The DNA sequences used in the construction of recombinant DNA molecules can originate from any  species . For example, plant DNA can be joined to bacterial DNA, or human DNA can be joined with fungal DNA. In addition, DNA sequences that do not occur anywhere in nature can be created by the  chemical synthesis of DNA  and incorporated into recombinant DNA molecules. Using recombinant DNA technology and synthetic DNA, any DNA sequence can be created and introduced into living organisms.


Proteins that can result from the expression of recombinant DNA within living cells are termed  recombinant proteins . When recombinant DNA encoding a protein is introduced into a host organism, the recombinant protein is not necessarily produced. [ 1 ]  Expression of foreign proteins requires the use of specialized expression vectors and often necessitates significant restructuring by
foreign coding sequences. [ 2 ]


Recombinant DNA differs from genetic recombination in that the former results from artificial methods while the latter is a normal biological process that results in the remixing of existing DNA sequences in essentially all organisms.


Molecular cloning is the laboratory process used to produce recombinant DNA. [ 3 ] [ 4 ] [ 5 ] [ 6 ]  It is one of two most widely used methods, along with  polymerase chain reaction  (PCR), used to direct the replication of any specific DNA sequence chosen by the experimentalist. There are two fundamental differences between the methods. One is that molecular cloning involves replication of the DNA within a living cell, while PCR replicates DNA in the test tube, free of living cells. The other difference is that cloning involves cutting and pasting DNA sequences, while PCR amplifies by copying an existing sequence.


Formation of recombinant DNA requires a  cloning vector , a DNA molecule that replicates within a living cell. Vectors are generally derived from  plasmids  or  viruses , and represent relatively small segments of DNA that contain necessary genetic signals for replication, as well as additional elements for convenience in inserting foreign DNA, identifying cells that contain recombinant DNA, and, where appropriate, expressing the foreign DNA. The choice of vector for molecular cloning depends on the choice of host organism, the size of the DNA to be cloned, and whether and how the foreign DNA is to be expressed. [ 7 ]  The DNA segments can be combined by using a variety of methods, such as restriction enzyme/ligase cloning or  Gibson assembly . [ citation needed ]


In standard cloning protocols, the cloning of any DNA fragment essentially involves seven steps: (1) Choice of host organism and cloning vector, (2) Preparation of vector DNA, (3) Preparation of DNA to be cloned, (4) Creation of recombinant DNA, (5) Introduction of recombinant DNA into the host organism, (6) Selection of organisms containing recombinant DNA, and (7) Screening for clones with desired DNA inserts and biological properties. [ 6 ]
 These steps are described in some detail in a related article ( molecular cloning ).


DNA expression requires the transfection of suitable host cells. Typically, either bacterial, yeast, insect, or mammalian cells (such as  Human Embryonic Kidney cells  or  CHO cells ) are used as host cells. [ 8 ]


Following transplantation into the host organism, the foreign DNA contained within the recombinant DNA construct may or may not be  expressed . That is, the DNA may simply be replicated without expression, or it may be  transcribed  and  translated  and a recombinant protein is produced. Generally speaking, expression of a foreign gene requires restructuring the gene to include sequences that are required for producing an  mRNA  molecule that can be used by the host's  translational apparatus  (e.g.  promoter ,  translational initiation signal , and  transcriptional terminator ). [ 9 ]  Specific changes to the host organism may be made to improve expression of the ectopic gene. In addition, changes may be needed to the coding sequences as well, to optimize translation, make the protein soluble, direct the recombinant protein to the proper cellular or extracellular location, and stabilize the protein from degradation. [ 10 ] [ 11 ] [ 12 ]


In most cases, organisms containing recombinant DNA have apparently normal  phenotypes . That is, their appearance, behavior and metabolism are usually unchanged, and the only way to demonstrate the presence of recombinant sequences is to examine the DNA itself, typically using a polymerase chain reaction (PCR) test. [ 13 ]  Significant exceptions exist, and are discussed below.


