Klenow fragment

The Klenow fragment is a large protein fragment produced when DNA polymerase I from E. coli is enzymatically cleaved by the protease subtilisin. First reported in 1970, it retains the 5’ → 3’ polymerase activity and the 3’ → 5’ exonuclease activity for removal of precoding nucleotides and proofreading, but loses its 5' → 3' exonuclease activity.

The other smaller fragment formed when DNA polymerase I from E. coli is cleaved by subtilisin retains the 5'→ 3' exonuclease activity but does not have the other two activities exhibited by the Klenow fragment (i.e. 5'-> 3' polymerase activity, and 3'->5' nuclease activity).

Because the 5' → 3' exonuclease activity of DNA polymerase I from E. coli makes it unsuitable for many applications, the Klenow fragment, which lacks this activity, can be very useful in research. The Klenow fragment is extremely useful for research-based tasks such as:

Synthesis of double-stranded DNA from single-stranded templates

Filling in recessed 3' ends of DNA fragments

Digesting away protruding 3' overhangs

Preparation of radioactive DNA probes

The Klenow fragment was also the original enzyme used for greatly amplifying segments of DNA in the polymerase chain reaction (PCR) process, before being replaced by thermostable enzymes such as Taq polymerase

Just as the 5' → 3' exonuclease activity of DNA polymerase I from E.coli can be undesirable, the 3' → 5' exonuclease activity of Klenow fragment can also be undesirable for certain applications. This problem can be overcome by introducing mutations in the gene that encodes Klenow. This results in forms of the enzyme being expressed that retain 5' → 3' polymerase activity, but lack any exonuclease activity (5' → 3' or 3' → 5'). This form of the enzyme is called the exo- Klenow fragment.

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Taq polymerase

Taq polymerase is a thermostable DNA polymerase named after the thermophilic bacterium Thermus aquaticus from which it was originally isolated. It is often abbreviated to "Taq Pol" (or simply "Taq"), and is frequently used in polymerase chain reaction (PCR), methods for greatly amplifying short segments of DNA.
T. aquaticus is a bacterium that lives in hot springs and hydrothermal vents, and Taq polymerase was identified as an enzyme able to withstand the protein-denaturing conditions (high temperature) required during PCR. Therefore it replaced the DNA polymerase from E.coli originally used in PCR . Taq's temperature optimum for activity is 75-80°C, with a halflife of 9 minutes at 97.5°C, and can replicate a 1000 base pair strand of DNA in less than 10 seconds at 72°C.
One of Taq's drawbacks is its relatively low replication fidelity. It lacks a 3' to 5' exonuclease proofreading activity, and has an error rate measured at about 1 in 9,000 nucleotides. Some thermostable DNA polymerases have been isolated from other thermophilic bacteria and archaea, such as Pfu DNA polymerase, possessing a proofreading activity, and are being used instead of (or in combination with) Taq for high-fidelity amplification.Taq makes DNA products that have A (Adenine) overhangs at their 3' ends. This may be useful in TA Cloning, whereby a cloning vector (such as a plasmid) is used which has a T (Thymine) 3' overhang, which complements with the A overhang of the PCR product, thus enabling ligation of the PCR product into the plasmid vector.

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Taq polymerase in PCR

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In the early 1980s Kary Mullis was working at Cetus Corporation on the application of synthetic DNAs to biotechnology. He was familiar with the use of DNA oligonucleotides as probes for binding to target DNA strands, as well as their use as primers for DNA sequencing and cDNA synthesis. In 1983, he began using two primers, one to hybridize to each strand of a target DNA, and adding DNA polymerase to the reaction. This led to exponential DNA replication, greatly amplifying the amounts of DNA between the primers.
However, after each round of replication the mixture needs to be heated above 90ºC to denature the newly formed DNA, allowing the strands to separate and act as templates in the next round of amplification. Unfortunately, this heating step also inactivates the DNA polymerase then being used, the Klenow fragment of the DNA Polymerase I from E. coli.
Use of the thermostable Taq polymerase eliminates the need for having to add new enzyme to the PCR reaction during the thermocycling process. A single closed tube in a relatively simple machine can be used to carry out the entire process. Thus, the use of Taq polymerase was the key idea that made PCR applicable to a large variety of molecular biology problems concerning DNA analysis.

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Significance

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Hoffmann-La Roche eventually bought the PCR and Taq patents from Cetus for $330 million, from which it may have received up to $2 billion in royalties. In 1989 Science Magazine named Taq polymerase its first "Molecule of the Year". Kary Mullis received the Nobel Prize in 1993, the only one awarded for research performed at a biotechnology company. By the early 1990s the PCR technique with Taq polymerase was being used in many areas, including basic molecular biology research, clinical testing, and forensics. It also began to find a pressing application in direct detection of the HIV virus in AIDS.

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Variants of PCR

Often only a small modification needs to be made to the 'standard' PCR protocol to achieve a desired goal:

One of the first adjustments made to PCR was the amplification of more than one target in a single tube. Multiplex-PCR can involve up to a dozen pairs of primers acting independently. This modification might be used simply to avoid having to prepare many individual reactions, or could allow the simultaneous analysis of multiple targets in a sample that has only a single copy of a genome. In testing for genetic disease mutations, six or more amplifications might be combined. In the standard protocol for DNA Fingerprinting, the 13 targets assayed are often amplified in groups of 3 or 4. Multiplex Ligation-dependent Probe Amplification (or MLPA) permits multiple targets to be amplified using only a single pair or primers, avoiding the resolution limitations of multiplex PCR

VNTR PCR involves few modifications to the basic PCR process, but instead targets areas of the genome that exhibit length variation. The analysis of the genotypes of the sample usually involves simple sizing of the amplification products by gel electrophoresis. Analysis of smaller VNTR segments known as Short Tandem Repeats (or STRs) is the basis for DNA Fingerprinting databases such as CODIS.

Asymmetric PCR is used to preferentially amplify one strand of the target DNA. It finds use in some types of sequencing and hybridization probing, where having only one of the two complementary strands of the product is advantageous. PCR is carried out as usual, but with a limiting amount of one of the primers. When it becomes depleted, continued replication leads to an arithmetic increase in extension of the other primer. A recent modification on this process, known as Linear-After-The-Exponential-PCR (or LATE-PCR), uses a limiting primer with a higher melting temperature Melting temperature (or Tm) than the excess primer to maintain reaction efficiency as the limiting primer concentration decreases mid-reaction. (Also see Overlap-extension PCR).

Some modifications are needed to perform long PCR. The original Klenow-based PCR process had trouble making a product larger than about 400 bp. However, early characterization of Taq polymerase showed that it could amplify targets up to several thousand bp long. Since then, modified protocols have allowed targets of over 50,000 bp to be amplified

Nested PCR, another early modification, can be used to increase the specificity of DNA amplification. Two sets of primers are used in two successive reactions. In the first, one pair of primers is used to generate DNA products, which may also contain products amplified from non-target areas. The products from the first PCR are then used to start a second, using one ('hemi-nesting') or two different primers whose binding sites are located (nested) within the first set. The specificity of all of the primers is combined, usually leading to a single product. Nested PCR is often more successful in specifically amplifying long DNA products than conventional PCR, but it requires more detailed knowledge of the sequence of the target.

Quantitative PCR (or Q-PCR) is used to measure the specific amount of target DNA (or RNA) in a sample. The normal PCR process is performed in a way that is largely qualitative - the amount of final product is only slightly proportional to the initial amount of target. By carefully running the amplification only within the phase of true exponential increase (avoiding the later 'plateau' phase), the amount of product is more proportional to the initial amount of target. Thermal cyclers have been developed which can monitor the amount of product during the amplification, allowing quantitation of samples containing a wide range of target copies. A method currently used is Quantitative Real-Time PCR. QRT-PCR methods use fluorescent dyes, such as Sybr Green, or fluorophore-containing DNA probes, such as TaqMan, to measure the amount of amplified product as the amplification progresses. It is often confusingly referred to as RT-PCR, the same acronym used for PCR combined with Reverse Transcriptase (see below), which itself might be used in conjunction with Q-PCR. More appropriate acronyms are QRT-PCR or RTQ-PCR.

Hot-start PCR is a technique that modifies the way that a PCR mixture is initially heated. During this step the polymerase is active, but the target has not yet been denatured and the primers may be able to bind to non-specific locations (or even to each other). The technique can be performed manually by heating the reaction components to the melting temperature (e.g. 95°C) before adding the polymerase. Alternatively, specialized systems have been developed that inhibit the polymerase's activity at ambient temperature, either by the binding of an antibody, or by the presence of covalently bound inhibitors that only dissociate after a high-temperature activation step. 'Hot-start/cold-finish PCR' is achieved with new hybrid polymerases that are inactive at ambient temperature and are only activated at elevated temperatures.

