Practical Biochemistry II (C-506) 2021 PDF

Assiut University Faculty of Science Chemistry Department Practical Biochemistry II (C-506) For Biochemistry Professional Diploma’ Students Prepared by: Prof. Nagwa Abo El-Maali 2021 Tabl...

Assiut University Faculty of Science Chemistry Department Practical Biochemistry II (C-506) For Biochemistry Professional Diploma’ Students Prepared by: Prof. Nagwa Abo El-Maali 2021 Table of content Subject Page 1. Enzyme-linked Immunosorbent Assay, ELISA 3 1.1 Enzyme Immunoassay for the Quantitative Determination of Vitamin D 11 1.2 Enzyme Immunoassay for the Quantitative Determination of Human Ferritin 18 Concentration in Human Serum 2. Electrophoresis 24 2.1 Principles of electrophoresis 24 2.2 Gel electrophoresis 27 2.2.1 Polyacrylamide gel electrophoresis (PAGE) 29 2.2.1.1 Native PAGE 35 2.2.1.2 SDS-PAGE 36 2.2.2 Isoelectric focusing 38 2.2.3 Two-dimensional (2D) electrophoresis. 41 2.3 Agarose gel electrophoresis 43 2.4 Staining methods 43 2.5 Specific protein detection methods: Western blot 45 2.6 Typical examples of protein-separating gel electrophoresis 46 3. Polymerase Chain Reaction (PCR) 54 4. Real-time quantitative PCR (RT-PCR). 59 HCV Detection by Real-Time PCR 72 5. Complete Blood Count (CBC) 76 2 1. Enzyme-linked Immunosorbent Assay, ELISA The ELISA, or Enzyme-Linked ImmunoSorbent Assay, is an analytical biochemistry technique that uses antibodies to detect the presence of specific biomolecules (i.e. peptides, proteins, antigens and hormones) in a complex sample. These samples can be single proteins or complex mixtures like cellular lysates, it was first described by Engvall and Perlmann in 1971. The assay uses a solid-phase type of enzyme immunoassay (EIA) to detect the presence of a ligand (commonly a protein) in a liquid sample using antibodies directed against the protein to be measured. ELISA has been used as a diagnostic tool in medicine, plant pathology, and biotechnology, as well as a quality control check in various industries. In the simplest form of an ELISA, antigens from the sample to be tested are attached to a surface. Then, a matching antibody- antibodies are produced by living organisms as part of their immune response and recognize specific recognition sites (epitopes) on foreign molecules (antigens)- is applied over the surface so it can bind the antigen. This antibody is linked to an enzyme and then any unbound antibodies are removed. In the final step, a substance containing the enzyme's substrate is added. If there was binding the subsequent reaction produces a detectable signal, most commonly a color change. Performing an ELISA involves at least one antibody with specificity for a particular antigen. The sample with an unknown amount of antigen is immobilized on a solid support (usually a polystyrene microtiter plate) either non-specifically (via adsorption to the surface) or specifically (via capture by another antibody specific to the same antigen, in a "sandwich" ELISA). After the antigen is immobilized, the detection antibody is added, forming a complex with the antigen. The detection antibody can be covalently linked to an enzyme or can itself be detected by a secondary antibody that is linked to an enzyme through bioconjugation. 3 Between each step, the plate is typically washed with a mild detergent solution to remove any proteins or antibodies that are non-specifically bound. After the final wash step, the plate is developed by adding an enzymatic substrate to produce a visible signal, which indicates the quantity of antigen in the sample. ELISA can perform other forms of ligand binding assays instead of strictly "immuno" assays, though the name carried the original "immuno" because of the common use and history of development of this method. The technique essentially requires any ligating reagent that can be immobilized on the solid phase along with a detection reagent that will bind specifically and use an enzyme to generate a signal that can be properly quantified. In between the washes, only the ligand and its specific binding counterparts remain specifically bound or "immunosorbed" by antigen-antibody interactions to the solid phase, while the nonspecific or unbound components are washed away. Unlike other spectrophotometric wet lab assay formats where the same reaction well (e.g., a cuvette) can be reused after washing, the ELISA plates have the reaction products immunosorbed on the solid phase, which is part of the plate, and so are not easily reusable. Two of the most popular enzyme reporting systems are alkaline phosphatase (AP) and horse radish peroxidase (HRP). While both have been used success- fully in the biological community, up until now only the use of AP has been reported in the literature for ELISA applied to cultural heritage. Types There are many ELISA tests for particular molecules that use the matching antibodies. ELISA tests are broken into several types of tests based on how the analytes and antibodies are bonded and used. The major types are described: 4  Direct ELISA The steps of direct ELISA follows the mechanism: A buffered solution of the antigen to be tested for is added to each well (usually 96-well plates) of a microtiter plate, where it is given time to adhere to the plastic through charge interactions. A solution of nonreacting protein, such as bovine serum albumin or casein, is added to each well in order to cover any plastic surface in the well which remains uncoated by the antigen. The primary antibody with an attached (conjugated) enzyme is added, which binds specifically to the test antigen coating the well. A substrate for this enzyme is then added. Often, this substrate changes color upon reaction with the enzyme. The higher the