If the rDNA sequences encode a gene that is expressed, then the presence of RNA and/or protein products of the recombinant gene can be detected, typically using  RT-PCR  or  western hybridization  methods. [ 13 ]  Gross phenotypic changes are not the norm, unless the recombinant gene has been chosen and modified so as to generate biological activity in the host organism. [ 14 ]  Additional phenotypes that are encountered include toxicity to the host organism induced by the recombinant gene product, especially if it is  over-expressed  or expressed within inappropriate cells or tissues. [ citation needed ]


In some cases, recombinant DNA can have deleterious effects even if it is not expressed. One mechanism by which this happens is  insertional inactivation , in which the rDNA becomes inserted into a host cell's gene. In some cases, researchers use this phenomenon to " knock out " genes to determine their biological function and importance. [ 15 ]  Another mechanism by which rDNA insertion into chromosomal DNA can affect gene expression is by inappropriate activation of previously unexpressed host cell genes. This can happen, for example, when a recombinant DNA fragment containing an active promoter becomes located next to a previously silent host cell gene, or when a host cell gene that functions to restrain gene expression undergoes insertional inactivation by recombinant DNA. [ citation needed ]


Recombinant DNA is widely used in  biotechnology , medicine and research. Today, recombinant proteins and other products that result from the use of DNA technology are found in essentially every pharmacy, physician or veterinarian office, medical testing laboratory, and biological research laboratory. In addition, organisms that have been manipulated using recombinant DNA technology, as well as products derived from those organisms, have found their way into many farms,  supermarkets ,  home medicine cabinets , and even pet shops, such as those that sell  GloFish  and other  genetically modified animals .


The most common application of recombinant DNA is in basic research, in which the technology is important to most current work in the biological and biomedical sciences. [ 13 ]  Recombinant DNA is used to identify, map and sequence genes, and to determine their function. rDNA probes are employed in analyzing gene expression within individual cells, and throughout the tissues of whole organisms. Recombinant proteins are widely used as reagents in laboratory experiments and to generate antibody probes for examining protein synthesis within cells and organisms. [ 4 ]


Many additional practical applications of recombinant DNA are found in industry, food production, human and veterinary medicine, agriculture, and bioengineering. [ 4 ]  Some specific examples are identified below.


Found in  rennet ,  chymosin  is the enzyme responsible for hydrolysis of  κ - casein  to produce para- κ -casein and  glycomacropeptide , which is the first step in formation of  cheese , and subsequently  curd , and  whey . [ 16 ]  It was the first genetically engineered food additive used commercially. Traditionally, processors obtained chymosin from rennet, a preparation derived from the fourth stomach of milk-fed calves. Scientists engineered a non-pathogenic strain (K-12) of  E. coli  bacteria for large-scale laboratory production of the enzyme. This microbiologically produced recombinant enzyme, identical structurally to the calf derived enzyme, costs less and is produced in abundant quantities. Today about 60% of U.S. hard cheese is made with genetically engineered chymosin. In 1990, FDA granted chymosin " generally recognized as safe " (GRAS) status based on data showing that the enzyme was safe. [ 17 ]


Recombinant human  insulin  has almost completely replaced insulin obtained from animal sources (e.g. pigs and cattle) for the treatment of  type 1 diabetes . A variety of different recombinant insulin preparations are in widespread use. [ 18 ]  Recombinant insulin is synthesized by inserting the human insulin gene into  E. coli , or yeast (Saccharomyces cerevisiae) [ 19 ]  which then produces insulin for human use. [ 20 ]  Insulin produced by E. coli requires further  post translational modifications  (e.g. glycosylation) whereas yeasts are able to perform these modifications themselves by virtue of being more complex host organisms. The advantage of recombinant human insulin is after chronic use patients don't develop an immune defence against it the way animal sourced insulin stimulates the human immune system. [ 21 ]


Administered to patients whose pituitary glands generate insufficient quantities to support normal growth and development. Before recombinant HGH became available, HGH for therapeutic use was obtained from pituitary glands of cadavers. This unsafe practice led to some patients developing  Creutzfeldt–Jakob disease . Recombinant HGH eliminated this problem, and is now used therapeutically. [ 22 ]  It has also been misused as a performance-enhancing drug by athletes and others. [ 23 ] [ 24 ]