Another simple modification can also decrease non-specific amplification. In Touchdown PCR, the temperature used to anneal the primers is gradually decreased in later cycles. The annealing temperature in the early cycles is usually 3-5°C above the standard Tm of the primers used, while in the later cycles it is a similar amount below the Tm. The initial higher annealing temperature leads to greater specificity for primer binding, while the lower temperatures permit more efficient amplification to the end of the reaction.

Other common modifications to PCR allow it to amplify low copy targets. The original report on Taq polymerase showed how the use of up to 60 cycles could amplify targets diluted to just one copy per reaction tube. A later report showed how multiple genetic loci could be amplified and analyzed from a single sperm. Modified protocols have allowed the identification of just one copy of the HIV genome within the DNA of up to 70,000 host cells.

Assembly PCR (also known as Polymerase Cycling Assembly or PCA) is the artificial synthesis of long DNA structures by performing PCR on a pool of long oligonucleotides with short overlapping segments. The oligonucleotide building blocks alternate between sense and antisense directions, and the overlaps determine the order of oligonucleotides, thereby selectively producing the final long DNA product.

In Colony PCR, bacterial colonies are rapidly screened by PCR for correct DNA vector constructs. Colonies are sampled with a sterile toothpick and dabbed into a master mix. To free the DNA for amplification, PCR is either started with an extended time at 95°C (when standard polymerase is used), or with a shortened denaturation step at 100°C and special chimeric DNA polymerase. Colonies from the master mix that shows the desired product are then tested individually.

The Digital polymerase chain reaction simultaneously amplifies thousands of samples, each in a separate droplet within an emulsion.

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What Is a Primer ?

A primer is a strand of nucleic acid that serves as a starting point for DNA replication. They are required because the enzymes that catalyze replication, DNA polymerases, can only add new nucleotides to an existing strand of DNA. The polymerase starts replication at the 3'-end of the primer, and copies the opposite strand.

In most cases of natural DNA replication, the primer for DNA synthesis and replication is a short strand of RNA (which can be made de novo). This RNA is produced by primase, and is later removed and replaced with DNA by a repair polymerase.

Many of the laboratory techniques of biochemistry and molecular biology that involve DNA polymerase, such as DNA sequencing and the polymerase chain reaction (PCR), require primers. These primers are usually short, chemically synthesized oligonucleotides, with a length of about twenty bases. They are hybridized to a target DNA, which is then copied by the polymerase.

Uses of synthetic primers

DNA sequencing is used to determine the nucleotides in a DNA strand; the chain termination method (dideoxy sequencing or Sanger method) uses a primer as a start marker for the chain reaction.

In PCR, primers are used to determine the DNA fragment to be amplified by the PCR process. The length of primers is usually not more than 30 nucleotides, and they match exactly the beginning and the end of the DNA fragment to be amplified. They direct replication towards each other - the extension of one primer by polymerase then becomes the template for the other, leading to an exponential increase in the target segment.
It is worth noting that primers are not essentially always necessary for DNA synthesis and can in fact be used by viral polymerases, e.g. influenza, for RNA synthesis.

PCR primer design

The melting temperature of a primer is defined as the temperature at which 50% of that same DNA molecule species form a stable double helix and the other 50% have been separated to single strand molecules. The melting temperature required increases with the length of the primer. Primers that are too short would anneal at several positions on a long DNA template, which would result in non-specific copies. On the other hand, the length of a primer is limited by the temperature required to melt it. Melting temperatures that are too high, i.e., above 80°C, can also cause problems since the DNA polymerases used for PCR are less active at such temperatures. The optimum length of a primer is generally from 20 to 30 nucleotides with a melting temperature between about 55°C and 65°C.

Pairs of primers should have the similar melting temperatures as annealing in a PCR reaction occurs for both simultaneously. A primer with a Tm significantly higher than the reaction's annealing temperature may mishybridize and extend at an incorrect location along the DNA sequence, while Tm significantly lower than the annealing temperature may fail to anneal and extend at all.

Primer sequences need to be chosen to uniquely select for a region of DNA, avoiding the possibility of mishybridization to a similar sequence nearby. A commonly used method is BLAST search whereby all the possible regions to which a primer may bind can be seen. Both the nucleotide sequence as well as the primer itself can be BLAST searched. Alternatively use of software such as Beacon designer, may yeild to specific primers. Mononucleotide repeats should be avoided, as loop formation can occur and contribute to mishybridization. Primers should not easily anneal with other primers in the mixture (either other copies of same or the reverse direction primer); this phenomenon can lead to the production of 'primer dimer' products contaminating the mixture. Primers should also not anneal strongly to themselves, as internal hairpins and loops could hinder the annealing with the template DNA.

Degenerate primers

Sometimes degenerate primers are used. These are actually mixtures of similar, but not identical, primers. They may be convenient if the same gene is to be amplified from different organisms, as the genes themselves are probably similar but not identical. The other use for degenerate primers is when primer design is based on protein sequence. As several different codons can code for one amino acid, it is often difficult to deduce which codon is used in a particular case. Therefore primer sequence corresponding to the amino acid isoleucine might be "ATH", where A stands for adenine, T for thymine, and H for adenine, thymine, or cytosine, according to the genetic code for each codon, using the IUPAC symbols for degenerate bases. Use of degenerate primers can greatly reduce the specificity of the PCR amplification. The problem can be partly solved by using touchdown PCR.

Degenerate primers are widely used and extremely useful in the field of microbial ecology. They allow for the amplification of genes from thus far uncultivated microorganisms or allow the recovery of genes from organisms where genomic information is not available. Usually, degenerate primers are designed by aligning gene sequencing found in GenBank. Differences among sequences are accounted for by using IUPAC degeneracies for individual bases. PCR primers are then synthesized as a mixture of primers corresponding to all permutations.

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Real-time PCR

In molecular biology, real-time polymerase chain reaction, also called quantitative real time polymerase chain reaction (qPCR) or kinetic polymerase chain reaction, is a laboratory technique based on the polymerase chain reaction, which is used to amplify and simultaneously quantify a targeted DNA molecule. It enables both detection and quantification (as absolute number of copies or relative amount when normalized to DNA input or additional normalizing genes) of a specific sequence in a DNA sample.
The procedure follows the general principle of polymerase chain reaction; its key feature is that the amplified DNA is quantified as it accumulates in the reaction in real time after each amplification cycle. Two common methods of quantification are the use of fluorescent dyes that intercalate with double-stranded DNA, and modified DNA oligonucleotide probes that fluoresce when hybridized with a complementary DNA.
Frequently, real-time polymerase chain reaction is combined with reverse transcription polymerase chain reaction to quantify low abundance messenger RNA (mRNA), enabling a researcher to quantify relative gene expression at a particular time, or in a particular cell or tissue type.
Although real-time quantitative polymerase chain reaction is often marketed as RT-PCR, it should not be confused with reverse transcription polymerase chain reaction, also known as RT-PCR.
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Background

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Cells in all organisms regulate their cellular activities by activating or deactivating the expression of their genes. Gene expression is usually directly proportional to the number of copies of messenger RNA (mRNA) of a particular gene in a cell or tissue.
Traditionally, the expression level of a gene has been estimated by visualizing the abundance of its mRNA transcript in a sample with a technique called northern blotting. In this method, purified RNA is separated by agarose gel electrophoresis, transferred to a solid matrix (such as a nylon membrane), and probed with a specific DNA probe that is complementary to the gene of interest. Although this technique is still used to assess gene expression, it requires relatively large amounts of RNA and provides only qualitative or semiquantitative information of mRNA levels.
In order to robustly detect and quantify gene expression from small amounts of RNA, amplification of the gene transcript is necessary. The polymerase chain reaction is a common method for amplifying DNA; for mRNA-based PCR the RNA sample is first reverse transcribed to cDNA with reverse transcriptase.
Development of PCR technologies based on reverse transcription and fluorophores permits measurement of DNA amplification during PCR in real time, i.e., the amplified product is measured at each PCR cycle. The data thus generated can be analysed by computer software to calculate relative gene expression in several samples, or mRNA copy number. Real-time PCR can also be applied to the detection and quantification of DNA in samples to determine the presence and abundance of a particular DNA sequence in these samples.

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Real-time PCR using double-stranded DNA dyes

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A DNA-binding dye binds to all double-stranded (ds)DNA in a PCR reaction, causing fluorescence of the dye. An increase in DNA product during PCR therefore leads to an increase in fluorescence intensity and is measured at each cycle, thus allowing DNA concentrations to be quantified. However, dsDNA dyes such as SYBR Green will bind to all dsDNA PCR products, including nonspecific PCR products (such as "primer dimers"). This can potentially interfere with or prevent accurate quantification of the intended target sequence. The reaction is prepared as usual, with the addition of fluorescent dsDNA dye. The reaction is run in a thermocycler, and after each cycle, the levels of fluorescence are measured with a detector; the dye only fluoresces when bound to the dsDNA (i.e., the PCR product). With reference to a standard dilution, the dsDNA concentration in the PCR can be determined.
Like other real-time PCR methods, the values obtained do not have absolute units associated with it (i.e. mRNA copies/cell). As described above, a comparison of a measured DNA/RNA sample to a standard dilution will only give a fraction or ratio of the sample relative to the standard, allowing only relative comparisons between different tissues or experimental conditions. To ensure accuracy in the quantification, it is usually necessary to normalize expression of a target gene to a stably expressed gene (see below). This can correct possible differences in RNA quantity or quality across experimental samples.