concentration of the primary antibody present in the serum, the stronger the color change. Often, a spectrometer is used to give quantitative values for color strength. The enzyme acts as an amplifier; even if only few enzyme-linked antibodies remain bound, the 5 Direct ELISA diagram enzyme molecules will produce many signal molecules. Within common-sense limitations, the enzyme can go on producing color indefinitely, but the more antibody is bound, the faster the color will develop. A major disadvantage of the direct ELISA is that the method of antigen immobilization is not specific; when serum is used as the source of test antigen, all proteins in the sample may stick to the microtiter plate well, so small concentrations of analyte in serum must compete with other serum proteins when binding to the well surface. The sandwich or indirect ELISA provides a solution to this problem, by using a "capture" antibody specific for the test antigen to pull it out of the serum's molecular mixture. ELISA may be run in a qualitative or quantitative format. Qualitative results provide a simple positive or negative result (yes or no) for a sample. The cutoff between positive and negative is determined by the analyst and may be statistical. Two or three times the standard deviation (error inherent in a test) is often used to distinguish positive from negative samples. In quantitative ELISA, the optical density (OD) of the sample is compared to a standard curve, which is typically a serial dilution of a known-concentration solution of the target molecule. For example, if a test sample returns an OD of 1.0, the point on the standard curve that gave OD = 1.0 must be of the same analyte concentration as the sample. The use and meaning of the names "indirect ELISA" and "direct ELISA" differs in the literature and on web sites depending on the context of the experiment. When the presence of an antigen is analyzed, the name "direct ELISA" refers to an ELISA in which only a labeled primary antibody is used, and the term "indirect ELISA" refers to an ELISA in which the antigen is bound by the primary antibody which then is detected by a labeled secondary antibody. In the latter case a sandwich ELISA is clearly distinct from an indirect ELISA. When the "primary" antibody is of interest, e.g. in the case of immunization analyses, this antibody is directly detected by the secondary antibody and the term "indirect ELISA" applies to a setting with two antibodies. 6  Sandwich ELISA A "sandwich" ELISA is used to detect sample antigen. The steps are: 1. A surface is prepared to which a known quantity of capture antibody is bound. 2. Any nonspecific binding sites on the surface are blocked. 3. The antigen-containing sample is applied to the plate, and captured by antibody. 4. The plate is washed to remove unbound antigen. 5. A specific antibody is added and binds to antigen (hence the 'sandwich': the antigen is stuck between two antibodies). This primary antibody could also be in the serum of a donor to be tested for reactivity towards the antigen. 6. Enzyme-linked secondary antibodies are applied as detection antibodies that also bind specifically to the antibody's Fc region (nonspecific). 7. The plate is washed to remove the unbound antibody-enzyme conjugates. 8. A chemical is added to be converted by the enzyme into a color or fluorescent or electrochemical signal. 9. The absorbance or fluorescence or electrochemical signal (e.g., current) of the plate wells is measured to determine the presence and quantity of antigen A sandwich ELISA. (1) Plate is coated with a capture antibody; (2) sample is added, and any antigen present binds to capture antibody; (3) detecting antibody is added and binds to antigen; (4) enzyme-linked secondary antibody is added and binds to detecting antibody; (5) substrate is added and is converted by enzyme to detectable form. 7 The image above includes the use of a secondary antibody conjugated to an enzyme, though, in the technical sense, this is not necessary if the primary antibody is conjugated to an enzyme (which would be direct ELISA). However, the use of a secondary-antibody conjugate avoids the expensive process of creating enzyme-linked antibodies for every antigen one might want to detect. By using an enzyme-linked antibody that binds the Fc region of other antibodies, this same enzyme-linked antibody can be used in a variety of situations. Without the first layer of "capture" antibody, any proteins in the sample (including serum proteins) may competitively adsorb to the plate surface, lowering the quantity of antigen immobilized. Use of the purified specific antibody to attach the antigen to the plastic eliminates a need to purify the antigen from complicated mixtures before the measurement, simplifying the assay, and increasing the specificity and the sensitivity of the assay. A sandwich ELISA used for research often needs validation because of the risk of false positive results. Competitive ELISA A third use of ELISA is through competitive binding. The steps for this ELISA are somewhat different from the first two examples: 1. Unlabeled antibody is incubated in the presence of its antigen (sample). 2. These bound antibody/antigen complexes are then added to an antigen-coated well. 3. The plate is washed, so unbound antibodies are removed. (The more antigen in the sample, the more Ag-Ab complexes are formed and so there are less unbound antibodies available to bind to the antigen in the well, hence "competition".) 4. The secondary antibody, specific to the primary antibody, is added. This second antibody is coupled to the enzyme. 5. A substrate is added, and remaining enzymes elicit a chromogenic or fluorescent signal. 