It is the recombinant form of  factor VIII , a blood-clotting protein that is administered to patients with the bleeding disorder  hemophilia , who are unable to produce factor VIII in quantities sufficient to support normal blood coagulation. [ 25 ]  Before the development of recombinant factor VIII, the protein was obtained by processing large quantities of human blood from multiple donors, which carried a very high risk of transmission of  blood borne infectious diseases , for example HIV and hepatitis B.


Hepatitis B  infection can be successfully controlled through the use of a recombinant  subunit   hepatitis B vaccine , which contains a form of the hepatitis B virus surface antigen that is produced in yeast cells. The development of the recombinant subunit vaccine was an important and necessary development because hepatitis B virus, unlike other common viruses such as  polio virus , cannot be grown  in vitro . [ 26 ]


Recombinant antibodies  (rAbs) are produced in vitro by the means of expression systems based on mammalian cells. Their monospecific binding to a specific epitope makes rAbs eligible not only for research purposes, but also as therapy options against certain cancer types, infections and autoimmune diseases. [ 27 ]


Each of the three widely used methods for  diagnosing HIV infection  has been developed using recombinant DNA. The antibody test ( ELISA  or  western blot ) uses a recombinant HIV protein to test for the presence of  antibodies  that the body has produced in response to an HIV infection. The DNA test looks for the presence of HIV genetic material using  reverse transcription polymerase chain reaction  (RT-PCR). Development of the RT-PCR test was made possible by the molecular cloning and sequence analysis of HIV genomes.  HIV testing page from US Centers for Disease Control (CDC)


Golden rice  is a recombinant variety of rice that has been engineered to express the enzymes responsible for  β-carotene  biosynthesis. [ 14 ]  This variety of rice holds substantial promise for reducing the incidence of  vitamin A deficiency  in the world's population. [ 28 ]  Golden rice is not currently in use, pending the resolution of regulatory and intellectual property issues. [ 29 ]


Commercial varieties of important agricultural crops (including soy, maize/corn, sorghum, canola, alfalfa and cotton) have been developed that incorporate a recombinant gene that results in resistance to the herbicide  glyphosate  (trade name  Roundup ), and simplifies weed control by glyphosate application. [ 30 ]  These crops are in common commercial use in several countries.


Bacillus thuringiensis  is a bacterium that naturally produces a protein ( Bt toxin ) with insecticidal properties. [ 28 ]  The bacterium has been applied to crops as an insect-control strategy for many years, and this practice has been widely adopted in agriculture and gardening. Recently, plants have been developed that express a recombinant form of the bacterial protein, which may effectively control some insect predators. Environmental issues associated with the use of these  transgenic  crops have not been fully resolved. [ 31 ]


The idea of recombinant DNA was first proposed by Peter Lobban, a graduate student of Prof.  Dale Kaiser  in the Biochemistry Department at Stanford University Medical School. [ 32 ]  The first publications describing the successful production and intracellular replication of recombinant DNA appeared in 1972 and 1973, from  Stanford  and  UCSF . [ 33 ] [ 34 ] [ 35 ] [ 36 ]  In 1980  Paul Berg , a professor in the Biochemistry Department at Stanford and an author on one of the first papers [ 33 ]  was awarded the Nobel Prize in Chemistry for his work on nucleic acids "with particular regard to recombinant DNA".  Werner Arber ,  Hamilton Smith , and  Daniel Nathans  shared the 1978  Nobel Prize in Physiology or Medicine  for the discovery of  restriction endonucleases  which enhanced the techniques of rDNA technology. [ citation needed ]