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Fluorescent reporter probe method

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Using fluorescent reporter probes is the most accurate and most reliable of the methods, but also the most expensive. It uses a sequence-specific RNA or DNA-based probe to quantify only the DNA containing the probe sequence; therefore, use of the reporter probe significantly increases specificity, and allows quantification even in the presence of some non-specific DNA amplification. This potentially allows for multiplexing - assaying for several genes in the same reaction by using specific probes with different-coloured labels, provided that all genes are amplified with similar efficiency.
It is commonly carried out with an RNA-based probe with a fluorescent reporter at one end and a quencher of fluorescence at the opposite end of the probe. The close proximity of the reporter to the quencher prevents detection of its fluorescence; breakdown of the probe by the 5' to 3' exonuclease activity of the taq polymerase breaks the reporter-quencher proximity and thus allows unquenched emission of fluorescence, which can be detected. An increase in the product targeted by the reporter probe at each PCR cycle therefore causes a proportional increase in fluorescence due to the breakdown of the probe and release of the reporter.

1. The PCR reaction is prepared as usual and the reporter probe is added.
2. As the reaction commences, during the annealing stage of the PCR both probe and primers anneal to the DNA target.
3. Polymerisation of a new DNA strand is initiated from the primers, and once the polymerase reaches the probe, its 5'-3-exonuclease degrades the probe, physically separating the fluorescent reporter from the quencher, resulting in an increase in fluorescence.
4. Fluorescence is detected and measured in the real-time PCR thermocycler, and its geometric increase corresponding to exponential increase of the product is used to determine the threshold cycle (CT) in each reaction.

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Quantitation

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Quantitating gene expression by traditional methods presents several problems. Firstly, detection of mRNA on a Northern blot or PCR products on a gel or Southern blot is time-consuming and does not allow precise quantitation. Also, over the 20-40 cycles of a typical PCR reaction, the amount of product reaches a plateau determined more by the amount of primers in the reaction mix than by the input template/sample.
Relative concentrations of DNA present during the exponential phase of the reaction are determined by plotting fluorescence against cycle number on a logarithmic scale (so an exponentially increasing quantity will give a straight line). A threshold for detection of fluorescence above background is determined. The cycle at which the fluorescence from a sample crosses the threshold is called the cycle threshold, Ct. Since the quantity of DNA doubles every cycle during the exponential phase, relative amounts of DNA can be calculated, e.g. a sample whose Ct is 3 cycles earlier than another's has 23 = 8 times more template.
Amounts of RNA or DNA are then determined by comparing the results to a standard curve produced by RT-PCR of serial dilutions (e.g. undiluted, 1:4, 1:16, 1:64) of a known amount of RNA or DNA. As mentioned above, to accurately quantify gene expression, the measured amount of RNA from the gene of interest is divided by the amount of RNA from a housekeeping gene measured in the same sample to normalize for possible variation in the amount and quality of RNA between different samples. This normalization permits accurate comparison of expression of the gene of interest between different samples, provided that the expression of the reference (housekeeping) gene used in the normalization is very similar across all the samples. Choosing a reference gene fulfilling this criterion is therefore of high importance, and often challenging, because only very few genes show equal levels of expression across a range of different conditions or tissues.
Applications of real-time polymerase chain reaction

There are numerous applications for real-time polymerase chain reaction in the laboratory. It is commonly used for both diagnostic and research applications.
Diagnostically real-time PCR is applied to rapidly detect the presence of genes involved in infectious diseases, cancer and genetic abnormalities. In the research setting, real-time PCR is mainly used to provide highly sensitive quantitative measurements of gene transcription.
The technology may be used in determining how the genetic expression of a particular gene changes over time, such as in the response of tissue and cell cultures to an administration of a pharmacological agent, progression of cell differentiation, or in response to changes in environmental conditions.
Also, the technique is used in Environmental microbiology, for example to quantify resistance genes in water samples.

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What is a Genetic Marker

A genetic marker is a known DNA sequence. It can be described as a variation, which may arise due to mutation or alteration in the genomic loci, that can be observed. A genetic marker may be a short DNA sequence, such as a sequence surrounding a single base-pair change (single nucleotide polymorphism, SNP), or a long one, like minisatellites.

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Some commonly used types of genetic markers are

• RFLP (or Restriction fragment length polymorphism)
• AFLP (or Amplified fragment length polymorphism)
• RAPD (or Random amplification of polymorphic DNA)
• VNTR (or Variable number tandem repeat)
• Microsatellite polymorphism
• SNP (or Single nucleotide polymorphism)
• STR (or Short tandem repeat)
• SFP (or Single feature polymorphism)

They can be further categorized as dominant or co-dominant. Dominant markers allow for analyzing many loci at one time, e.g. RAPD. A primer amplifying a dominant marker could amplify at many loci in one sample of DNA with one PCR reaction. Co-dominant markers analyze one locus at a time. A primer amplifying a co-dominant marker would yield one targeted product.

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Uses:

Genetic markers can be used to study the relationship between an inherited disease and its genetic cause (for example, a particular mutation of a gene that results in a defective protein). It is known that pieces of DNA that lie near each other on a chromosome tend to be inherited together. This property enables the use of a marker, which can then be used to determine the precise inheritance pattern of the gene that has not yet been exactly localized.
Genetic markers have to be easily identifiable, associated with a specific locus, and highly polymorphic, because homozygotes do not provide any information. Detection of the marker can be direct by RNA sequencing, or indirect using allozymes.

Some of the methods used to study the genome or phylogenetics are RFLP, Amplified fragment length polymorphism (AFLP), RAPD, SSR.

Insulin production

Genetic markers also play a role in genetic engineering, as they can be used to produce normal, functioning proteins to replace defective ones. The damaged or faulty section of DNA is removed and replaced with the identical, but functioning, gene sequence from another source.
This is done by removal of the faulty section of DNA and its replacement with the functioning gene from another source, usually a human donor. These gene sections are placed in solution with bacterial cells, a small number of which take up the genetic material and reproduce the new DNA sequence. Engineers need to know which bacteria have been successful in duplicating these genes so another gene is added, altering the bacteria's resistance to antibiotics. Replica plating or a fermenter is used to grow enough bacteria to test resistance to antibiotics. It is important that the cultures are not mixed.This process can be used as a treatment for diabetes mellitus. Bacterial DNA often has two resistency genes: one for tetracycline and one for ampicillin. The insulin gene can be inserted in the middle of the ampicillin gene after it has been removed using restriction endonucleases. If the gene has been taken up, the bacteria both produces insulin and is also no longer ampicillin resistant. The bacteria are then allowed to grow on an agar plate containing a culture medium. The bacteria grow and produce colonies on the agar jelly. A piece of filter paper can be placed onto the top of this agar plate so that the exact positions of the colonies are remembered. This produces a copy which can then be transferred onto a second agar plate containing ampicillin. All of the bacteria that are not resistant to ampicillin will die. These locations on the second plate show the places on the first plate where bacteria are not resistant and therefore produce insulin. Another similar method is followed, in which an epitope sequence is added to insert. When the insert is expressed so is the epitope. Then this epitope can be effectively bound using an antibody on a filter paper. And the expressing colonies can be easily selected.

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What is a Gene

A gene is a locatable region of genomic sequence, corresponding to a unit of inheritance, which is associated with regulatory regions, transcribed regions and/or other functional sequence regions. The physical development and phenotype of organisms can be thought of as a product of genes interacting with each other and with the environment. A concise definition of a gene, taking into account complex patterns of regulation and transcription, genic conservation and non-coding RNA genes, has been proposed by Gerstein et al."A gene is a union of genomic sequences encoding a coherent set of potentially overlapping functional products".
Colloquially, the term gene is often used to refer to an inheritable trait which is usually accompanied by a phenotype as in ("tall genes" or "bad genes") -- the proper scientific term for this is allele.
In cells, genes consist of a long strand of DNA that contains a promoter, which controls the activity of a gene, and coding and non-coding sequence. Coding sequence determines what the gene produces, while non-coding sequence can regulate the conditions of gene expression. When a gene is active, the coding and non-coding sequence is copied in a process called transcription, producing an RNA copy of the gene's information. This RNA can then direct the synthesis of proteins via the genetic code. But some RNAs are used directly, for example as part of the ribosome. These molecules resulting from gene expression, whether RNA or protein, are known as gene products.
Genes often contain regions that do not encode products, but regulate gene expression। The genes of eukaryotic organisms can contain regions called introns that are removed from the messenger RNA in a process called splicing. The regions encoding gene products are called exons. In eukaryotes, a single gene can encode multiple proteins, which are produced through the creation of different arrangements of exons through alternative splicing. In prokaryotes (bacteria and archaea), introns are less common and genes often contain a single uninterrupted stretch of DNA, called a cistron, that codes for a product. Prokaryotic genes are often arranged in groups called operons with promoter and operator sequences that regulate transcription of a single long RNA. This RNA contains multiple coding sequences. Each coding sequence is preceded by a Shine-Dalgarno sequence that ribosomes recognize.