6. The reaction is stopped to prevent eventual saturation of the signal. Some competitive ELISA kits include enzyme-linked antigen rather than enzyme-linked antibody. The labeled antigen competes for primary antibody binding sites 8  with the sample antigen (unlabeled). The less antigen in the sample, the more labeled antigen is retained in the well and the stronger the signal. Commonly, the antigen is not first positioned in the well. For the detection of HIV antibodies, the wells of microtiter plate are coated with the HIV antigen. Two specific antibodies are used, one conjugated with enzyme and the other present in serum (if serum is positive for the antibody). Cumulative competition occurs between the two antibodies for the same antigen, causing a stronger signal to be seen. Sera to be tested are added to these wells and incubated at 37 °C, and then washed. If antibodies are present, the antigen- antibody reaction occurs. No antigen is left for the enzyme-labelled specific HIV antibodies. These antibodies remain free upon addition and are washed off during washing. Substrate is added, but there is no enzyme to act on it, so a positive result shows no color change. Reverse ELISA A fourth ELISA test does not use the traditional wells. This test leaves the antigens suspended in the test fluid. 1. Unlabeled antibody is incubated in the presence of its antigen (sample) 2. A sufficient incubation period is provided to allow the antibodies to bind to the antigens. 3. The sample is then passed through the Scavenger container. This can be a test tube or a specifically designed flow through channel. The surface of the Scavenger container or channel has “Scavenger Antigens” bound to it. These can be identical or sufficiently similar to the primary antigens that the free antibodies will bind. 4. The Scavenger container must have sufficient surface area and sufficient time to allow the Scavenger Antigens to bind to all the excess Antibodies introduced into the sample. 5. The sample, that now contains the tagged and bound antibodies, is passed through a detector. This device can be a flow cytometer or other device that illuminates the tags and registers the response. 9  This test allows multiple antigens to be tagged and counted at the same time. This allows specific strains of bacteria to be identified by two (or more) different color tags. If both tags are present on a cell, then the cell is that specific strain. If only one is present, it is not. This test is done, generally, one test at a time and cannot be done with the microtiter plate. The equipment needed is usually less complicated and can be used in the field. Commonly used enzymatic markers The following are lists the enzymatic markers commonly used in ELISA assays, which allow the results of the assay to be measured upon completion. OPD (o-phenylenediamine dihydrochloride) turns amber to detect HRP (Horseradish Peroxidase), which is often used to as a conjugated protein. TMB (3,3',5,5'-tetramethylbenzidine) turns blue when detecting HRP and turns yellow after the addition of sulfuric or phosphoric acid. ABTS (2,2'-Azinobis [3-ethylbenzothiazoline-6-sulfonic acid]-diammonium salt) turns green when detecting HRP. PNPP (p-Nitrophenyl Phosphate, Disodium Salt) turns yellow when detecting alkaline phosphatase. Applications Because the ELISA can be performed to evaluate either the presence of antigen or the presence of antibody in a sample, it is a useful tool for determining serum antibody concentrations (such as with the HIV test or West Nile virus). It has also found applications in the food industry in detecting potential food allergens, such as milk, peanuts, walnuts, almonds, and eggs and as serological blood test for coeliac disease. ELISA can also be used in toxicology as a rapid presumptive screen for certain classes of drugs. There are ELISA tests to detect various kind of diseases, such as dengue, malaria, Chagas disease, Johne's disease. ELISA tests also are extensively employed for in vitro diagnostics in medical laboratories. The other uses of ELISA include: detection of Mycobacterium antibodies in tuberculosis detection of rotavirus in feces detection of hepatitis B markers in serum detection of hepatitis C markers in serum detection of enterotoxin of E. coli in feces detection of HIV antibodies in blood samples detection of SARS-CoV-2 antibodies in blood samples. These videos, we will show you the basics of how to perform an ELISA in your classroom laboratory. https://www.youtube.com/watch?v=ilbptHXFQ9Q https://www.youtube.com/watch?v=zR_xlV5v_f4 10  2. Electrophoresis 2.1. Principles of electrophoresis Electrophoresis is a method used to separate charged particles from one another based on differences in their migration speed. In the course of electrophoresis, two electrodes (typically made of an inert metal, e.g. platinum) are immersed in two separate buffer chambers. The two chambers are not fully isolated from each other. Charged particles can migrate from one chamber to the other (Figure 1). Fig.1 The principle of electrophoresis. By using an electric power supply, electric potential (E) is generated between the two electrodes. Due to the electric potential, electrons move by a wire between the two electrodes. More specifically, electrons move from the anode to the cathode. Hence, the anode will be positively charged, while the cathode will be negatively charged. Electrons driven to the cathode will leave the electrode and participate in a reduction reaction with water generating hydrogen gas and hydroxide ions. In the meantime, at the positive anode an oxidation reaction occurs. Electrons released from water molecules enter the electrode generating oxygen gas and free protons (which immediately form hydroxonium ions with water molecules). The amount of electrons leaving the cathode equals the amount of electrons entering the cathode. The two buffer chambers are interconnected such that charged particles can migrate between the two chambers. 