipid-DNA (e.g., liposomes) protein-DNA, or lipid-protein-DNA, and viral vectors. The physicochemical methods of gene delivery are summarized in Table 2-2 and Table 2-3.
Viral vectors can be fragments of viral DNA containing the DNA to be transferred or the viral particle itself.56 The viral particle is rendered replication defective through the manipulation of the viral DNA, and the end product is a nonpathogenic viral vector carrying the genetic information of therapeutic interest. Many different types of virus have been used for genetransfer purposes. The viruses holding most promise for future application in biomedicine (retroviruses, adenoviruses, and adenoassociated viruses) are discussed in the following sections.
Retroviral Vectors. The Moloney murine leukemia virus (MoMuLV) was the first virus used in developing a vector system for gene transfer. Parts of the retroviral genome that are involved in the replication of the MoMuLV—gag, pol, and env—are removed and replaced by a gene of interest. What remains of the retrovirus are the long terminal repeats harboring regulatory elements, integration signals, and transcription promoters, and a packaging signal. To produce retroviral vectors carrying the gene of interest, one must use a packaging cell line harboring the gag, pol, and env genes, which have previously been incorporated into the genome of such packaging cells. The vectors are produced in high titers in the packaging cells, purified, and injected into the organism (in vivo gene therapy) or put in contact with cells collected from the patient and maintained in cell culture (ex vivo gene therapy). Retroviral vectors have the ability to integrate their genomic material into the host genome as a result of the presence of the remaining retroviral regulatory sequences in the form of the long terminal repeats. Once integrated, the inserted gene can be expressed to produce the desired protein.
Advantages of retroviral vectors are that they are well characterized, they can be produced in high titers, and they have a high efficiency of infection. Disadvantages are the limited size of DNA (7–8 kb) that retrovirus vectors can accommodate, the requirement that the target cells be in cell division to allow for the integration of the vectors, and the potential for insertional mutagenesis due to retroviral vector integration at random in the genome. The latter feature has the potential to interrupt important genes in the cell, with serious consequences that include oncogenesis through the activation of protooncogenes or inactivation of tumorsuppressor genes.
Lentiviruses are a group of retroviruses that have the ability to infect and integrate their genome even in cells that are not dividing. Thus, the use of lentiviruses as gene therapy vectors is broader than that of other retroviruses and is under investigation.
Adenoviral Vectors. The human adenoviruses are nonenveloped DNA viruses with a linear double-stranded DNA of ~36 kb, encapsulated in an icosahedral capsoid measuring 70 to 100 nm in diameter. Adenoviruses are known pathogens in humans, and most, if not all, adult humans have been exposed to adenoviruses and have antibodies against adenovirus antigens.
An adenovirus does not depend on host-cell division for its replication, and the chromosome rarely integrates into the genome, remaining episomal in most cases. Integration seems to occur mainly in the presence of high levels of infection in dividing cells, but this event does not contribute significantly to the utility of these viruses as vectors. Adenoviral vectors have a broad spectrum of cell infectivity that includes virtually all postmitotic and mitotic cells; they also can be produced in high titers.
The genome of the adenovirus encodes ~15 proteins. Viral gene expression occurs in a coordinated fashion and is mainly directed by the E1A and E1B genes, localized within the 5′ region of the adenoviral genome. These genes have transactivation functions for the transcription of various viral and host-cell genes. Because E1 genes are involved in adenoviral replication, their removal renders the virus replication incompetent. The removal also creates room for insertion of a foreign gene of therapeutic interest. An exogenous DNA can also replace the E3 region, which produces a product enabling the virus to evade the immune system. Packaging cells carrying adenoviral genes that provide transcomplementation functions are required to produce defective adenoviral vectors. Packaging cells of the NIH-293 cell lineage are human embryo kidney (HEK) cells that have been previously transformed with type 5 adenovirus. These cells retain the E1A and E1B regions of the viral genome covalently linked to their genomic DNA.
Disadvantages of the adenoviral vectors include the short duration of transgene expression, because the vector usually does not integrate stably into the host-cell genome. Further, the size of the inserted foreign DNA sequence is limited, and cellular and humoral immune responses are typically triggered against the adenoviral particles or against the host cell that eventually expresses adenoviral proteins, thus limiting the longevity of the adenovirus vector.