The total set of genes in an organism is known as its genome. An organism's genome size is generally lower in prokaryotes, both in number of base pairs and number of genes, than even single-celled eukaryotes. However, there is no clear relationship between genome sizes and complexity in eukaryotic organisms. One of the largest known genomes belongs to the single-celled amoeba Amoeba dubia, with over 670 billion base pairs, some 200 times larger than the human genome. The estimated number of genes in the human genome has been repeatedly revised downward since the completion of the Human Genome Project; current estimates place the human genome at just under 3 billion base pairs and about 20,000–25,000 genes. A recent Science article gives a number of 20,488 protein-coding genes, with perhaps 100 more yet to be discovered. The gene density of a genome is a measure of the number of genes per million base pairs (called a megabase, Mb); prokaryotic genomes have much higher gene densities than eukaryotes. The gene density of the human genome is roughly 12–15 genes per megabase pair.

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What is Stellite DNA

Satellite DNA consists of highly repetitive DNA, and is so called because repetitions of a short DNA sequence tend to produce a different frequency of the nucleotides adenine, cytosine, guanine and thymine, and thus have a different density from bulk DNA - such that they form a second or 'satellite' band when genomic DNA is separated on a density gradient.
Length


A repeated pattern can be between 1 base pair long (a mononucleotide repeat) to several thousand base pairs long, and the total size of a satellite DNA block can be several megabases without interruption. Most satellite DNA is localized to the telomeric or the centromeric region of the chromosome. The nucleotide sequence of the repeats is fairly well conserved across a species. However, variation in the length of the repeat is common. For example, minisatellite DNA is a short region (1-5kb) of 20-50 repeats. The difference in length of the minisatellites is the basis for DNA fingerprinting.

Satellite DNA, at least the microsatellite variety, is thought to have originated by slippage of a replicated chromosome against its template.

Microsatellites are often found in transcription units. Often the base pair repetition will disrupt proper protein synthesis, leading to diseases such as myotonic dystrophy.

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What is Selfish DNA

Selfish DNA refers to those sequences of DNA which, in their purest form, have two distinct properties:

(1) the DNA sequence spreads by forming additional copies of itself within the genome; and

(2) it makes no specific contribution to the reproductive success of its host organism. This idea was sketched briefly by Richard Dawkins in his 1976 book The Selfish Gene and was explicitly exposed in two 1980 articles in Nature magazine. According to one of these articles:

So, the selfish DNA can be considered an efficient replicator that follows another way of increasing in number.

Examples

• Transposons copy themselves to different loci inside the genome. These elements constitute a large fraction of eukaryotic genome sizes (C-values): about 45% of the human genome is composed of transposons and their defunct remnants.
• Homing endonuclease genes cleave DNA at its own site on the homologous chromosome, triggering the DNA double-stranded break repair system, which "repairs" the break by copying the HEG onto the homologous chromosome. HEGs have been characterized in yeast, and can only survive by passing between multiple isolated populations or species. Supernumerary B chromosomes are nonessential chromosomes that are transmitted in higher-than-expected frequencies, which leads to their accumulation in progenies.

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What is Junk DNA

In molecular biology, "junk" DNA is a provisional label for the portions of the DNA sequence of a chromosome or a genome for which no function has yet been identified. Scientists fully expect to find functions for some, but definitely not all, of this provisionally classified collection. About 80-90% of the human genome has been designated as "junk", including most sequences within introns and most intergenic DNA. While much of this sequence may be an evolutionary artifact that serves no present-day purpose, some is believed to function in ways that are not currently understood. Moreover, the conservation of some junk DNA over many millions of years of evolution may imply an essential function. Some consider the "junk" label as something of a misnomer, but others consider it apposite as junk is stored away for possible new uses, rather than thrown out; others prefer the term "noncoding DNA" (although junk DNA often includes transposons that encode proteins with no clear value to their host genome). However it now appears that, although protein-coding DNA makes up barely 2% of the human genome, about 80% of the bases in the genome may be transcribed, but transcription by itself does not necessarily imply function.

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Broadly, the science of functional genomics has developed widely accepted techniques to characterize protein-coding genes, RNA genes, and regulatory regions. In the genomes of most plants and animals, however, these together constitute only a small percentage of genomic DNA (less than 2% in the case of humans). The function, if any, of the remainder remains under investigation. Most of it can be identified as repetitive elements that have no known biological function for their host (although they are useful to geneticists for analyzing lineage and phylogeny). Still, a large amount of sequence in these genomes falls under no existing classification other than "junk".
Overall genome size, and by extension the amount of junk DNA, appears to have little relationship to organism complexity: the genome of the unicellular Amoeba dubia has been reported to contain more than 200 times the amount of DNA in humans".
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Hypotheses of origin and function
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There are some hypotheses, none conclusively established, from the most academic to the less expected, for how junk DNA arose and why it persists in the genome:

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  These chromosomal regions could be composed of the now-defunct remains of ancient genes, known as pseudogenes, which were once functional copies of genes but have since lost their protein-coding ability (and, presumably, their biological function). After non-functionalization, pseudogenes are free to acquire genetic noise in the form of random mutations.
• 8% of human junk DNA has been shown to be formed by retrotransposons of Human Endogenous Retroviruses (HERVs), although as much as 25% is recognisably formed of retrotransposons. This is a lower limit on how much of the genome is retrotransposons because older remains might not be recognizable having accumulated too much mutation. New research suggests that genome size variation in at least two kinds of plants is mostly because of retrotransposons.
• In 1997, Steven Sparks proposed that "The end purpose of this "excess DNA" must be to reduce the probability of transcribable genes being cut by chromosomal crossover. Gametes can survive only when their important, transcribed genes are saved from meiotic cutting by being surrounded with "buffer DNA".
• Junk DNA might provide a reservoir of sequences from which potentially advantageous new genes can emerge. In this way, it may be an important genetic basis for evolution.
• Some junk DNA could simply be spacer material that allows enzyme complexes to form around functional elements more easily. In this way, the junk DNA could serve an important function even though the actual sequence information it contains is irrelevant.
• Some portions of junk DNA could serve presently unknown regulatory functions, controlling the expression of certain genes, the development of an organism from embryo to adult, and/or development of certain organs/organelles.
• More and more scientists believe that in fact regulatory layer(s) in the "junk DNA", such as through non-coding RNAs, altogether contain genetic programming at least on par with, and possibly much more important than protein coding genes. But still how much of the 98% would be involved in such activity is unknown.

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Hok/sok system

The host killing/suppressor of killing system, it is also known as hok/sok system, in molecular biology, is a postsegregational killing system of the plasmid R1 of Escherichia coli(or is it).

In simple words, the system is controlled by two genes, hok and sok, coding respectively what can be thought of as a long-lived poison and a short-lived antidote. After cell division, daughter cells without a copy of the plasmid die, as the poison is still active from the parent cell, while the short-lived antidote is not stopping the poison anymore. Only cells with a plasmid can produce more antidote and survive. For this reason, the killing system is "postsegregational", since cell death occurs after segregation of the plasmid.

The hok gene codes for a 52 amino acid toxic protein which causes cell death by depolarization of the cell membrane. The translation of hok mRNA is, however, inhibited by the transcript of the sok gene, which is an antisense regulator and binds to the hok mRNA, forming a duplex which is recognized by the RNase III and degraded. The killing mechanism is obtained through differential decay rates of the hok and sok transcripts: while hok mRNA is quite stable, sok-RNA is rapidly degraded, which would allow hok to be expressed; however the higher rate of transcription of sok compensate, leaving hok mRNA untranslated in plasmid-containing cells. The loss of plasmid causes the hok mRNA not to be inhibited anymore by sok antisense, leading to protein expression and cell death.

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El Tor

El Tor is the name given to a particular strain of the bacterium Vibrio cholerae, the causative agent of cholera. Also known as O1, it has been the dominant strain in the seventh global pandemic. It is distinguished from the classic strain at a genetic level, although both are in the O1 serogroup and both contain Inaba, Ogawa and Hikojima serotypes. It was first identified in 1905 at a camp in El-Tor, Egypt.

El Tor was identified again in an outbreak in 1937 but the pandemic did not arise until 1961 in Sulawesi. El Tor spread through Asia (Bangladesh in 1963, India in 1964) and then into the Middle East, Africa and Europe. From North Africa it spread into Italy by 1973. In the late 1970s there were small outbreaks in Japan and in the South Pacific.