24 These particles are driven by the electric potential between the two electrodes. Negatively charged ions, called anions, move towards the positively charged anode, while positively charged ions, called cations, move towards the negatively charged cathode. The two electrodes are immersed in two separate buffer chambers. The two chambers are connected such that charged particles can migrate from one chamber to the other. By using a power supply, electric potential difference is generated between the two electrodes. As a result, electrons flow from one of the electrodes, the anode, towards the other electrode, the cathode. Electrons from the cathode are taken up by water molecules of the buffer, resulting in a chemical reaction which generates hydrogen gas and hydroxide ions. In the other buffer chamber, water molecules transfer electrons to the anode an in another chemical reaction that generates oxygen gas and protons. (Protons are immediately taken up by water molecules to form hydroxonium ions.) As charged particles can migrate between the two chambers due to the electric potential difference, positive ions (cations) move towards the negatively charged cathode while negatively charged ions (anions) move towards the positively charged anode. Different ions migrate at different speeds dictated by their sizes and by the number of charges they carry. As a result, different ions can be separated from each other by electrophoresis. It is very important to understand the basic physics describing the dependence of the speed of the ion as a function of the number of charges on the ion, the size of the ion, the magnitude of the applied electric field and the nature of the medium in which the ions migrate. By understanding these basic relationships, the principles of the many different specific electrophoresis methods become comprehensible. The fundamental principle of electrophoresis is illustrated in Figure 1. The mathematical description of the force during electrophoresis is simple. An electric force Fe is exerted on the charged particle. The magnitude of the electric force equals the product of the charge q of the particle and the electric field E generated between the two electrodes: (1) Dimensions of the electric field E are defined either in newton/coulomb or volt/cm units. During electrophores is the magnitude of the electric field E is defined in volt/cm units. It can be easily calculated using the value of the voltage (volt) set by the electric power supply and the distance of the two electrodes (cm). 25 Particles having different electrophoretic mobility, i.e. those that migrate at different speeds in the same medium and electric field, can be separated by electrophoresis. In biochemical and molecular biological studies, the most typical charged molecules that are analysed and separated by electrophoresis are proteins and nucleic acids. Electrophoresis is always performed by using a special medium, most often a gel. The corresponding methods are therefore denoted as gel electrophoresis. 2.2 Gel electrophoresis The principle of electrophoresis does not assume any particular requirements about the nature of the liquid medium in which the ions are separated. Yet, in the great majority of currently used electrophoretic applications, the medium has a three-dimensional network structure, i.e. the medium is a gel. At the very beginning when the technique was invented, electrophoresis was performed without using a gel matrix. Charged particles were migrated in a homogeneous liquid phase. However, it soon became apparent that the use of a liquid medium raises at least three major difficulties. One is that the separation of different ions in an ordinary liquid is rather inefficient. It is so because a significant factor of an effective separation should be a marked size-dependent drag force exerted by the medium on the particles. Although even ordinary liquids do interfere with the migration speed of the particles in a size-dependent manner, this size dependence is quite moderate. The other big problem has a simple technical origin. In liquid phase, even very small levels of temperature inhomogeneity trigger convection that significantly compromises the resolution of the separation. Finally, in an ordinary liquid phase, the extent of diffusion is high and, in the typical timeframe of the generally slow electrophoresis experiments, diffusion decreases the efficiency of the separation. All three problems had been dealt with when, instead of ordinary liquids, gels were introduced as a medium. The gel provides a three-dimensional molecular network structure to the liquid medium. It prevents convections and also lowers the rate of diffusion. Moreover, perhaps the most dramatic advantageous effect of the gel is that it acts as a molecular sieve: it interferes only slightly with the 27 movement of small molecules, but drastically slows down the motion of large molecules. All gels are characterised by an average pore size. Molecules much smaller than the mean pore diameter are almost unaffected by the presence of the gel, while those that are larger than the pores practically do not migrate in the gel. When ions with sizes in the range of the pore size are migrated through the gel by electrophoresis, the gel exerts a pronounced size-dependent dragging force on them. As a consequence, the pore size distribution of the gel determines an operational size range in which different ions can be separated. Looking at this from the