Molecular Biology for the Clinician: Understanding Current Methods
The basics of molecular biology involve the replication of deoxyribonucleic acid and its transcription and translation into proteins. Biochemical assays such as the Southern blot analysis, polymerase chain reaction (PCR), Northern blot analysis, reverse-transcriptase PCR, microarray technology, Western blot analysis, immunohistochemistry, enzyme-linked immunosorbent assay, and flow cytometry utilize various aspects of molecular biology. To understand these assays requires some basic understanding of the principles of molecular biology. This paper provides basic information on the methodology and techniques used in these assays.
Introduction to Basic Molecular Biology
The nucleus of all mammalian cells contains a specified number of chromosomes that differs between species. Diploid cells have two sets of chromosomes, with one copy being inherited from each parent cell. Including the sex chromosomes, humans have 46 chromosomes, dogs have 78 chromosomes, and cats have 34 chromosomes.1 Chromosomes divide by mitosis, and all somatic cells undergo mitosis. Genetic material in chromosomes is composed of deoxyribonucleic acid (DNA). Genes that are on the same chromosome show linked inheritance (i.e., they tend to stay together when DNA undergoes replication). Genes that are on different chromosomes segregate independently (i.e., they recombine at random from one generation to the next). A genome consists of the entire set of chromosomes for any organism.
Cells also contain ribonucleic acid (RNA), which is found in abundance in both the cytoplasm and the nucleus. In the nucleus, RNA is concentrated in nucleoli and is attached to the chromosomes.2 Ribonucleic acid and DNA are constructed of nucleotide building blocks. Each nucleotide contains a phosphate group linked to a sugar group containing five carbon atoms, which in turn is linked to either a purine or a pyrimidine base. Purine bases include adenine (A) and guanine (G). Pyrimidine bases include cytosine (C), thymine (T), and uracil (U). Thymine is only found in DNA, and U is only found in RNA.
Deoxyribose is the sugar of DNA, and ribose is the sugar of RNA. In both DNA and RNA, the nucleotides are joined by covalent bonds linking the phosphate group of one nucleotide to a hydroxyl group on the sugar of the adjacent nucleotide. These linkages connect the 5′ carbon atom of one deoxyribose (or ribose) residue to the 3′ carbon atom of the deoxyribose (ribose) in the adjacent nucleotide.1 The four different bases of DNA are attached in any order along the spine of the helix, giving DNA its specificity. Genetic information is carried in the sequence of the bases, like letters and words making up sentences. The orientation of the sugar-phosphate spine defines the direction of the nucleic acid chains. Nucleic acid sequences are referred to as 5′ to 3′, because the terminal nucleotide at one end of the chain has a free 5′ (amino) group, and the terminal nucleotide at the other end has a free 3′ (carboxy) group. The 5′ terminus is to the left of the 3′ terminus. Chains of DNA and RNA elongate from the 5′ end to the 3′ end; therefore, transcription begins at the 5′ end and terminates at the 3′ end.2
Deoxyribonucleic acid is composed of two strands that form a double helix [Figure 1]. The two strands are connected by hydrogen bonds between the nitrogenous bases. The bases pair up in a complementary manner. Guanine hydrogen atoms bond only with C, and A hydrogen atoms bond only with T. The model originally described by Watson and Crick requires that the two chains run in an antiparallel manner, meaning that one strand runs in the 5′ to 3′ direction and its partner runs in the 3′ to 5′ direction.2 Guanine-cytosine base pairs are more stable and stronger than AT base pairs, because GC base pairs have three hydrogen bonds and AT base pairs only have two bonds.
Precise replication of genetic material is crucial. Replication of parental DNA strands requires that the strands separate to allow each to become a template for synthesis of a new complementary strand. The separation is transient, and the replication fork is the point where the DNA duplex is disrupted, forming a separated junction. Each of the daughter strands is identical to the original parental strand and consists of one parental strand and one newly synthesized strand. Specific enzymes catalyze synthesis of DNA. For example, DNA polymerase catalyzes DNA synthesis. An interesting limitation is that all polymerases work only from the 5′ to 3′ direction. The replication fork moves from the 5′ to 3′ direction on the leading strand but from 3′ to 5′ on the lagging strand (i.e., the DNA strand on the opposite side of the replication fork that is being replicated in short fragments). Although the lagging strand appears to replicate backward, it actually synthesizes DNA in the appropriate 5′ to 3′ direction, because RNA primers bind to the lagging strand and synthesize short fragments of DNA called Okazaki fragments.3 These fragments are later joined together by a series of enzymes to create an intact strand.