Stanford University  applied for a U.S. patent on recombinant DNA on November 4, 1974, listing the inventors as  Herbert W. Boyer  (professor at the  University of California, San Francisco ) and  Stanley N. Cohen  (professor at  Stanford University ); this patent, U.S. 4,237,224A, was awarded on December 2, 1980. [ 37 ] [ 38 ]  The first licensed drug generated using recombinant DNA technology was human insulin, developed by  Genentech  and licensed by  Eli Lilly and Company . [ 39 ]


Scientists associated with the initial development of recombinant DNA methods recognized that the potential existed for organisms containing recombinant DNA to have undesirable or dangerous properties. At the 1975  Asilomar Conference on Recombinant DNA , these concerns were discussed and a voluntary moratorium on recombinant DNA research was initiated for experiments that were considered particularly risky. This moratorium was widely observed until the US  National Institutes of Health  developed and issued formal guidelines for rDNA work. Today, recombinant DNA molecules and recombinant proteins are usually not regarded as dangerous. However, concerns remain about some organisms that express recombinant DNA, particularly when they leave the laboratory and are introduced into the environment or food chain. These concerns are discussed in the articles on  genetically modified organisms  and  genetically modified food controversies . Furthermore, there are concerns about the by-products in biopharmaceutical production, where recombinant DNA result in specific protein products. The major by-product, termed  host cell protein , comes from the host expression system and poses a threat to the patient's health and the overall environment. [ 40 ] [ 41 ]


RMCE  ( recombinase-mediated cassette exchange ) is a procedure in  reverse genetics  allowing the systematic, repeated modification of higher eukaryotic genomes by targeted integration, based on the features of  site-specific recombination  processes (SSRs). For RMCE, this is achieved by the clean exchange of a preexisting  gene cassette  for an analogous cassette carrying the "gene of interest" (GOI).


The genetic modification of mammalian cells is a standard procedure for the production of correctly modified proteins with pharmaceutical relevance. To be successful, the transfer and expression of the  transgene  has to be highly efficient and should have a largely predictable outcome. Current developments in the field of  gene therapy  are based on the same principles. Traditional procedures used for transfer of GOIs are not sufficiently reliable, mostly because the relevant  epigenetic  influences have not been sufficiently explored: transgenes integrate into chromosomes with low efficiency and at  loci  that provide only sub-optimal conditions for their  expression . As a consequence the newly introduced information may not be realized (expressed), the gene(s) may be lost and/or re-insert and they may render the target cells in unstable state. It is exactly this point where RMCE enters the field. The procedure was introduced in 1994  [ 1 ]  and it uses the tools yeasts and  bacteriophages [ 2 ]  have evolved for the efficient replication of important genetic information:


Most yeast strains contain circular, plasmid-like DNAs called "two-micron circles". The persistence of these entities is granted by a recombinase called  "flippase" or "Flp" . Four  monomers  of this enzyme associate with two identical short (48 bp) target sites, called  FRT  ("flip-recombinase targets"), resulting in their  crossover . The outcome of such a process depends on the  relative orientation  of the participating FRTs leading to


This spectrum of options could be extended significantly by the generation of spacer mutants for  extended 48 bp  FRT  sites  (cross-hatched half-arrows in Figure 1). Each mutant Fn recombines with an identical mutant Fn with an efficiency equal to the wildtype sites (F x F). A cross-interaction (F x Fn) is strictly prevented by the particular design of these components. This sets the stage for the situation depicted in Figure 1A:


First applied for the  Tyr-recombinase  Flp, this novel procedure is not only relevant to the rational construction of biotechnologically significant cell lines, but it also finds increasing use for the systematic generation of  stem cells . Stem cells can be used to replace damaged tissue or to generate transgenic animals with largely pre-determined properties.