Adenoassociated Viral Vectors. Adenoassociated virus (AAV) is a small nonenveloped, nonpathogenic DNA virus belonging to the Parvoviridae family. The AAV genome is a single-strand DNA molecule of 4681 bases, including two inverted long terminal repeats (ITRs). ITRs are 145-base-long palindromic sequences involved in the regulation of the AAV cell cycle. They are located in the 5′ and 3′ terminal portions of the viral genome and serve as origins and initiators for DNA replication. Flanked by the ITRs, two large open-reading frames code for a regulatory protein and a structural protein, called rep and cap, respectively. The protein coding sequence located in the 5′ region (rep gene) encodes four nonstructural proteins involved in the genomic replication. The 3′ region contains the cap gene, which encodes three structural proteins required for the formation of the viral capsid. AAV is capable of replication in a cell only in the presence of a helper virus (adenovirus or herpes virus) that provides by transcomplementation the helper factors that are essential for its replication. In the absence of a helper virus, the AAV genome preferentially integrates into a specific site on the short arm of chromosome 19, between q13.3 and qter, called AAVS1. The ITRs as well as a rep transcript play an important role in this process, which results in a latent infection in mitotic as well as in postmitotic cells. Episomal virus and insertion in nonspecific sites has been documented.
Among the advantages of AAV as a potential gene vector in human gene therapy are the lack of relation to human diseases, broad infectivity spectrum, and ability to stably integrate into the host genome. This integration can occur in cells that are not dividing, although at a lower frequency than in dividing cells. The site-directed integration is also a most favorable property of AAV.
Immunization with transfer of genetic material represents a novel approach to vaccination.57,58 The technology involves transferal of a gene (encoding an antigenic protein cloned in an expression vector) to a host, leading to the induction of an immune response. Direct gene transfer may be undertaken using either viral vectors or recombinant plasmid DNA. Viral vectors have the disadvantages of being derived from pathogens (like traditional vaccines based on attenuated virus), and, therefore, are of limited interest for the purpose of immunization. In contrast, DNA plasmids encoding antigens are more frequently used because they do not have the inconvenience of classic vaccines: they are safe, inexpensive, easy to produce, heat stable, and amenable to genetic manipulation.
Currently, two main delivery systems are available for gene vaccination. Plasmid DNA is injected intramuscularly, or DNA is coated onto gold beads and transferred into the epidermis or dermis by a bioballistic process (gene gun). The intramuscular injection is the most widely used method for immunization, and it consists of direct injection of naked DNA into skeletal muscle. Plasmid DNA in some instances is injected into muscle directly in saline solution or after injection of toxins or a local anesthetic to cause necrosis and regeneration of the injected muscle, thereby increasing the expression of the encoded antigen and amplifying the immunologic response. It is unclear whether the elevated antigen in regenerating muscle cells is due to an increased expression of the antigen gene or to the antigen contained within antigen-presenting cells that are recruited to the site of tissue damage. Humoral and cell-mediated immune responses have been induced by the direct intramuscular injection of plasmid DNA endocrine immunogens. An antibody response was first reported against an influenza virus protein in mice, and specific cytotoxic T-cell responses were also detected in different systems (i.e., human immunodeficiency virus infection and hepatitis B) after genetic immunization. Protective immunity was first demonstrated in mice injected intramuscularly with nucleoprotein DNA of influenza virus. In this model, researchers have indicated that both CD4+ and CD8+ T cells contributed to the protection. Protective immune responses have also been demonstrated in mice against Leishmania major, Plasmodium yoelii, Mycobacterium tuberculosis, dengue virus, and herpes simplex virus.
The bioballistic (gene gun) method uses a helium gas pressuredriven device to deliver gold particles coated with plasmid directly into the skin. When gene vaccines are administered by gene gun technology, most of the plasmid DNA is taken up by keratinocytes and some dermal fibroblasts; they become transfected and produce the encoded antigen. Humoral responses using bioballistic approaches were demonstrated using plasmids encoding human growth hormone and human a-antitrypsin.
The nature of the immune response elicited by these DNA vaccination approaches is not clearly understood. In an initial study, the suggestion was made that DNA vaccination elicits a cell-mediated immune response, because passive transfer of serum from immune mice did not engender protective immunity. Depletion experiments demonstrated that both CD4+ and CD8+ cell populations were involved in host protection against infection. However, some investigators have also reported an induction of immunoglobulins after gene gun–mediated immunization.

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