A problem does exist, however, in that the end of the lagging strand is not fully replicated, leaving an overhanging strand of DNA that is unpaired on the leading strand. This occurs every time DNA undergoes replication; it is called the “end replication problem,” because the DNA duplex becomes shorter by approximately 50 to 200 base pairs each time it undergoes replication. Continued shortening would be a serious problem if the chromosomes did not have protective caps at the ends, called telomeres. Telomeres protect the genomic DNA from degradation and shortening that could lead to mutations. Telomeric ends are repeated base pairs of TTAGGG that do not encode any genes.4 Once the telomere becomes critically shortened by continuous DNA replication, an as-yet unknown signal stops cell replication forever. This cell then enters a senescent state. It does not necessarily die at that point and may survive for a certain amount of time. Each cell type has a set number of divisions that it undergoes in its lifetime before it becomes senescent, and this fixed number defines one of the major theories of aging.5
Cells that require unlimited proliferative capabilities like cancer cells, germ-line cells, and some stem cells have acquired the ability to lengthen their telomeres. One of the telomeric ends has a 3′ segment that provides a template for the addition of individual bases. Telomere transferase (telomerase) is a ribonucleoprotein complex that uses an RNA template for synthesizing the telomeric DNA. This complex also has a catalytic subunit (telomerase reverse transcriptase [TRT]) that acts upon its own RNA template. The cells that have telomerase activity can maintain their telomeres and theoretically live forever.5
Protein Synthesis
Deoxyribonucleic acid must be converted to messenger RNA (mRNA), which is the working copy of the genetic information, so that it can then be translated into protein synthesis. Accessory proteins, called transcription factors, control the expression of different genes by up-regulating (turning on) or down-regulating (turning off) them. Transcription factors bind to specific areas on genes, called promoters and enhancers, in order to control gene expression. Once mRNA is made, it is sent out into the cytoplasm, and the ribosomes (translators) translate the mRNA into the amino acid sequences of protein. It was once thought that one gene made one protein; now it is known that one gene can make more than one protein via post-translational modifications. The central dogma of molecular biology is that through DNA transcription and RNA translation, proteins are synthesized.
Codons are the groups of nucleotides that code for amino acids. All codons contain three successive nucleotides. Many amino acids are specified by more than one codon. There are a possible 64 combinations of the three bases, and 61 of these are used to code for specific amino acids. The other three combinations (UAA, UAG, UGA) code for stop signals, which act like a period at the end of a sentence. The codon AUG encodes the start codon and also codes for methionine, so it has two potential jobs. The AUG combinations that start polypeptides are all closely preceded by a purine-rich sequence that distinguishes it from the AUG that codes for methionine.2
Mutations are changes in the coding sequence of a gene. Mutations are important, because they are responsible for inherited disorders and are the source of phenotypic variation (i.e., natural selection).
Techniques Common to All Assays
Gel Electrophoresis
Gel electrophoresis is the most widely used assay method in molecular biology. Photographs of gel electrophoresis are often shown in publications to evaluate, analyze, and demonstrate the results of an experiment. The concept of electrophoresis utilizes the premise that positive charges attract negative charges and vice versa. Both DNA and RNA are negatively charged and move at the same speed toward a positive electrode when dissolved in solution. The sizes of the DNA or RNA molecules can be used to identify them when they are run through a gel matrix. A gel is a semisolid meshwork of cross-linked polymers made of agarose. Gels have gaps in the meshwork that allow DNA or RNA molecules to migrate through them based on size. The smaller molecules migrate more quickly and easily, while larger molecules migrate more slowly. Small fragments of DNA are often run through polyacrylamide or sieving agarose gels. These types of gels have very small pore sizes that allow better resolution of the smaller molecules.
Visualization of DNA or RNA molecules is often done after they are run through gels. The most common method of visualizing DNA or RNA is by staining the agarose gel with ethidium bromide, which intercalates itself throughout the nucleic acid structure.3 Nucleic acids that have ethidium bromide bound to them fluoresce bright orange under ultraviolet light and can be photographed. Radioactive labeling is another method used to visualize nucleic acids.