It has been previously established that coexpression of both Cre and Flp recombinases catalyzes the exchange of sequences flanked by single loxP and FRT sites integrated into the genome at a random location. However, these studies did not explore whether such an approach could be used to modify conditional mouse alleles carrying single or multiple loxP and FRT sites. dual RMCE (dRMCE; Osterwalder et al., 2010) was recently developed as a re-engineering tool applicable to the vast numbers of mouse conditional alleles that harbor wild-type loxP and FRT sites and therefore are not compatible with conventional RMCE. The general dRMCE strategy takes advantage of the fact that most conditional alleles encode a selection cassette flanked by FRT sites, in addition to loxP sites that flank functionally relevant exons ('floxed' exons). The FRT-flanked selection cassette is in general placed outside the loxP-flanked region, which renders these alleles directly compatible with dRMCE. Simultaneous expression of Cre and Flp recombinases induces cis recombination and formation of the deleted allele, which then serves as a 'docking site' at which to insert the replacement vector by trans recombination. The correctly replaced locus would encode the custom modification and a different drug-selection cassette flanked by single loxP and FRT sites. dRMCE therefore appears as a very efficient tool for targeted re-engineering of thousands of mouse alleles produced by the IKMC consortium.


Multiplexing setups rely on the fact that each F-Fn pair (consisting of a wildtype  FRT  site and a mutant called "n") or each Fn-Fm pair (consisting of two mutants, "m" and "n") constitutes a unique "address" in the genome. A prerequisite are differences in four out of the eight spacer positions (see Figure 1B). If the difference is below this threshold, some cross-interaction between the mutants may occur leading to a faulty deletion of the sequence between the heterospecific (Fm/Fn or F/Fn) sites.


13 FRT-mutants  [ 3 ] [ 4 ]  have meanwhile become available, which permit the establishment of several unique genomic addresses  side-by side  (for instance F-Fn and Fm-Fo). These addresses will be recognized by donor plasmids that have been designed according to the same principles, permitting successive (but also synchronous) modifications at the predetermined  loci . These modifications can be driven to completion in case the compatible donor plasmid(s) are provided at an excess (mass-action principles). Figure 2 illustrates one use of the multiplexing principle: the stepwise extension of a coding region in which a basic expression unit is provided with  genomic insulators ,  enhancers , or other  cis -acting elements.


A recent variation of the general concept is based on PhiC31  (an integrase of the Ser-class) , which permits introduction of another RMCE target at a secondary site  after  the first RMCE-based modification has occurred. This is due to the fact that each phiC31-catalyzed exchange destroys the attP and attB sites it has addressed  [ 2 ]  converting them to  att R and  att L  product sites, respectively. While these changes permit the subsequent mounting of new (and most likely remote) targets, they do not enable addressing several RMCE targets  in parallel , nor do they permit "serial RMCE", i.e. successive, stepwise modifications at a given genomic locus.


This is different for Flp-RMCE, for which the post-RMCE status of  FRT s corresponds to their initial state. This property enables the intentional, repeated mobilization of a target cassette by the addition of a new donor plasmid with compatible architecture. These "multiplexing-RMCE" options open unlimited possibilities for serial- and parallel specific modifications of pre-determined RMCE-targets  [ 5 ]


Generation of transgenic knock-out/-in mice and their genetic modification by RMCE. [ 6 ] [ 7 ]


Insertion of a target cassette in a mammalian host cell line (CHO DG44 in suspension culture) and exchange with an ER stress reporter construct via targeted integration (RMCE). [ 8 ]


Recombineering  ( recombi nation-mediated genetic engi neering ) [ 1 ]   is a genetic and  molecular biology  technique based on  homologous recombination  systems, as opposed to the older/more common method of using  restriction enzymes  and  ligases  to combine DNA sequences in a specified order. Recombineering is widely used for bacterial genetics, in the generation of target vectors for making a conditional  mouse knockout , and for modifying DNA of any source often contained on a  bacterial artificial chromosome  (BAC), among other applications.


Although developed in bacteria, much of the inspiration for recombineering techniques came from methods first developed in  Saccharomyces cerevisiae [ 2 ]  where a  linear plasmid  was used to target genes or clone genes off the chromosome.  In addition, recombination with single-strand  oligonucleotides  (oligos) was first shown in  Saccharomyces cerevisiae . [ 3 ]  Recombination was observed to take place with oligonucleotides as short as 20 bases.