The extent of the pandemic has been due to the relative mildness (lower expression level) of El Tor, the disease has many more asymptomatic carriers than is usual, outnumbering active cases by up to 50:1. El Tor also remains in the body for longer and survives better than other known types. The actual infection is also relatively mild, or at least rarely fatal. Additionally El Tor is capable of host-to-host transmission, unlike the classic strain

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Ice-minus bacteria

Ice-minus bacteria is a nickname given to a variant of the common bacterium Pseudomonas syringae (P. syringae). This strain of P. syringae lacks the ability to produce a certain surface protein, usually found on wild-type "ice-plus" P. syringae. The "ice-plus" protein (Ina protein, "Ice nucleation-active" protein) found on the outer bacterial cell wall acts as the nucleating centers for ice crystals. This facilitates ice formation, hence the designation "ice-plus." The ice-minus variant of P. syringae is a mutant, lacking the gene responsible for ice-nucleating surface protein production. This lack of surface protein provides a less favorable environment for ice formation. Both strains of P. syringae occur naturally, but recombinant DNA technology has allowed for the synthetic removal or alteration of specific genes, enabling the creation of the ice-minus strain.

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Economic importance:

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The success of the agricultural world is heavily dependent on the weather. Cold weather conditions are directly responsible for the appearance of frost on plants and most importantly, crops. In the United States alone, it has been estimated that frost accounts for approximately $1 billion in crop damage each year. As P. syringae commonly inhabits plant surfaces, its ice nucleating nature incites frost development, freezing the buds of the plant and destroying the occurring crop. The introduction of an ice-minus strain of P. syringae to the surface of plants would incur competition between the strains. Should the ice-minus strain win out, the ice nucleate provided by P. syringae would no longer be present, lowering the level of frost development on plant surfaces at normal water freezing temperature (0oC). Even if the ice-minus strain does not win out, the amount of ice nucleate present from ice-plus P. syringae would be reduced due to competition. Decreased levels of frost generation at normal water freezing temperature would translate into a lowered quantity of crops lost due to frost damage, rendering higher crop yields overall.

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Controversy

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At the time of Dr. Lindow's work on ice-minus P. syringae, genetic engineering was considered to be very controversial. The controversy primarily revolved around fears of introducing new organisms that may permanently disrupt the ecosystem. The fear was that the introduction of ice-minus bacteria to the environment would eliminate bacterial and plant varieties. This was true in the case of the gypsy moth's accidental introduction into the U.S. Without a predator in the U.S., the gypsy moth is still causing overwhelming destruction to the hardwood forests of northeastern U.S.

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Naked DNA

Naked DNA is histone-free DNA that is passed from cell to cell during a gene transfer process called transformation or transfection. In transformation , purified or naked DNA is taken up by the recipient cell which will give the recipient cell a new characteristic or phenotype. Transfection differs from transformation since the DNA is not generally incorporated into the cell's genome, it is only transiently expressed.

In the field of DNA vaccines or genetic immunization, the term "naked DNA" was coined by Vical to mean DNA delivered free from agents which promote transfection. Research on the use of naked DNA for DNA vaccinations and gene therapy has shown some initial success, but have not yet resulted in any generally available therapy.

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VECTOR

In molecular biology, a vector is any vehicle used to transfer foreign genetic material to another cell.
The vector itself is generally a DNA sequence that consists of an insert (transgene) and a larger sequence that serves of the "backbone" of the vector. The purpose of a vector to transfer genetic information to another cell is typically to isolate, multiply, or express the insert in the target cell. Vectors called expression vectors (expression constructs) specifically are for the expression of the transgene in the target cell, and generally have a promoter sequence that drives expression of the transgene. Simpler vectors called transcription vectors are only capable of being transcribed but not translated: they can be replicated in a target cell but not expressed, unlike expression vectors. Transcription vectors are used to amplify their insert.
Insertion of a vector into the target cell is generally called transfection, although insertion of a viral vector is often called transduction.

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Characteristics:

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Two common vectors are plasmids and viral vectors.

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Plasmids

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Plasmids are double-stranded generally circular DNA sequences that are capable of automatically replicating in a host cell. Plasmid vectors minimalistically consist of an origin of replication that allows for semi-independent replication of the plasmid in the host and also the transgene insert. Modern plasmids generally have many more features, notably including a "multiple cloning site" which includes nucleotide overhangs for insertion of an insert, and multiple restriction enzyme consensus sites to either side of the insert. In the case of plasmids utilized as transcription vectors, incubating bacteria with plasmids generates hundreds or thousands of copies of the vector within the bacteria in hours, and the vectors can be extracted from the bacteria, and the multiple cloning site can be restricted by restriction enzymes to excise the hundredfold or thousandfold amplified insert. These plasmid transcription vectors characteristically lack crucial sequences that code for polyadenylation sequences and translation termination sequences in translated mRNAs, making expression of transcription vectors impossible.

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Viral vectors

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Viral vectors are generally genetically-engineered viruses carrying modified viral DNA or RNA that has been rendered noninfectious, but still contain viral promoters and also the transgene, thus allowing for translation of the transgene through a viral promoter. However, because viral vectors frequently are lacking infectious sequences, they require helper viruses or packaging lines for large-scale transfection. Viral vectors are often designed for permanent incorporation of the insert into the host genome, and thus leave distinct genetic markers in the host genome after incorporating the transgene. For example, retroviruses leave a characteristic retroviral integration pattern after insertion that is detectable and integrates that the viral vector has incorporated into the host genome.

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Transcription

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Transcription is a necessary component in all vectors: the premise of a vector is to multiply the insert (although expression vectors later also drive the translation of the multiplied insert). Thus, even stable expression is determined by stable transcription, which generally depends on promoters in the vector. However, expression vectors have a variety of expression patterns: constituitive (consistent expression) or inducible (expression only under certain conditions or chemicals). This expression is based on different promoter activities, not post-transcriptional activities, thus, these two different types of expression vectors depend on different types of promoters.
Viral promoters are often used for constitutive expression in plasmids and in viral vectors because they normally reliably force constant transcription in many cell lines and types.
Inducible expression depends on promoters that respond to the induction conditions: for example, the murine mammary tumor virus promoter only initiates transcription after dexamethasone application and the Drosphilia heat shock promoter only iniates after high temperatures.

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Expression

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Expression vectors require not only transcription but translation of the vector's insert, thus requiring more components than simpler transcription-only vectors. Expression vectors require sequences that encode for:

• Polyadenylation tail: Creates a polyadenylation tail at the end of the transcribed pre-mRNA that protects the mRNA from exonucleases and ensures transcriptional and translational termination: stabilizes mRNA production.
• Minimal UTR length: UTRs contain specific characteristics that may impede transcription or translation, and thus the shortest UTRs or none at all are encoded for in optimal expression vectors.
• Kozak sequence: Vectors should encode for a Kozak sequence in the mRNA, which assembles the ribosome for translation of the mRNA.

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Features

Modern vectors may encompass additional features besides the transgene insert and a backbone:

• Promoter: Necessary component for all vectors: used to drive transcription of the vector's transgene.
• Genetic markers: Genetic markers for viral vectors allow for confirmation that the vector has integrated with the host genomic DNA.
• Antibiotic resistance: Vectors with antibiotic-resistance open reading frames allow for identification of which cells have uptaken the vector through antibiotic selection.
• Epitope: Vector contains a sequence for a specific epitope that is incorporated into the expressed protein. Allows for antibody identification of cells expressing the vector.
• β-galactosidase: Vector's multiple cloning site contains sequence for β-galactosidase, an enzyme that digests galactose, to either side of the region intended for an insert. If the insert has not successfully ligated into the vector, cells expressing the empty vector will generate β-galactosidase and digest galactose. However, cells that express a vector with a transgene will have the coding sequence for β-galactosidase and be unable to digest galactose, and a subsequent color dye for galactose (X-gal) subsequently identifies cells expressing a vector with an insert, although it is unknown whether the insert is the intended one.
• Targeting sequence: Expression vectors may include encoding for a targeting sequence in the finished protein that directs the expressed protein to a specific organelle in the cell.