Hybridization
The principles of hybridization, also called annealing, are often used in molecular biology to identify DNA or RNA products. Hybridization is the binding of a labeled probe to membrane-bound DNA or RNA. Nucleic acid hybridization is affected by the size of the nucleic acids being hybridized, the overall number of guanine and cytosine molecules (G + C content), the concentration of the nucleic acids, and how close the target matches the probe. Probes are molecules that are tagged with radioactivity, fluorescent agents, or digoxigenin, and they are used to detect another molecule. If the probe used is not 100% compatible with the target, it is said to be mismatched. Numerous molecular techniques use hybridization, including Southern and Northern blot analysis and the polymerase chain reaction (PCR).
Techniques for DNA Assays
Polymerase Chain Reaction
The PCR assay was invented in 1987, and today it is the most widely used molecular technique in all of science.2 Its applications include clinical diagnostics, genetic and forensic analyses, and genetic engineering. On the most basic level, PCR is used to amplify specific DNA sequences. Most PCR assays are based on known sequences of DNA or segments of genes.
A component of a PCR is the template, which is DNA that has the target sequence within it (e.g., the patient’s sample). Only a trace amount of template is required for a PCR. A pair of primers is also necessary. A primer is a short segment of single-stranded DNA that matches a portion of the target sequence. Most PCRs use a forward primer, which is located at the 5′ end of the target sequence, and a reverse primer, which is located at the 3′ end of the target sequence on the complementary strand of DNA. Next, a heat-resistant enzyme drives the reaction. The most commonly used enzyme is derived from Thermus aquaticus and is called Taq polymerase. Other necessary ingredients for the PCR are a large supply of nucleotides for the creation of the DNA strands and an appropriate buffer.
The mixture of these ingredients is placed into a thermocycler that is programmed to rapidly change temperature for a preset number of cycles (30 to 50). The mixture is heated to 90°C in order to denature or separate the template’s double-stranded DNA. Then the temperature is dropped to between 50° and 65°C, which allows the primers to bind or anneal to the complementary sequence on the template DNA (i.e., the target sequence). Lastly, the temperature is raised to 70°C for 1 to 2 minutes, which allows the Taq polymerase to elongate the new DNA strands beginning at the primers. This elongation always occurs in a 5′ to 3′ direction. Each cycle doubles the amount of DNA present, and by the end of 30 cycles, there are >1 million copies of double-stranded DNA present. This PCR product is then visualized using agarose gel electrophoresis stained with ethidium bromide [Figure 2].
Cloning
In the present context, cloning a gene or a piece of that gene refers to obtaining the DNA that makes up that particular gene, using molecular techniques. The DNA is then used in subsequent experiments such as PCR.2 For example, if the gene of interest is not known for the species being assayed, then the most common approach used to assess a segment of DNA is to align the sequences of this gene with those of two or three closely related species to compare their homology. In areas that are exact, short sequences are chosen as the primers in the forward and reverse direction to encompass an area of a few hundred base pairs. The DNA of the species of interest is used as the template, and the chosen primers are added to the mixture, which also includes nucleotides (for making the newly created DNA), Taq polymerase (which drives the reaction), and other necessary ingredients. Most initial experiments use a lower annealing temperature and a higher number of cycles, which allows for promiscuous binding of the primers to areas that are not exactly correct—resulting in multiple bands on the agarose gel. Even though a band is expected at a certain point/size, it is also possible to amplify garbage (i.e., material unrelated to desirable material being amplified). In order to decipher the band of DNA product that the assay is seeking, either Southern blot hybridization or sequencing of that DNA product can be used. In a Southern blot analysis, a DNA probe is designed within the expected sequence that does not include either of the primer sequences. This probe is then used to confirm or refute the DNA products amplified by PCR. A more common alternative is to sequence the PCR product. This is explained further later.
Southern Blot Analysis
Southern blot analysis is used to detect DNA that has been immobilized on a nylon membrane by using a DNA probe.2 The DNA of interest is separated using agarose gel electrophoresis. The DNA of interest may be a PCR product, a large piece of DNA (e.g., plasmid), or genomic DNA. Plasmid DNA is a circular molecule of double-stranded helical DNA that is used in gene cloning. Genomic DNA is chromosomal DNA isolated from a eukaryotic cell. Since plasmid and genomic DNA strands are very large, they cannot be run through an agarose gel without being cut into smaller pieces, which is performed by restriction enzyme digestion.2 Restriction enzymes are made by bacteria, and they cut DNA strands at specific sites. Once the fragmented DNA has undergone agarose gel electrophoresis and detection with ethidium bromide, the DNA from the gel is transferred to a nylon membrane, and then the membrane is incubated with a salt solution containing a labeled DNA probe. The DNA probe only binds to DNA fragments on the membrane that are similar to the probe, and the probe can then be detected using autoradiography. An example of samples evaluated by Southern blot analysis is seen in Figure 3.