Recombineering is based on homologous recombination in  Escherichia coli  mediated by  bacteriophage  proteins, either RecE/RecT from Rac prophage  [ 4 ]  or Redαβδ from  bacteriophage lambda . [ 5 ] [ 6 ]  The  lambda Red recombination system is now most commonly used and the first demonstrations of Red  in vivo  genetic engineering were independently made by Kenan Murphy [ 7 ]  and Francis Stewart. [ 4 ] [ 5 ]  However, Murphy's experiments required expression of RecA and also employed long homology arms. Consequently, the implications for a new DNA engineering technology were not obvious. The Stewart lab showed that these homologous recombination systems mediate efficient recombination of linear DNA molecules flanked by homology sequences as short as 30 base pairs (40-50 base pairs are more efficient) into target  DNA  sequences in the absence of RecA. Now the homology could be provided by oligonucleotides made to order, and standard  recA  cloning hosts could be used, greatly expanding the utility of recombineering.


Recombineering utilizes linear DNA substrates that are either double-stranded (dsDNA) or single-stranded (ssDNA). Most commonly, dsDNA recombineering has been used to create gene replacements, deletions, insertions, and inversions. Gene cloning [ 6 ] [ 8 ]  and gene/protein tagging (His tags etc., see  [ 9 ] ) is also common. For gene replacements or deletions, usually a cassette encoding a drug-resistance gene is made by PCR using bi-partite primers. These primers consist of (from 5’→3’) 50 bases of homology to the target region, where the cassette is to be inserted, followed by 20 bases to prime the drug resistant cassette. The exact junction sequence of the final construct is determined by primer design. [ 10 ] [ 11 ]  These events typically occur at a frequency of approximately 10 4 /10 8 cells that survive  electroporation . Electroporation is the method used to transform the linear substrate into the recombining cell.


In some cases, one desires a deletion with no marker left behind, to make a gene fusion, or to make a point mutant in a gene.  This can be done with two rounds of recombination. [ 12 ]  In the first stage of recombineering, a selection marker on a cassette is introduced to replace the region to be modified. In the second stage, a second counterselection marker (e.g. sacB) on the cassette is selected against following introduction of a target fragment containing the desired modification. Alternatively, the target fragment could be flanked by  loxP  or  FRT  sites, which could be removed later simply by the expression of the  Cre  or FLP recombinases, respectively.
A novel selection marker "mFabI" was also developed to increase recombineering efficiency. [ 13 ]


Recombineering with ssDNA provided a breakthrough both in the efficiency of the reaction and the ease of making point mutations. [ 1 ]  This technique was further enhanced by the discovery that by avoiding the methyl-directed mismatch repair system, the frequency of obtaining recombinants can be increased to over 10 7 /10 8  viable cells. [ 14 ]  This frequency is high enough that alterations can now be made without selection. With optimized protocols, over 50% of the cells that survive electroporation contain the desired change. Recombineering with ssDNA only requires the Red Beta protein; Exo, Gamma and the host recombination proteins are not required. As proteins homologous to Beta and RecT are found in many bacteria and bacteriophages (>100 as of February 2010), recombineering is likely to work in many different bacteria. [ 15 ]  Thus, recombineering with ssDNA is expanding the genetic tools available for research in a variety of organisms. To date, recombineering has been performed in  E. coli ,  S. enterica ,  Y. pseudotuberculosis ,   S. cerevisiae  and  M. tuberculosis . [ 16 ] [ 17 ] [ 18 ] [ 19 ] [ 20 ] [ 21 ]


In the year 2010, it has been demonstrated that ssDNA recombination can occur in the absence of known recombination functions. [ 22 ]  Recombinants were found at up to 10 4 /10 8  viable cells. This Red-independent activity has been demonstrated in  P. syringae ,  E. coli ,  S. enterica serovar typhimurium  and  S. flexneria .