opposite point of view, each separation problem defines an optimal pore size to be applied. The gel has to fulfill several general criteria to be applicable for biochemical electrophoresis. It needs to be hydrophilic, chemically stable (should not participate in chemical reactions during electrophoresis), neutral (free of electric charges, otherwise it would act as an ion exchanger) and mechanically resistant (should not be too elastic or too rigid as such gels would be difficult to handle). Furthermore, as the separated ions (mostly proteins and nucleic acids) need to be visualised in the gel by some kind of staining procedure, the gel should be transparent, and should not strongly bind the dyes used for staining. Finally, and very importantly, the experimenter should be able to adjust the pore size during the preparation of the gel. The size range of molecules (ions) studied in molecular biology is extremely broad. No single gel- forming compound is known that could cover the entire corresponding range of applicable pore sizes. Two compounds are dominantly used for gel electrophoresis: polyacrylamide and agarose. Polyacrylamide gels typically provide much smaller pores than do agarose gels. The polyacrylamide gel is formed by the radical polymerisation of acrylamide monomers. This process alone would lead to very long polymer chains instead of a three-dimensional gel. The three-dimensional network is brought about by the incorporation of N,N'-methylenebisacrylamide into the polymerising chains, which results in crosslinks between the long chains. The polyacrylamide gel is held together by covalent bonds. The pore size of polyacrylamide gels can be adjusted via the concentration of the acrylamide monomer and the ratio of the crosslinking agent, N,N'-methylenebisacrylamide. The pore size of polyacrylamide gels corresponds to a relatively low value (compared to that of agarose gels). 28 Polyacrylamide gels are used typically for the electrophoresis of proteins and relatively small nucleic acids. In comparison, the agarose gel is formed via non-covalent interactions between long polysaccharide chains. The pore size of agarose gels is much larger than that of acrylamide gels. Accordingly, agarose gels are used typically for the electrophoresis of large nucleic acids. The pore size of the agarose gel can be controlled via the concentration of the agarose solution. As the interaction between agarose molecules is non-covalent, the gel is formed by a physical (in contrast to a chemical) process. A suspension of agarose is heated up until the system reaches a sol state and then it is left to cool down to room temperature to reach the gel state. The following sections review the various polyacrylamide- and agarose-based electrophoresis methods. 2.2.1 Polyacrylamide gel electrophoresis (PAGE) PAGE method in general Polyacrylamide gels can be used for the separation and analysis of proteins and relatively small nucleic acid molecules. For example, when it was first invented, Sanger's DNA sequencing method applied PAGE to separate linear single-stranded DNA molecules based on their length. The resolution of the PAGE method is so high that, in the size range of about 10-1000 nucleotide units, it is capable of separating DNA molecules that differ in length only by a single monomer unit. In the case of single-stranded DNA, individual molecules are separated solely based on their length. This is due to the fact that, in the case of DNA (or RNA), the number of negative charges is a simple linear function of the number of monomer units (i.e. the length of the molecule). In other words, the specific charge (number of charges per particle mass) is invariant, i.e. it is the same for all DNA molecules. It is so because each monomer unit has one phosphate moiety that carries the negative charge. When an appropriate denaturing agent, such as urea, is added to the DNA sample and the gel is heated, the shape of the varying-length linear DNA molecules becomes identical. As a consequence, denatured molecules will be separated exclusively based on their size. (We will see the same principle at the SDS-PAGE method that separates denatured proteins almost exclusively based on their size (molecular weight)). 29 There are several PAGE methods (SDS-PAGE, isoelectric focusing, 2D PAGE) that can be applied mostly for the separation of proteins based on distinct molecular properties. At a given pH, different proteins carry different amounts of electric charge. Moreover, different proteins have different shapes and sizes, too. Consequently, during electrophoresis, proteins are separated by a complex combination of their charge, shape and size. PAGE separation of proteins provides high resolution. However, as three independent molecular properties simultaneously influence electrophoretic mobility, it will provide limited room for precise interpretation. For example, when two proteins are compared, it remains hidden what makes one of them migrate faster: a larger number of electric charges, a smaller size, or a more spherical shape. Nevertheless, even the simplest PAGE method, which will be referred to as native PAGE, provides many particular advantages. In order to increase the analytical applicability of the PAGE technology, several variations of the method have been established to separate proteins based on a single molecular property. As we will see, SDS-PAGE separates proteins based primarily on molecular weight, while isoelectric focusing separates proteins exclusively based on isoelectric point. In the presence of suitable initiator and catalyst compounds, acrylamide can readily polymerise in a radical process. (Acrylamide is harmful by inhalation or skin contact, and thus it should