Sequencing of DNA
Nucleotides (A, T, C, G) are the letters of the genetic alphabet. A DNA sequence is the specific order in which the nucleotides are “spelled out” to create genes. Therefore, DNA sequencing is a technique that elucidates the order of the nucleotides that make up a piece of DNA that has been amplified by PCR or by cloning. Currently, a sequence of =700 nucleotides can be obtained from a sequencing reaction. Larger pieces of DNA must be cut into smaller pieces and then sequenced. Two methods are commonly used to determine DNA sequences. The first is the enzymatic dideoxy (Sanger) method, and the second is the chemical (Maxam-Gilbert) method.6
Real-Time PCR
Real-time PCR is used to quantify the number of copies of DNA or mRNA present in a sample. This technology is especially useful in diagnostic medicine, because infectious agents causing clinical disease can be distinguished from normal or background flora. The amplification function of the PCR has a sigmoidal shape that becomes saturated after 20 to 40 cycles, because the conditions become less optimal with each cycle [Figure 4]. The saturation point is the flat plateau of the sigmoid curve, and all samples eventually reach this point even though there are differing amounts of DNA present in each sample. A band seen on an agarose gel following traditional PCR is representative of this plateau (i.e., the entire number of cycles of the PCR). Expression levels of the DNA sequence of interest are compared during the exponential phase of the PCR, long before saturation of the reaction occurs.
The way in which real-time PCR monitors the amount of DNA being amplified is by using fluorescent dyes that bind only to double-stranded DNA. The fluorescence increases with each cycle as the DNA amplification increases. A computer program uses an algorithm to analyze the curves for each sample as well as for the standard controls. Following mathematical calculations, data for each sample include the copy number of DNA, which can then be used to achieve a diagnosis. In addition, the copy number of DNA is directly related to cycle number. With a higher copy number, the sigmoid curve reaches the plateau at a lower cycle number. For example, in Figure 4, sample A has a higher expression than sample B, because the cycle number where it crossed the threshold of the plateau (saturation point) is lower.
Techniques for RNA Assays
Virtually all mature mRNAs include a long 3′ polyadenosine sequence (AAA repeated 200 to 300 times). This 3′ polyadenosine tail is thought to protect the young mRNA from being destroyed by RNA degrading enzymes. It is also an exceedingly useful feature that is exploited in some molecular techniques.
Northern Blot Analysis
Northern blot analysis is the most sensitive technique for detecting mRNA expression levels.7 The RNA is separated by size on a denaturing, formaldehyde, agarose electrophoretic gel. Since RNA has a negative charge, it migrates toward the cathode when a current is applied to the gel. Similar to other forms of electrophoresis, mobility of the molecule in the gel is inversely proportional to its size, such that large fragments do not migrate as rapidly as shorter fragments. After separation by electrophoresis, the RNA is transferred to a nitrocellulose membrane, and the RNA is cross-linked to the membrane by ultraviolet (UV) irradiation.
Specific RNA sequences can be detected on the blotting membrane by hybridization (i.e., by binding of a labeled probe to the membrane-bound RNA). The probe can be labeled by coupling to a protein such as digoxigenin, which in turn can be detected using an antidigoxigenin antibody conjugated to alkaline phosphatase that converts a color substrate. This staining provides an amplified signal. Alternatively, the probe can be radiolabeled and detected by autoradiography. The results of hybridization reveal the number, size, and abundance of transcripts, with the quantity of the product being approximately proportional to the amount of the specific RNA present. The concentration of a housekeeping gene, such as 18s-rRNA, can be measured on the same Northern blot to evaluate the relative signal strength. While this technique is time-consuming and requires significant amounts of RNA for analysis, it is still commonly used, because it is extremely quantitative and, therefore, provides an accurate assessment of mRNA levels. One example of how this technique has been used in veterinary medicine can be seen in the paper by Green (et al.).8 The researchers examined the mRNA from dogs affected with cardiomyopathy to determine if genetic mutations associated with the disease resulted in abnormal expression of mRNA.