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Polymerase Chain Reaction (PCR - 2)

Variations on the basic PCR technique:

1. Allele-specific PCR: This diagnostic or cloning technique is used to identify or utilize single-nucleotide polymorphisms (SNPs) (single base differences in DNA). It requires prior knowledge of a DNA sequence, including differences between alleles, and uses primers whose 3' ends encompass the SNP. PCR amplification under stringent conditions is much less efficient in the presence of a mismatch between template and primer, so successful amplification with an SNP-specific primer signals presence of the specific SNP in a sequence. See SNP genotyping for more information.
2. Assembly PCR or Polymerase Cycling Assembly (PCA): Assembly PCR is the artificial synthesis of long DNA sequences by performing PCR on a pool of long oligonucleotides with short overlapping segments. The oligonucleotides alternate between sense and antisense directions, and the overlapping segments determine the order of the PCR fragments thereby selectively producing the final long DNA product.
3. Asymmetric PCR: Asymmetric PCR is used to preferentially amplify one strand of the original DNA more than the other. It finds use in some types of sequencing and hybridization probing where having only one of the two complementary stands is required. PCR is carried out as usual, but with a great excess of the primers for the chosen strand. Due to the slow (arithmetic) amplification later in the reaction after the limiting primer has been used up, extra cycles of PCR are required. A recent modification on this process, known as Linear-After-The-Exponential-PCR (LATE-PCR), uses a limiting primer with a higher melting temperature (Melting temperatureTm) than the excess primer to maintain reaction efficiency as the limiting primer concentration decreases mid-reaction.
4. Colony PCR: Bacterial colonies (E.coli) can be rapidly screened by PCR for correct DNA vector constructs. Selected bacterial colonies are picked with a sterile toothpick and dabbed into the PCR master mix or sterile water. The PCR is started with an extended time at 95˚C when standard polymerase is used or with a shortened denaturation step at 100˚C and special chimeric DNA polymerase.
5. Helicase-dependent amplification: This technique is similar to traditional PCR, but uses a constant temperature rather than cycling through denaturation and annealing/extension cycles. DNA Helicase, an enzyme that unwinds DNA, is used in place of thermal denaturation.
6. Hot-start PCR: This is a technique that reduces non-specific amplification during the initial set up stages of the PCR. The technique may be performed manually by heating the reaction components to the melting temperature (e.g., 95˚C) before adding the polymerase. Specialized enzyme systems have been developed that inhibit the polymerase's activity at ambient temperature, either by the binding of an antibody or by the presence of covalently bound inhibitors that only dissociate after a high-temperature activation step. Hot-start/cold-finish PCR is achieved with new hybrid polymerases that are inactive at ambient temperature and are instantly activated at elongation temperature.
7. Intersequence-specific (ISSR) PCR: a PCR method for DNA fingerprinting that amplifies regions between some simple sequence repeats to produce a unique fingerprint of amplified fragment lengths.
8. Inverse PCR: a method used to allow PCR when only one internal sequence is known. This is especially useful in identifying flanking sequences to various genomic inserts. This involves a series of DNA digestions and self ligation, resulting in known sequences at either end of the unknown sequence
9. Ligation-mediated PCR: This method uses small DNA linkers ligated to the DNA of interest and multiple primers annealing to the DNA linkers; it has been used for DNA sequencing, genome walking, and DNA footprinting.
10. Methylation-specific PCR (MSP): The MSP method was developed by Stephen Baylin and Jim Herman at the Johns Hopkins School of Medicine, and is used to detect methylation of CpG islands in genomic DNA. DNA is first treated with sodium bisulfite, which converts unmethylated cytosine bases to uracil, which is recognized by PCR primers as thymine. Two PCRs are then carried out on the modified DNA, using primer sets identical except at any CpG islands within the primer sequences. At these points, one primer set recognizes DNA with cytosines to amplify methylated DNA, and one set recognizes DNA with uracil or thymine to amplify unmethylated DNA. MSP using qPCR can also be performed to obtain quantitative rather than qualitative information about methylation.
11. Multiplex Ligation-dependent Probe Amplification (MLPA): permits multiple targets to be amplified with only a single primer pair, thus avoiding the resolution limitations of multiplex PCR.
12. Multiplex-PCR: The use of multiple, unique primer sets within a single PCR mixture to produce amplicons of varying sizes specific to different DNA sequences. By targeting multiple genes at once, additional information may be gained from a single test run that otherwise would require several times the reagents and more time to perform. Annealing temperatures for each of the primer sets must be optimized to work correctly within a single reaction, and amplicon sizes, i.e., their base pair length, should be different enough to form distinct bands when visualized by gel electrophoresis.
13. Nested PCR: increases the specificity of DNA amplification, by reducing background due to non-specific amplification of DNA. Two sets of primers are being used in two successive PCRs. In the first reaction, one pair of primers is used to generate DNA products, which besides the intended target, may still consist of non-specifically amplified DNA fragments. The product(s) are then used in a second PCR with a set of primers whose binding sites are completely or partially different from and located 3' of each of the primers used in the first reaction. Nested PCR is often more successful in specifically amplifying long DNA fragments than conventional PCR, but it requires more detailed knowledge of the target sequences.
14. Overlap-extension PCR: is a genetic engineering technique allowing the construction of a DNA sequence with an alteration inserted beyond the limit of the longest practical primer length.
15. Quantitative PCR (Q-PCR): is used to measure the quantity of a PCR product (preferably real-time). It is the method of choice to quantitatively measure starting amounts of DNA, cDNA or RNA. Q-PCR is commonly used to determine whether a DNA sequence is present in a sample and the number of its copies in the sample. Quantitative real-time PCR. It is often confusingly known as RT-PCR (Real Time PCR) or RQ-PCR. QRT-PCR or RTQ-PCR are more appropriate contractions. RT-PCR commonly refers to reverse transcription PCR (see below), which is often used in conjunction with Q-PCR. QRT-PCR methods use fluorescent dyes, such as Sybr Green, or fluorophore-containing DNA probes, such as TaqMan, to measure the amount of amplified product in real time.
16. RT-PCR: (Reverse Transcription PCR) is a method used to amplify, isolate or identify a known sequence from a cellular or tissue RNA. The PCR is preceded by a reaction using reverse transcriptase to convert RNA to cDNA. RT-PCR is widely used in expression profiling, to determine the expression of a gene or to identify the sequence of an RNA transcript, including transcription start and termination sites and, if the genomic DNA sequence of a gene is known, to map the location of exons and introns in the gene. The 5' end of a gene (corresponding to the transcription start site) is typically identified by an RT-PCR method, named RACE-PCR, short for Rapid Amplification of cDNA Ends.
17. TAIL-PCR: Thermal asymmetric interlaced PCR is used to isolate unknown sequence flanking a known sequence. Within the known sequence TAIL-PCR uses a nested pair of primers with differing annealing temperatures; a degenerate primer is used to amplify in the other direction from the unknown sequence.
18. Touchdown PCR: a variant of PCR that aims to reduce nonspecific background by gradually lowering the annealing temperature as PCR cycling progresses. The annealing temperature at the initial cycles is usually a few degrees (3-5˚C) above the Tm of the primers used, while at the later cycles, it is a few degrees (3-5˚C) below the primer Tm. The higher temperatures give greater specificity for primer binding, and the lower temperatures permit more efficient amplification from the specific products formed during the initial cycles.
19. PAN-AC: This method uses isothermal conditions for amplification, and may be used in living cells.

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Polymerase Chain Reaction (PCR - 1)

The polymerase chain reaction (PCR) is a technique widely used in molecular biology. It derives its name from one of its key components, a DNA polymerase used to amplify (i.e., replicate) a piece of DNA by in vitro enzymatic replication. As PCR progresses, the DNA thus generated is itself used as template for replication. This sets in motion a chain reaction in which the DNA template is exponentially amplified. With PCR it is possible to amplify a single or few copies of a piece of DNA across several orders of magnitude, generating millions or more copies of the DNA piece. PCR can be performed without restrictions on the form of DNA, and it can be extensively modified to perform a wide array of genetic manipulations.

Almost all PCR applications employ a heat-stable DNA polymerase, such as Taq polymerase, an enzyme derived from the bacterium Thermus aquaticus. This DNA polymerase enzymatically assembles a new DNA strand from DNA building blocks, the nucleotides, using single-stranded DNA as template and DNA oligonucleotides (also called DNA primers) required for initiation of DNA synthesis. The vast majority of PCR methods use thermal cycling, i.e., alternately heating and cooling the PCR sample to a defined series of temperature steps. These different temperature steps are necessary to bring about physical separation of the strands in a DNA double helix (DNA melting), and permit DNA synthesis by the DNA polymerase to selectively amplify the target DNA. The power and selectivity of PCR are primarily due to selecting primers that are highly complementary to the DNA region targeted for amplification, and to the thermal cycling conditions used.

Developed in 1983 by Kary Mullis, PCR is now a common and often indispensable technique used in medical and biological research labs for a variety of applications. These include DNA cloning for sequencing, DNA-based phylogeny, or functional analysis of genes; the diagnosis of hereditary diseases; the identification of genetic fingerprints (used in forensics and paternity testing); and the detection and diagnosis of infectious diseases. Mullis won the Nobel Prize for his work on PCR.

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PCR Principle And Procedure:

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PCR is used to amplify specific regions of a DNA strand (the DNA target). This can be a single gene, a part of a gene, or a non-coding sequence. Most PCR methods typically amplify DNA fragments of up to 10 kilo base pairs (kb), although some techniques allow for amplification of fragments up to 40 kb in size.
A basic PCR set up requires several components and reagents. These components include:

• DNA template that contains the DNA region (target) to be amplified.
• One or more primers, which are complementary to the DNA regions at the 5' (five prime) and 3' (three prime) ends of the DNA region.
• a DNA polymerase such as Taq polymerase or another DNA polymerase with a temperature optimum at around 70°C.
• Deoxynucleoside triphosphates (dNTPs; also very commonly and erroneously called deoxynucleotide triphosphates), the building blocks from which the DNA polymerases synthesizes a new DNA strand
• Buffer solution, providing a suitable chemical environment for optimum activity and stability of the DNA polymerase.
• Divalent cations, magnesium or manganese ions; generally Mg2+ is used, but Mn2+ can be utilized for PCR-mediated DNA mutagenesis, as higher Mn2+ concentration increases the error rate during DNA synthesis
• Monovalent cation potassium ions.