be handled with care.) This reaction would lead to very long polyacrylamide chains, yielding a highly viscous liquid instead of a gel. As already mentioned, these long chains need to be cross-linked to form a three-dimensional network. This is achieved by mixing N,N'-methylenebisacrylamide into the acrylamide solution. In essence, N,N'-methylenebisacrylamide is composed of two acrylamide molecules covalently interconnected via a methylene moiety. When, during the polymerization reaction, the acrylamide groups of N,N'-methylenebisacrylamide molecules become incorporated in the long polyacrylamide chains, cross-links are formed between the polyacrylamide chains leading to a gel (Figure 2). In the course of electrophoresis, ions (proteins or nucleic acids) are separated in this gel. 30  Figure 2. Molecular structure of the polyacrylamide gel. The three-dimensional molecular network comes into being by a radical polymerisation of acrylamide monomers and cross-linking N,N'-methylenebisacrylamide components. Without any modification, polyacrylamide electrophoresis separates macromolecular ions based on a combination of charge, size and shape. Size (and shape) separation is due to the molecular sieving property of the gel. The size range in which molecules can be separated is dictated by the average pore size of the gel. In the case of polyacrylamide gels, this can be controlled through the concentration of the acrylamide monomer and the proportion of the cross-linking N,N'- methylenebisacrylamide. The acrylamide concentration can be set in the range of about 4-20 % as this is the range in which the mechanical properties of the gel are appropriate. Below this range the gel will be too soft and it will not keep its shape, while above this range it will be too rigid and prone to break. The optimal proportion of the N,N'-methylenebisacrylamide component is 1-3 % relative to the acrylamide component. The polyacrylamide gel possesses all advantageous properties necessary for a good electrophoresis medium, i.e. it is hydrophilic, free of electric charges and chemically stable. A further very important property of the polyacrylamide gel is that it does not participate in any non-specific or specific binding interaction with proteins. Furthermore, the polyacrylamide gel does not interfere with common protein staining reactions. 31  complexes and the pH of the gel buffer, as this will determine where individual proteins will migrate in the gel. 2.2.1.2 SDS-PAGE SDS-PAGE is an electrophoresis method to separate proteins. However, unlike in the case of native PAGE, here the proteins migrate in their denatured state. As it was mentioned in the general introduction to traditional (native) PAGE, the migration velocity of proteins is a function of their size, shape and the number of electric charges they carry. As the velocity is a complex function of these properties, native PAGE cannot be used to estimate the molecular mass of proteins. The traditional native PAGE method is similarly unable to assess whether a purified protein is composed of a single subunit or multiple subunits. Even a multi-subunit protein may migrate in a single sharp band. SDS-PAGE (Figure 4) was introduced to analyse such cases and to allow the estimation of the molecular mass of single-subunit proteins or those of individual subunits of multi-subunit proteins. SDS-PAGE is the most prevalent PAGE method currently in use. Figure 4. SDS polyacrylamide gel electrophoresis. SDS (sodium dodecyl sulphate) is an anionic detergent that unfolds proteins and provides them with extra negative charges. The amount of the associated SDS molecules—and therefore the number of charges—is proportional to the length of the polypeptide chain. The SDS gel separates individual polypeptide chains (monomeric proteins and subunits of multimeric proteins) according to their size.The velocity of the proteins is an inverse linear function of the logarithm of their molecular mass. Proteins of known molecular mass can be used to establish a calibration curve (a descending line) along which the unknown molecular mass of other proteins can be estimated. 36  3. Polymerase Chain Reaction (PCR) 3.1 Introduction Polymerase chain reaction (PCR) was invented by Mullis in 1983 and patented in 1985. Its principle is based on the use of DNA polymerase which is an in vitro replication of specific DNA sequences. This method can generate tens of billions of copies of a particular DNA fragment (the sequence of interest, DNA of interest, or target DNA) from a DNA extract (DNA template). If the sequence of interest is present in the DNA extract, it is possible to selectively replicate it (we speak of amplification) in very large numbers. The power of PCR is based on the fact that the amount of matrix DNA is not, in theory, a limiting factor. We can therefore amplify nucleotide sequences from infinitesimal amounts of DNA extract. PCR is therefore a technique of purification or cloning. DNA extracted from an organism or sample containing DNAs of various origins is not directly analyzable. It contains many mass of nucleotide sequences. It is therefore necessary to isolate and purify the sequence or sequences that are of interest, whether it is the sequence of a gene or noncoding sequences (introns, transposons, mini or microsatellites). From such a mass of sequences that constitutes the matrix DNA, the PCR can therefore select one or more sequences and amplify them by replication to tens of billions of copies. Once the reaction is complete, the amount of matrix DNA that is not in the area of interest will not have varied. In contrast, the amount of the amplified sequence(s) (the DNA of interest) will be very big. PCR makes it possible to amplify