Reverse-Transcriptase PCR
The amount of RNA to be measured is often well below the threshold detected by conventional Northern blot analysis. Reverse-transcriptase PCR (RT-PCR) can be used to assess lower levels of mRNA expression in samples by relying on the amplification of a DNA template via PCR. Prior to amplification through PCR, the single-stranded mRNA is converted into double-stranded complementary DNA (cDNA). To make a DNA copy of the mRNA, the enzyme, reverse transcriptase, is used, as well as a primer that binds to the polyadenosine tail of mRNA. Once the cDNA is made, the RNA is degraded by the enzyme, RNAse H, and DNA polymerases are used to build the complementary strand. The result is a collection of DNA molecules that represents all the specific mRNA contained within the cell. Similar to PCR of DNA, primers are designed for a specific gene, and they allow exponential amplification by PCR. The resulting PCR product is subjected to agarose gel electrophoresis to separate the PCR products by size. Ethidium bromide intercalates into the DNA and can be visualized using UV light.
An important advantage to RT-PCR is that only a small amount of biological sample is necessary to determine the mRNA level. The limitation of this method for determining gene expression is that RT-PCR combined with visualization using ethidium bromide is only semiquantitative, owing to the fact that the PCR is linear only within a range of cycles. While RT-PCR is not well suited for the quantitative comparison of mRNA expression, the technique is useful in screening tissues for the expression of a gene. For example, RT-PCR-based assays are used in the clinical setting to detect viruses.
Real-Time RT-PCR
Real-time RT-PCR is the same technique as real-time PCR; however, the starting material is mRNA rather than DNA. If comparisons regarding gene expression levels are made in the saturation phase, large initial differences in mRNA expression may be lost; therefore, it is necessary to monitor the time kinetics of amplification. The technology of real-time RT-PCR developed out of the need for a quantitative technique to determine gene expression levels in small samples.
Measurement of mRNA for a particular gene by real-time RT-PCR is not fundamentally different from RT-PCR. Both techniques require mRNA initially to be reverse transcribed to create cDNA. The cDNA is then subjected to PCR. The main difference is that the instrument where the PCR is carried out is both a thermocycler and a detection device. The reaction mixture contains a fluorescent probe that intercalates into the DNA during synthesis. At the end of each PCR cycle, the amount of fluorescence is assessed. Expression levels of analyzed genes of interest are compared during the exponential phase of the PCR, long before saturation of the reaction occurs. Based on an algorithm, a threshold level of fluorescence must be reached for a mRNA to be considered expressed, and the threshold is expressed as a cycle number. Expression of mRNA is directly related to cycle number. With higher relative expression, a lower cycle number is required to reach the threshold or plateau [Figure 4]. The copy number of the mRNA gene of interest can be standardized against the copy number of a housekeeping gene, or it can be determined from a standard curve. Real-time RT-PCR has been used in the authors’ laboratory to determine changes in gene expression during corneal wound healing. Different treatments, such as tetracycline, have been found to alter the time and duration of expression of particular genes important for cell migration.9
Microarray
A microarray is a tool for analyzing the relative expression level of a gene by determining the amount of mRNA present. A microarray can be considered a reverse Northern blot technique. In a Northern blot assay, probes are allowed to search for and bind to the appropriate RNA that has been bound to a membrane. In a microarray, probes for hundreds to tens of thousands of transcripts are immobilized on the surface of a slide or chip, and a labeled RNA solution is then added. The technology allows DNA to be spotted or synthesized directly onto a solid surface in an ordered, predetermined fashion at an extremely high density, facilitating quick and efficient analysis of the expression of thousands of genes in a single experiment.10
As with both RT-PCR and real-time RT-PCR, mRNA is isolated from a specific tissue or cell line and used to generate cDNA. Unlike the other techniques, the cDNA is labeled with different fluorescent tags (typically red and green) so that the samples can be differentiated upon imaging. Next, the cDNA mixture is hybridized with the array, probing the target cDNA for the presence of known genetic sequences that are anchored to the array slide. To detect the target cDNA that reflects gene expression in an experimental sample, the cDNA solution is hybridized with the array, causing the fluorescent-labeled molecules to bind to the sites on the array with DNA sequences that correspond to the genes expressed in each cell.10
After the hybridization step is complete and the target cDNA is bound to the array, the microarray is placed in a scanner that consists of specifically designed lasers, a microscope, and a detector. The fluorescent tags are excited by the laser, and the intensity of the red or green signal corresponds to the amount of DNA present in each sample. Spots that are yellow are equally expressed in the sample. The microscope and detector generate a digital picture of the array, and the intensity of the red-yellow-green spectrum is analyzed for each spot on the microarray.