The PCR is commonly carried out in a reaction volume of 15-100 μl in small reaction tubes (0.2-0.5 ml volumes) in a thermal cycler. The thermal cycler allows heating and cooling of the reaction tubes to control the temperature required at each reaction step (see below). Many modern thermal cyclers make use of the Peltier effect which permits both heating and cooling of the block holding the PCR tubes simply by reversing the electric current. Thin-walled reaction tubes permit favorable thermal conductivity to allow for rapid thermal equilibration. Most thermal cyclers have heated lids to prevent condensation at the top of the reaction tube. Older thermocyclers lacking a heated lid require a layer of oil on top of the reaction mixture or a ball of wax inside the tube.

The PCR usually consists of a series of 20 to 35 repeated temperature changes called cycles; each cycle typically consists of 2-3 discrete temperature steps. Most commonly PCR is carried out with cycles that have three temperature steps (Fig. 2). The cycling is often preceded by a single temperature step (called hold) at a high temperature (>90°C), and followed by one hold at the end for final product extension or brief storage. The temperatures used and the length of time they are applied in each cycle depend on a variety of parameters. These include the enzyme used for DNA synthesis, the concentration of divalent ions and dNTPs in the reaction, and the melting temperature (Tm) of the primers.

• Initialization step: This step consists of heating the reaction to a temperature of 94-96°C (or 98°C if extremely thermostable polymerases are used), which is held for 1-9 minutes. It is only required for DNA polymerases that require heat activation by hot-start PCR.
• Denaturation step: This step is the first regular cycling event and consists of heating the reaction to 94-98°C for 20-30 seconds. It causes melting of DNA template and primers by disrupting the hydrogen bonds between complementary bases of the DNA strands, yielding single strands of DNA.
• Annealing step: The reaction temperature is lowered to 50-65°C for 20-40 seconds allowing annealing of the primers to the single-stranded DNA template. Typically the annealing temperature is about 3-5 degrees Celsius below the Tm of the primers used. Stable DNA-DNA hydrogen bonds are only formed when the primer sequence very closely matches the template sequence. The polymerase binds to the primer-template hybrid and begins DNA synthesis.
• Extension/elongation step: The temperature at this step depends on the DNA polymerase used; Taq polymerase has its optimum activity temperature at 75-80°C, and commonly a temperature of 72°C is used with this enzyme. At this step the DNA polymerase synthesizes a new DNA strand complementary to the DNA template strand by adding dNTP's that are complementary to the template in 5' to 3' direction, condensing the 5'-phosphate group of the dNTPs with the 3'-hydroxyl group at the end of the nascent (extending) DNA strand. The extension time depends both on the DNA polymerase used and on the length of the DNA fragment to be amplified. As a rule-of-thumb, at its optimum temperature, the DNA polymerase will polymerize a thousand bases in one minute.
• Final elongation: This single step is occasionally performed at a temperature of 70-74°C for 5-15 minutes after the last PCR cycle to ensure that any remaining single-stranded DNA is fully extended.
• Final hold: This step at 4-15°C for an indefinite time may be employed for short-term storage of the reaction.
• To check whether the PCR generated the anticipated DNA fragment (also sometimes referred to as the amplimer or amplicon), agarose gel electrophoresis is employed for size separation of the PCR products. The size(s) of PCR products is determined by comparison with a DNA ladder (a molecular weight marker), which contains DNA fragments of known size, run on the gel alongside the PCR products

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Northern Blot

The northern blot is a technique used in molecular biology research to study gene expression. It takes its name from the similarity of the procedure to the Southern blot procedure, named for biologist Edwin Southern, used to study DNA, with the key difference that, in the northern blot, RNA , rather than DNA, is the substance being analyzed by electrophoresis and detection with a hybridization probe. This technique was developed in 1977 by James Alwine, David Kemp, and George Stark at Stanford University.
The gels may be run on either agarose or denaturing polyacrylamide gels depending on the size of the RNA to be detected. A notable difference in the procedure in case of agarose gels, (as compared with the Southern blot) is the addition of formaldehyde which acts as a denaturant. For smaller fragments denaturing polyacrylamide urea gels are employed.
As in the Southern blot, the hybridization probe may be made from DNA or RNA.
A variant of the procedure known as the reverse northern blot was occasionally (although, infrequently) used. In this procedure, the substrate nucleic acid (that is affixed to the membrane) is a collection of isolated DNA fragments, and the probe is RNA extracted from a tissue and radioactively labelled.
The use of DNA microarrays that have come into widespread use in the late 1990s and early 2000s is more akin to the reverse procedure, in that they involve the use of isolated DNA fragments affixed to a substrate, and hybridization with a probe made from cellular RNA. Thus the reverse procedure, though originally uncommon, enabled the one-at-a-time study of gene expression using northern analysis to evolve into gene expression profiling, in which many (possibly all) of the genes in an organism may have their expression monitored.

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Southern Blot

A Southern blot is a method routinely used in molecular biology to check for the presence of a DNA sequence in a DNA sample. Southern blotting combines agarose gel electrophoresis for size separation of DNA with methods to transfer the size-separated DNA to a filter membrane for probe hybridization. The method is named after its inventor, the British biologist Edwin Southern. Other blotting methods (i.e., western blot, northern blot, southwestern blot) that employ similar principles, but using RNA or protein, have later been named in reference to Southern's name. As the technique was eponymously named, Southern blot should be capitalised, whereas northern and western blots should not.

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Method:

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1. Restriction endonucleases are used to cut high-molecular-weight DNA strands into smaller fragments.
2. The DNA fragments are then electrophoresed on an agarose gel to separate them by size.
3. If some of the DNA fragments are larger than 15 kb, then prior to blotting, the gel may be treated with an acid, such as dilute HCl, which depurinates the DNA fragments, breaking the DNA into smaller pieces, thus allowing more efficient transfer from the gel to membrane.
4. If alkaline transfer methods are used, the DNA gel is placed into an alkaline solution (typically containing sodium hydroxide) to denature the double-stranded DNA. The denaturation in an alkaline environment provides for improved binding of the negatively charged DNA to a positively charged membrane, separates it into single DNA strands for later hybridization to the probe (see below), and destroys any residual RNA that may still be present in the DNA.
5. A sheet of nitrocellulose (or, alternatively, nylon) membrane is placed on top of (or below, depending on the direction of the transfer) the gel. Pressure is applied evenly to the gel (either using suction, or by placing a stack of paper towels and a weight on top of the membrane and gel), to ensure good and even contact between gel and membrane. Buffer transfer by capillary action from a region of high water.
   potential to a region of low water potential (usually filter paper and paper tissues) is then used to move the DNA from the gel on to the membrane; ion exchange interactions bind the DNA to the membrane due to the negative charge of the DNA and positive charge of the membrane.
6. The membrane is then baked, i.e., exposed to high temperature (60 to 100 °C) (in the case of nitrocellulose) or exposed to ultraviolet radiation (nylon) to permanently and covalently crosslink the DNA to the membrane.
7. the membrane is then exposed to a hybridization probe—a single DNA fragment with a specific sequence whose presence in the target DNA is to be determined. The probe DNA is labelled so that it can be detected, usually by incorporating radioactivity or tagging the molecule with a fluorescent or chromogenic dye. In some cases, the hybridization probe may be made from RNA, rather than DNA. To ensure the specificity of the binding of the probe to the sample DNA, most common hybridization methods use salmon testes (sperm) DNA for blocking of the membrane surface and target DNA, deionized formamide, and detergents such as SDS to reduce non-specific binding of the probe.
8. After hybridization, excess probe is washed from the membrane, and the pattern of hybridization is visualized on X-ray film by autoradiography in the case of a radioactive or fluorescent probe, or by development of color on the membrane if a chromogenic detection method is used.

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Hybridization of the probe to a specific DNA fragment on the filter membrane indicates that this fragment contains DNA sequence that is complementary to the probe.
The transfer step of the DNA from the electrophoresis gel to a membrane permits easy binding of the labeled hybridization probe to the size-fractionated DNA

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Western Blot

The western blot (alternately, immunoblot) is a method to detect a specific protein in a given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) (Figure 1) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target protein. There are now many reagent companies that specialise in providing antibodies (both monoclonal and polyclonal antibodies) against many thousands of different proteins. Commercial antibodies can be expensive, though the unbound antibody can be reused between experiments. This method is used in the fields of molecular biology, biochemistry, immunogenetics and other molecular biology disciplines.
Other related techniques include using antibodies to detect proteins in tissues and cells by immunostaining and enzyme-linked immunosorbent assay (ELISA).