a signal from a background noise, so it is a molecular cloning method, and clone comes back to purity. There are many applications of PCR. It is a technique now essential in cellular and molecular biology. It permits, especially in a few hours, the “acellular cloning” of a DNA fragment through an automated system, which usually takes several days with standard techniques of molecular cloning. On the other hand, PCR is widely used for diagnostic purposes to detect the presence of a specific DNA sequence of this or that organism in a biological fluid. It is also used to make genetic fingerprints, whether it is the genetic identification of a person in the context of a judicial inquiry, or the identification of animal varieties, plant, or microbial for food quality testing, diagnostics, or varietal selection. PCR is still essential for performing sequencing or site-directed mutagenesis. 54 Finally, there are variants of PCR such as real-time PCR, competitive PCR, PCR in situ, RT-PCR, etc.At present, the revolutionary evolutions of the molecular biological research are based on the PCR technique which provides the suitable and specific products especially in the field of the characterization and the conservation of the genetic diversity. Several applications are possible in downstream of the PCR technique: (1) the establishment of a complete sequence of the genome of the most important livestock breeds; (2) development of a technology measuring scattered polymorphisms at loci throughout the genome (e.g., SNP detection methods); and (3) the development of a microarray technology to measure gene transcription on a large scale. The study of biological complexity is a new frontier that requires high throughput molecular technology, high speed and computer memory, new approaches to data analysis, and the integration of interdisciplinary skills. 3.2 Principle of the PCR PCR makes it possible to obtain, by in vitro replication, multiple copies of a DNA fragment from an extract. Matrix DNA can be genomic DNA as well as complementary DNA obtained by RT- PCR from a messenger RNA extract (poly-A RNA), or even mitochondrial DNA. It is a technique for obtaining large amounts of a specific DNA sequence from a DNA sample. This amplification is based on the replication of a double-stranded DNA template. It is broken down into three phases: a denaturation phase, a hybridization phase with primers, and an elongation phase. The products of each synthesis step serve as a template for the following steps, thus exponential amplification is achieved. The polymerase chain reaction is carried out in a reaction mixture which comprises the DNA extract (template DNA), Taq polymerase, the primers, and the four deoxyribonucleoside triphosphates (dNTPs) in excess in a buffer solution. The tubes containing the mixture reaction are subjected to repetitive temperature cycles several tens of times in the heating block of a thermal cycler (apparatus which has an enclosure where the sample tubes are deposited and in which the temperature can vary, very quickly and precisely, from 0 to 100°C by Peltier effect). The apparatus allows the programming of the duration and the succession of the cycles of temperature steps. Each cycle includes three periods of a few tens of seconds. The process of the PCR is subdivided into three stages as follows: 55 3.3 The denaturation It is the separation of the two strands of DNA, obtained by raising the temperature. The first period is carried out at a temperature of 94°C, called the denaturation temperature. At this temperature, the matrix DNA, which serves as matrix during the replication, is denatured: the hydrogen bonds cannot be maintained at a temperature higher than 80°C and the double-stranded DNA is denatured into single-stranded DNA (single-stranded DNA). 3.4 Hybridization The second step is hybridization. It is carried out at a temperature generally between 40 and 70°C, called primer hybridization temperature. Decreasing the temperature allows the hydrogen bonds to reform and thus the complementary strands to hybridize. The primers, short single-strand sequences complementary to regions that flank the DNA to be amplified, hybridize more easily than long strand matrix DNA. The higher the hybridization temperature, the more selective the hybridization, the more specific it is. 3.5 Elongation The third period is carried out at a temperature of 72°C, called elongation temperature. It is the synthesis of the complementary strand. At 72°C, Taq polymerase binds to primed single-stranded DNAs and catalyzes replication using the deoxyribonucleoside triphosphates present in the reaction mixture. The regions of the template DNA downstream of the primers are thus selectively synthesized. In the next cycle, the fragments synthesized in the previous cycle are in turn matrix and after a few cycles, the predominant species corresponds to the DNA sequence between the regions where the primers hybridize. It takes 20–40 cycles to synthesize an analyzable amount of DNA (about 0.1 μg). Each cycle theoretically doubles the amount of DNA present in the previous cycle. It is recommended to add a final cycle of elongation at 72°C, especially when the sequence of interest is large (greater than 1 kilobase), at a rate of 2 minutes per kilobase. PCR makes it possible to amplify sequences whose size is less than 6 kilobases. The PCR reaction is extremely rapid, it lasts only a few hours (2–3 hours for a PCR of 30 cycles). 