Many genes show low expression levels, and the respective signals may be overwhelmed by background noise. As a result, few conclusions can be drawn; therefore, it is necessary to set a threshold level for transcripts that are included in the analysis. Furthermore, it is important to repeat the analysis with samples that have undergone the same processing (e.g., RNA extraction, cDNA synthesis) to ensure that the signal is consistent between samples and experimental runs. It should be noted that while microarray is a good technique to analyze global changes between two samples, it must be validated by other molecular biology techniques.
Several major categories of diagnostic questions can be addressed using microarrays as a starting point. These include scanning for the presence of a mutation in a specific disease gene, scanning for expression profiles that differentiate molecular subcategories of disease, scanning large-scale changes in DNA copy number that can accompany some acquired diseases, and the identification of markers associated with disease.11–13 Perhaps the most widely used application of microarrays in animal medicine is the rapid and accurate detection and genotyping of a range of animal pathogens, including bacteria, fungi, and viruses.14,15
In Situ Hybridization
In situ hybridization can be used to localize the expression of different transcripts in thinly sliced tissue sections.7 The principles underlying this technique are the same as those described above for Northern blotting. In the case of in situ hybridization, DNA or RNA probes (riboprobes) containing either radioactive or nonradioactive labels are used. In general, RNA probes offer increased sensitivity and specificity compared to DNA probes, although RNA probes are more difficult to produce and more labile than DNA probes. The oligonucleotide probe is generated to the mRNA of interest and is hybridized against the histological tissue section, which allows the cellular location of the transcript to be visualized. The power of this technique is that the location of mRNA expression can be visualized in the context of the whole tissue. In situ hybridization allows for an estimation of the level and location of expression of mRNA. An example of the technique is shown in Figure 5.
Protein Techniques
Western Blot Analysis
Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) is a technique that is used to separate proteins based on their molecular weight.7 The detergent SDS has a negative charge, and when added to the sample, it binds hydrophobic amino acid residues. Proteins bind proportionally to the same amount of SDS; therefore, the charge to mass ratio of all proteins is equivalent, and all SDS-protein complexes are negative. Protein samples are prepared either under reducing conditions where ß-mercaptoethanol is added and the sample is boiled, or under nonreducing conditions where ß-mercaptoethanol and heat are omitted. The specific sample preparation conditions are determined by experimental necessity.
Once the samples have been prepared for electrophoresis, they are loaded on the gel and an electrical field is applied. The rate of migration of the protein is inversely proportional to the logarithm of its molecular weight—meaning, larger proteins move slower through the gel because they encounter more resistance. Protein bands that result from differential migration can be stained for visualization with Coomassie blue or silver, and they can be compared against reference proteins with known molecular weights run on the same gel. While this is a powerful tool that allows examination of global changes in protein levels from samples, it lacks specificity.
Western immunoblotting utilizes antibody specificity to ascertain information about the levels of a particular protein in samples.7 In this technique, after proteins are separated on an SDS-PAGE gel, they are transferred to a membrane by applying a current across the gel and membrane in such a way that the proteins migrate from the gel to the membrane. After the proteins have been transferred, the membrane is incubated with an antibody (primary antibody) manufactured against the protein of interest. Following incubation with the primary antibody, the membrane is incubated with a secondary antibody that is conjugated to horseradish peroxidase and is capable of recognizing the primary antibody. For example, if the antibody against protein “X” was raised in a rabbit, the second antibody used would be an anti-rabbit horseradish peroxidase-conjugated antibody. This highly specific antibody-to-protein binding is detected using a peroxidase-dependent luminescence-generating system followed by exposure to X-ray film. The bands that appear reflect the size and relative amount of the studied protein. As seen in Figure 6, specific bands appearing at the appropriate weight/point confirm the protein of interest is present. As an internal control, the blots are stripped and re-probed with an antibody for a housekeeping gene (e.g., ß-actin). If the bands for the housekeeping gene have the same relative intensity, the protein concentrations in each lane are equal. The relative quantity of the protein of interest can then be determined. For example, in Figure 6, all lanes have approximately equal protein concentrations (based on the ß-actin bands), and lanes 1 and 2 have less protein (pAkt) present than lanes 3 and 4.
Western immunoblotting is an extremely powerful tool for examining the presence or absence of proteins in a sample and for comparing samples ascertained under different conditions. For example, western immunoblotting has been used to determine which proteins are overexpressed during cataract formation.16–18 The major limitation of Western immunoblotting is that it is semiquantitative.