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Steps in a western blot:

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Tissue preparation

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Samples may be taken from whole tissue or from cell culture. In most cases, solid tissues are first broken down mechanically using a blender (for larger sample volumes), using a homogenizer (smaller volumes), or by sonication. Cells may also be broken open by one of the above mechanical methods. However, it should be noted that bacteria, virus or environmental samples can be the source of protein and thus western blotting is not restricted to cellular studies only.
Assorted detergents, salts, and buffers may be employed to encourage lysis of cells and to solubilize proteins. Protease and phosphatase inhibitors are often added to prevent the digestion of the sample by its own enzymes.
A combination of biochemical and mechanical techniques – including various types of filtration and centrifugation – can be used to separate different cell compartments and organelles.

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Gel electrophoresis:

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The proteins of the sample are separated using gel electrophoresis. Separation of proteins may be by isoelectric point (pI), molecular weight, electric charge, or a combination of these factors. The nature of the separation depends on the treatment of the sample and the nature of the gel

By far the most common type of gel electrophoresis employs polyacrylamide gels and buffers loaded with sodium dodecyl sulfate (SDS). SDS-PAGE (SDS polyacrylamide gel electrophoresis) maintains polypeptides in a denatured state once they have been treated with strong reducing agents to remove secondary and tertiary structure (e.g. S-S disulfide bonds to SH and SH) and thus allows separation of proteins by their molecular weight. Sampled proteins become covered in the negatively charged SDS and move to the positively charged electrode through the acrylamide mesh of the gel. Smaller proteins migrate faster through this mesh and the proteins are thus separated according to size (usually measured in kilo Daltons, kD). The concentration of acrylamide determines the resolution of the gel - the greater the acrylamide concentration the better the resolution of lower molecular weight proteins. The lower the acrylamide concentration the better the resolution of higher molecular weight proteins. Proteins travel only in one dimension along the gel for most blots.
Samples are loaded into wells in the gel. One lane is usually reserved for a marker or ladder, a commercially available mixture of proteins having defined molecular weights, typically stained so as to form visible, coloured bands. An example of a ladder is the GE Full Range Molecular weight ladder (Figure 1). When voltage is applied along the gel, proteins migrate into it at different speeds. These different rates of advancement (different electrophoretic mobilities) separate into bands within each lane.
It is also possible to use a two-dimensional (2-D) gel which spreads the proteins from a single sample out in two dimensions. Proteins are separated according to isoelectric point (pH at which they have neutral net charge) in the first dimension, and according to their molecular weight in the second dimension.

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Transfer:

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In order to make the proteins accessible to antibody detection, they are moved from within the gel onto a membrane made of nitrocellulose or PVDF. The membrane is placed on top of the gel, and a stack of tissue papers placed on top of that. The entire stack is placed in a buffer solution which moves up the paper by capillary action, bringing the proteins with it. Another method for transferring the proteins is called electroblotting and uses an electric current to pull proteins from the gel into the PVDF or nitrocellulose membrane. The proteins move from within the gel onto the membrane while maintaining the organization they had within the gel. As a result of this "blotting" process, the proteins are exposed on a thin surface layer for detection (see below). Both varieties of membrane are chosen for their non-specific protein binding properties (i.e. binds all proteins equally well). Protein binding is based upon hydrophobic interactions, as well as charged interactions between the membrane and protein. Nitrocellulose membranes are cheaper than PVDF, but are far more fragile and do not stand up well to repeated probings.
The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie or Ponceau S dyes. Coomassie is the more sensitive of the two, although Ponceau S's water solubility makes it easier to subsequently destain and probe the membrane as described below.

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Blotting:

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Since the membrane has been chosen for its ability to bind protein, and both antibodies and the target are proteins, steps must be taken to prevent interactions between the membrane and the antibody used for detection of the target protein. Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein - typically Bovine serum albumin (BSA) or non-fat dry milk (both are inexpensive), with a minute percentage of detergent such as Tween 20. The protein in the dilute solution attaches to the membrane in all places where the target proteins have not attached. Thus, when the antibody is added, there is no room on the membrane for it to attach other than on the binding sites of the specific target protein. This reduces "noise" in the final product of the Western blot, leading to clearer results, and eliminates false positives.

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Detection:

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During the detection process the membrane is "probed" for the protein of interest with a modified antibody which is linked to a reporter enzyme, which when exposed to an appropriate substrate drives a colorimetric reaction and produces a colour. For a variety of reasons, this traditionally takes place in a two-step process, although there are now one-step detection methods available for certain applications.

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Two step:

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Primary antibody
Antibodies are generated when a host species or immune cell culture is exposed to the protein of interest (or a part thereof). Normally, this is part of the immune response, whereas here they are harvested and used as sensitive and specific detection tools that bind the protein directly.
After blocking, a dilute solution of primary antibody (generally between 0.5 and 5 micrograms/ml) is incubated with the membrane under gentle agitation. Typically, the solution is comprised of buffered saline solution with a small percentage of detergent, and sometimes with powdered milk or BSA. The antibody solution and the membrane can be sealed and incubated together for anywhere from 30 minutes to overnight. It can also be incubated at different temperatures, with warmer temperatures being associated with more binding, both specific (to the target protein, the "signal") and non-specific ("noise").

Secondary antibody

After rinsing the membrane to remove unbound primary antibody, the membrane is exposed to another antibody, directed at a species-specific portion of the primary antibody. This is known as a secondary antibody, and due to its targeting properties, tends to be referred to as "anti-mouse," "anti-goat," etc. Antibodies come from animal sources (or animal sourced hybridoma cultures); an anti-mouse secondary will bind to just about any mouse-sourced primary antibody. This allows some cost savings by allowing an entire lab to share a single source of mass-produced antibody, and provides far more consistent results. The secondary antibody is usually linked to

biotin or to a reporter enzyme such as alkaline phosphatase or horseradish peroxidase. This means that several secondary antibodies will bind to one primary antibody and enhances the signal.
Most commonly, a horseradish peroxidase-linked secondary is used in conjunction with a chemiluminescent agent, and the reaction product produces luminescence in proportion to the amount of protein. A sensitive sheet of photographic film is placed against the membrane, and exposure to the light from the reaction creates an image of the antibodies bound to the blot.
As with the ELISPOT and ELISA procedures, the enzyme can be provided with a substrate molecule that will be converted by the enzyme to a colored reaction product that will be visible on the membrane (see the figure below with blue bands).
A third alternative is to use a radioactive label rather than an enzyme coupled to the secondary antibody, such as labeling an antibody-binding protein like Staphylococcus Protein A with a radioactive isotope of iodine. Since other methods are safer, quicker and cheaper this method is now rarely used.

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Colorimetric detection:

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The colorimetric detection method depends on incubation of the western blot with a substrate that reacts with the reporter enzyme (such as peroxidase) that is bound to the secondary antibody. This converts the soluble dye into an insoluble form of a different color that precipitates next to the enzyme and thereby stains the nitrocellulose membrane. Development of the blot is then stopped by washing away the soluble dye. Protein levels are evaluated through densitometry (how intense the stain is) or spectrophotometry.

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Chemiluminescence:

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chemiluminescent detection methods depend on incubation of the western blot with a substrate that will luminesce when exposed to the reporter on the secondary antibody. The light is then detected by photographic film, and more recently by CCD cameras which captures a digital image of the western blot. The image is analysed by densitometry, which evaluates the relative amount of protein staining and quantifies the results in terms of optical density. Newer software allows further data analysis such as molecular weight analysis if appropriate standards are used. So-called "enhanced chemiluminescent" (ECL) detection is considered to be among the most sensitive detection methods for blotting analysis.

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Radioactive detection:

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Radioactive labels do not require enzyme substrates, but rather allow the placement of medical X-ray film directly against the western blot which develops as it is exposed to the label and creates dark regions which correspond to the protein bands of interest (see image to the right). The importance of radioactive detections methods is decliningbecause it is very expensive, health and safety risks are high and ECL provides a useful alternative.

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Fluorescent detection:

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The fluorescently labeled probe is excited by light and the emission of the excitation is then detected by a photosensor such as CCD camera equipped with appropriate emission filters which captures a digital image of the western blot and allows further data analysis such as molecular weight analysis and a quantitative western blot analysis. Fluorescence is considered to be among the most sensitive detection methods for blotting analysis.

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Medical diagnostic applications:

• The confirmatory HIV test employs a western blot to detect anti-HIV antibody in a human serum sample. Proteins from known HIV-infected cells are separated and blotted on a membrane as above. Then, the serum to be tested is applied in the primary antibody incubation step; free antibody is washed away, and a secondary anti-human antibody linked to an enzyme signal is added. The stained bands then indicate the proteins to which the patient's serum contains antibody.
• A Western blot is also used as the definitive test for Bovine spongiform encephalopathy (BSE, commonly referred to as 'mad cow disease').

Text Source: Wikipedia Liscence NGU