3.6 Primers To achieve selective amplification of nucleotide sequences from a DNA extract by PCR, it is essential to have least one pair of oligonucleotides. These oligonucleotides, which will serve as primers for replication, are synthesized chemically and must be the best possible complementarity 56 with both ends of the sequence of interest that one wishes to amplify. One of the primers is designed to recognize complementarily a sequence located upstream of the fragment 5′–3′ strand DNA of interest; the other to recognize, always by complementarity, a sequence located upstream complementary strand (3′–5′) of the same fragment DNA. Primers are single-stranded DNAs whose hybridization on sequences flanking the sequence of interest will allow its replication so selective. The size of the primers is usually between 10 and 30 nucleotides in order to guarantee a sufficiently specific hybridization on the sequences of interest of the matrix DNA. 3.7 Taq polymerase DNA polymerase allows replication. We use a DNA polymerase purified or cloned from of an extremophilic bacterium, Thermus aquaticus, which lives in hot springs and resists temperatures above 100°C. This polymerase (Taq polymerase) has the characteristic remarkable to withstand temperatures of around 100°C, which are usually sufficient to denature most proteins. Thermus aquaticus finds its temperature of comfort at 72°C, optimum temperature for the activity of its polymerase. 3.8 The reaction conditions The volumes of reaction medium vary between 10 and 100 μl. There are a multitude of reaction medium formulas. However, it is possible to define a standard formula that is suitable for most polymerization reactions. This formula has been chosen by most manufacturers and suppliers, who, moreover, deliver a ready-to-use buffer solution with Taq polymerase. Concentrated 10 times, its formula is approximately the following: 100 mM Tris-HCl, pH 9.0; 15 mM MgCl2, 500 mM KCl. It is possible to add detergents (Tween 20, Triton X-100) or glycerol in order to increase the conditions of stringency that make it harder and therefore more selective hybridization of the primers. This approach is generally used to reduce the level of nonspecific amplifications due to the hybridization of the primers on sequences without relationship with the sequence of interest. We can also reduce the concentration of KCl until eliminated or increase the concentration of MgCl2. Indeed, some pairs of primers work better with solutions enriched with magnesium. On the other hand, with high concentrations of dNTP, the concentration of magnesium should be increased because of stoichiometric interactions between magnesium and dNTPs that reduce the amount of free magnesium in the reaction medium. dNTPs (deoxyribonucleoside triphosphates) provide both 57 the energy and the nucleotides needed for DNA synthesis during the chain polymerization. They are incorporated in the reaction medium in excess, that is, about 200 μM final. Depending on the reaction volume chosen, the primer concentration may vary between 10 and 50 pmol per sample. Matrix DNA can come from any organism and even complex biological materials that include DNAs from different organisms. But to ensure the success of a PCR, it is still necessary that the DNA matrix is not too degraded. This criterion is obviously all the more crucial as the size of the sequence of interest is large. It is also important that the DNA extract is not contaminated with inhibitors of the polymerase chain reaction (detergents, EDTA, phenol, proteins, etc.). The amount of template DNA in the reaction medium initiate that the amplification reaction can be reduced to a single copy. The maximum quantity may in no case exceed 2 μg. In general, the amounts used are in the range of 10–500 ng of template DNA. The amount of Taq polymerase per sample is generally between 1 and 3 units. The choice of the duration of the temperature cycles and the number of cycles depends on the size of the sequence of interest as well as the size and the complementarity of the primers. The durations should be reduced to a minimum not only to save time but also to prevent risk of nonspecific amplification. For denaturation and hybridization of primers, 30 seconds are usually sufficient. For elongation, it takes 1 minute per kilobase of DNA of interest and 2 minutes per kilobase for the final cycle of elongation. The number of cycles, generally between 20 and 40, is inversely proportional to the abundance of DNA matrix. 3.9 PCR product detection and analysis The product of a PCR consists of one or more DNA fragments (the sequence or sequences of interest). The detection and analysis of the products can be very quickly carried out by agarose gel electrophoresis (or acrylamide). The DNA is revealed by ethidium bromide staining. Thus, the products are instantly visible by ultraviolet transillumination (280–320 nm). Very small products are often visible very close to the migration front in the form of more or less diffuse bands. They correspond to primer dimers and sometimes to the primers themselves. Depending on the reaction conditions, nonspecific DNA fragments may be amplified to a greater or lesser extent, forming net bands or “smear”. On automated systems, a fragment analyzer is now used. This apparatus uses the principle of capillary electrophoresis. Fragment detection is performed by a laser diode. This is only possible if the PCR is performed with primers coupled to fluorochromes. 58  4. Real-time quantitative PCR (RT-PCR) 4.1 Introduction The introduction of real-time PCR technology has revolutionized the field of molecular diagnostics and has enabled the shift of molecular diagnostics toward a high-throughput, automated technology with lower turnaround times. It allows the sensitive, specific and reproducible quantification of mRNA. Real-time PCR assays are characterized by a wide dynamic range of quantification of 7–8 logarithmic decades, a high technical sensitivity (