The biggest advantage of recombineering is that it obviates the need for conveniently positioned  restriction sites , whereas in conventional genetic engineering, DNA modification is often compromised by the availability of unique restriction sites. In engineering large constructs of >100 kb, such as the  Bacterial Artificial Chromosomes  (BACs), or chromosomes, recombineering has become a necessity. Recombineering can generate the desired modifications without leaving any 'footprints' behind. It also forgoes multiple  cloning  stages for generating intermediate  vectors  and therefore is used to modify DNA constructs in a relatively short time-frame. The homology required is short enough that it can be generated in synthetic oligonucleotides and recombination with short oligonucleotides themselves is incredibly efficient. Recently, recombineering has been developed for high throughput DNA engineering applications termed 'recombineering pipelines'. [ 23 ]  Recombineering pipelines support the large scale production of BAC  transgenes  and  gene targeting  constructs for functional genomics programs such as EUCOMM (European Conditional Mouse Mutagenesis Consortium) and KOMP (Knock-Out Mouse Program). Recombineering has also been automated, a process called "MAGE" -Multiplex Automated Genome Engineering, in the Church lab. [ 24 ]  With the development of CRISPR technologies, construction of  CRISPR interference  strains in  E. coli  requires only one-step oligo recombineering, providing a simple and easy-to-implement tool for gene expression control. [ 12 ] [ 25 ]
"Recombineering tools" and laboratory protocols have also been implemented for a number of plant species. These tools and procedures are customizable, scalable, and freely available to all researchers. [ 26 ]


In  molecular biology , a  reporter gene  (often simply  reporter ) is a  gene  that researchers attach to a  regulatory sequence  of another gene of interest in  bacteria ,  cell culture , animals or plants. Such genes are called reporters because the characteristics they confer on organisms expressing them are easily identified and measured, or because they are  selectable markers . Reporter genes are often used as an indication of whether a certain gene has been taken up by or expressed in the cell or organism population.


To introduce a reporter gene into an organism, scientists place the reporter gene and the gene of interest in the same  DNA construct  to be inserted into the cell or organism. For  bacteria  or  prokaryotic cells  in culture, this is usually in the form of a circular DNA molecule called a  plasmid . For  viruses , this is known as a  viral vector . It is important to use a reporter gene that is not natively expressed in the cell or organism under study, since the expression of the reporter is being used as a marker for successful uptake of the gene of interest. [ 1 ]


Commonly used reporter genes that induce visually identifiable characteristics usually involve  fluorescent  and  luminescent  proteins. Examples include the gene that encodes jellyfish  green fluorescent protein  (GFP), which causes  cells  that express it to glow green under blue or ultraviolet light, [ 2 ]  the enzyme  luciferase , which catalyzes a reaction with  luciferin  to produce light, and the red fluorescent protein from the gene  dsRed  [ fr ] . [ 3 ] [ 4 ] [ 5 ] [ 6 ] [ 7 ]  The  GUS  gene has been commonly used in plants but luciferase and  GFP  are becoming more common. [ 8 ] [ 9 ]


A common reporter in bacteria is the  E. coli   lacZ  gene, which encodes the protein  beta-galactosidase . [ 10 ]  This enzyme causes bacteria expressing the gene to appear blue when grown on a medium that contains the  substrate analog   X-gal . An example of a selectable marker which is also a reporter in bacteria is the  chloramphenicol acetyltransferase  (CAT) gene, which confers resistance to the antibiotic  chloramphenicol . [ 11 ]


Many methods of  transfection  and  transformation  – two ways of expressing a foreign or modified gene in an organism – are effective in only a small percentage of a population subjected to the techniques. [ 13 ] [ 14 ]  Thus, a method for identifying those few successful gene uptake events is necessary. Reporter genes used in this way are normally expressed under their own  promoter  (DNA regions that initiates gene transcription) independent from that of the introduced gene of interest; the reporter gene can be expressed  constitutively  (that is, it is "always on") or inducibly with an external intervention such as the introduction of  Isopropyl β-D-1-thiogalactopyranoside  (IPTG) in the β-galactosidase system. [ 10 ]  As a result, the reporter gene's expression is independent of the gene of interest's expression, which is an advantage when the gene of interest is only expressed under certain specific conditions or in tissues that are difficult to access. [ 1 ]