Immunohistochemical Staining
Immunohistochemistry is used to localize specific proteins in tissues (immunohistochemistry) or cells (immunocytochemistry).4 Immunohistochemistry often involves special staining techniques used by pathologists to further characterize tissue specimens. Because immunohistochemistry can be applied to formalin-fixed or frozen tissues and cells, it is easy to use after fine-needle aspiration, biopsy, or necropsy examination. It is not a quantifiable technique; however, if numerous samples are stained at the same time (with appropriate positive and negative controls), these samples can be compared, which makes the technique semiquantitative.
Immunohistochemistry has several requirements, including tissue or cells immobilized on a microscope slide, antibody against the protein of interest (primary antibody), appropriate buffers and alcohols, a secondary antibody that has been biotinylated, enzyme-conjugated streptavidin or another conjugate, a chromophore, and a counterstain. As with Western blotting techniques, the secondary antibody is typically immunoglobulins made against the species of the primary antibody. The enzyme conjugate can be horseradish peroxidase, alkaline phosphatase, glucose oxidase, or immunogold. The chromophore is a color-producing substrate system, so it gives the reaction a color. Typically, diaminobenzidine (DAB) or 3-amino-9-ethyl carbazole (AEC) is used as the chromophore. Diaminobenzidine imparts a brown color to positive sites, and AEC imparts a red color to positive sites. Depending on the situation, one chromophore may be better than the other. For example, in samples that contain melanin, AEC is superior, because it provides better contrast against the pigment present in the sample. The counterstain commonly used is hematoxylin, because it stains the nucleus of the cells blue, and nuclear proteins will have a brownish-blue hue. The stained samples can be permanently mounted for future evaluation.
Immunofluorescent staining is identical to immunohistochemistry, except that the primary or secondary antibody is labeled with a fluorescent tag. A fluorescent microscope is necessary to evaluate these samples, and often fluorescence does not last for more than a few weeks to months. In Figures 7A and 7B, one tissue sample is stained using immunohistochemistry (A), and the other is an example of immunofluorescence (B).
Enzyme-Linked Immunosorbent Assay
Enzyme-linked immunosorbent assay (ELISA) is a biochemical method used to detect the presence of an antigen or antibody in a sample. It has numerous advantages, including its simplicity, speed (a few hours), sensitivity, and relatively low cost. Also with ELISA, many samples can be evaluated at one time, there is potential for automation, and numerous kits are commercially available.19 Two antibodies are typically used; one is specific to the antigen or antibody of interest in the sample, and the other is coupled to an enzyme (enzyme-linked) that causes a signal or color change that is quantified by a spectrophotometer.19 Three main methods form the basis for all ELISAs: the direct, indirect, and sandwich methods.5
With the direct ELISA method, antigen is added to a microtiter plate and is passively adsorbed to the plate (solid phase). Following incubation, antibodies specific for the antigen are labeled with an enzyme conjugate and are added to the microtiter plate. Once the enzyme conjugate-labeled antibodies have bound to the antigens on the plate, excess unbound antibodies are washed away, and a substrate or chromophore solution is added that catalyzes the enzyme conjugate to produce a color change. This color change is quantified by a spectrophotometer.
The indirect ELISA method is similar to the direct method, but antibodies specific for the antigen bound to the microtiter plate are added to wells and incubated, and then excess unbound antibodies are washed away. Antibodies labeled with enzyme conjugate directed against the species of the original antibody are then added and incubated, and excess is washed away. A substrate or chromophore is added that catalyzes the enzyme conjugate to produce a color change, which is quantified by a spectrophotometer.
The sandwich ELISA can be divided into two types: the direct or indirect sandwich ELISA. The first four steps of both techniques are similar. Antibodies become passively attached to the solid phase (microtiter plate) and are called capture antibodies. These antibodies capture or bind the antigens added at the next step. Following incubation and washing, an antibody-antigen complex is attached to the solid phase, and the antigen is said to be trapped. With the direct sandwich ELISA technique, enzyme-conjugated antibodies directed against the antigen are added, and this completes the sandwich. Unbound conjugate is washed away, and a substrate or chromophore solution is added that catalyzes the enzyme conjugate to produce a color change, which is quantified by a spectrophotometer. With the indirect sandwich ELISA method, detection antibodies not labeled with an enzyme are then added to the antibody-antigen complex described above. Following incubation and washing, the detection antibodies are themselves identified with an antispecies enzyme conjugate that produces a color change. This color change is quantified by a spectrophotometer.
The advantage of ELISA-produced data is that they are generated relatively quickly and can be evaluated easily. This method is often used to compare numerous test and control samples in duplicate or triplicate in a quantitative manner. For example, the ELISA has been used to compare protein alterations in lenses exposed to an oxidative stressor (tertiary butyl hydroperoxide) for various periods of time.20
All ELISAs must be standardized and validated in order to achieve a level of confidence in the results. Standardization and validation procedures include optimization of the reagents and the protocol, determination of the repeatability of the assay, and determination of the sensitivity and specificity of the assay.19
Flow Cytometry
Flow cytometry is another method of evaluating cells. It is most commonly used to count cells, evaluate cell-surface phenotypic markers, or to analyze cell cycles, cytokine production, apoptosis, and cell viability.21 It is only useful for cells that are suspended and not clumped together, because the cells are passed within a stream of fluid in front of a light beam. Forward scatter and side scatter, as well as fluorescence detectors, are used in flow cytometry. The molecules to be detected on the cell scatter the light, and fluorescent chemicals in the particle may be excited to emit light at a lower frequency than the light source. The combination of fluorescence and scattered light is detected and analyzed by the machine, resulting in data shown as a one-dimensional histogram or as two-dimensional dot plots. An example of a blood sample evaluated by flow cytometry is seen in Figure 8. Newer software programs also allow three-dimensional analysis.
Conclusion
Molecular biological techniques, especially PCR assays and ELISA, have become powerful diagnostic tools in veterinary medicine. Another interesting application of these techniques is using biological samples from clinical animals to investigate the mechanisms of disease. Information obtained from animals allows the basic research models to be validated as acceptable models for studying specific disease processes. Ultimately, results from translational research may enhance disease prevention and lead to more effective and specific medical treatments.
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Citation: Journal of the American Animal Hospital Association 42, 5; 10.5326/0420326
Double helix structure of deoxyribonucleic acid. (Image courtesy of Mr. Timothy Vojt.)
Agarose gel electrophoresis of 10 polymerase chain reactions (PCRs). The bands seen in lanes 1 and 13 are molecular-weight markers that allow estimation of the size of the PCR products. Bands in lanes 2, 3, and 5 are positive for the reactions performed. The bands in lanes 6, 7, and 10 are positive controls for the samples run in lanes 2, 3, and 9. The reactions run in lanes 4, 8, and 9 are negative.
Illustration of a Southern blot analysis of genomic deoxyribonucleic acid (DNA) extracted from seven samples. The analysis was run on an agarose gel, transferred to a nylon membrane, and then a DNA probe was hybridized with the immobilized DNA on the membrane. The bands indicate hybridization (binding) to various sites on the DNA in each sample.
Sigmoidal curves generated by real-time polymerase chain reactions (PCRs). Sample A has a higher expression level than sample B, because the cycle number (i.e., the point where the curve reaches the plateau threshold) is lower.
In situ hybridization is illustrated in this figure. Cells were processed appropriately, and a deoxyribonucleic acid probe was used in this experiment. The chromogen used was diaminobenzidine (DAB), so the positive brown staining denotes messenger ribonucleic acid in the nuclei of these cells.
In this Western immunoblot of four samples, the first two lanes have lighter bands than the last two lanes. Since the ß-actin and p-Akt bands are similar in intensity, the samples in lanes 1 and 2 have less expression of p-Akt than the samples in lanes 3 and 4. The ß-actin protein is often used as an assay loading control.
Examples of immunohistochemical (A) and immunofluorescent (B) staining. In 7A, the blue staining is the counterstain, and the brown staining is positive for the protein of interest in the sample. In 7B, the blue color is the counterstain, and the green staining is positive for the protein of interest in the sample.
A dot plot showing the results of flow cytometry. In the sample, 8% of the cells (upper left quadrant) stain positively for cluster differentiation (CD) 8 with phycoerythrin (PE, red); 23% of the cells (lower right quadrant) stain positively for CD4 with fluorescein-5-isothiocyanate (FITC, green); and 1% of the cells (upper right quadrant) stain positive for both CD4 and CD8. The cells represented in the bottom left quadrant are not staining for either CD4 or CD8. (Image courtesy of Ms. Sarah Leavall and Dr. Mary Jo Burkhard.)