In the case of selectable-marker reporters such as CAT, the transfected population of bacteria can be grown on a substrate that contains  chloramphenicol . Only those cells that have successfully taken up the construct containing the CAT gene will survive and multiply under these conditions. [ 11 ]


Reporter genes can be used to assay for the expression of a gene of interest that is normally difficult to quantitatively assay. [ 1 ]  Reporter genes can produce a protein that has little obvious or immediate effect on the cell culture or organism. They are ideally not present in the native genome to be able to isolate reporter gene expression as a result of the gene of interest's expression. [ 1 ] [ 15 ]


To activate reporter genes, they can be expressed  constitutively , where they are directly attached to the gene of interest to create a  gene fusion . [ 16 ]  This method is an example of using  cis -acting  elements where the two genes are under the same promoter elements and are  transcribed  into a single  messenger RNA  molecule. The  mRNA  is then  translated  into protein. It is important that both proteins be able to properly  fold  into their active conformations and interact with their substrates despite being fused. In building the DNA construct, a segment of DNA coding for a flexible polypeptide linker region is usually included so that the reporter and the gene product will only minimally interfere with one another. [ 17 ] [ 18 ]  Reporter genes can also be expressed by  induction  during growth. In these cases,  trans -acting  elements, such as  transcription factors  are used to express the reporter gene. [ 19 ] [ 20 ]


Reporter gene assay have been increasingly used in  high throughput screening  (HTS) to identify small molecule inhibitors and activators of protein targets and pathways for  drug discovery  and  chemical biology .  Because the reporter enzymes themselves (e.g. firefly  luciferase ) can be direct targets of small molecules and confound the interpretation of HTS data, novel coincidence reporter designs incorporating artifact suppression have been developed. [ 21 ] [ 22 ]


Reporter genes can be used to assay for the activity of a particular promoter in a cell or organism. [ 23 ]  In this case there is no separate "gene of interest"; the reporter gene is simply placed under the control of the target promoter and the reporter gene product's activity is quantitatively measured. The results are normally reported relative to the activity under a "consensus" promoter known to induce strong gene expression. [ 24 ]


A more complex use of reporter genes on a large scale is in  two-hybrid screening , which aims to identify proteins that natively interact with one another  in vivo . [ 25 ]


A  restriction digest  is a procedure used in  molecular biology  to prepare  DNA  for analysis or other processing.  It is sometimes termed  DNA fragmentation , though this term is used for other procedures as well. In a restriction digest, DNA molecules are cleaved at specific restriction sites of 4-12 nucleotides in length by use of restriction enzymes which recognize these sequences.  [ 1 ]


The resulting digested DNA is very often selectively amplified using  polymerase chain reaction  (PCR), making it more suitable for analytical techniques such as  agarose gel electrophoresis , and  chromatography .  It is used in  genetic fingerprinting , plasmid  subcloning , and  RFLP analysis .


A given  restriction enzyme  cuts DNA segments within a specific  nucleotide sequence , at what is called a  restriction site . These  recognition sequences  are typically four, six, eight, ten, or twelve nucleotides long and generally  palindromic  (i.e. the same nucleotide sequence in the 5' – 3' direction). Because there are only so many ways to arrange the four nucleotides that compose DNA (Adenine, Thymine, Guanine and Cytosine) into a four- to twelve-nucleotide sequence, recognition sequences tend to occur by chance in any long sequence. Restriction enzymes specific to hundreds of distinct sequences have been identified and synthesized for sale to laboratories, and as a result, several potential "restriction sites" appear in almost any gene or locus of interest on any chromosome. Furthermore, almost all artificial plasmids include a (often entirely synthetic)  polylinker  (also called "multiple cloning site") that contains dozens of restriction enzyme recognition sequences within a very short segment of DNA.  This allows the insertion of almost any specific fragment of DNA into plasmid  vectors , which can be efficiently "cloned" by insertion into replicating bacterial cells.


After restriction digest, DNA can then be analysed using  agarose gel electrophoresis .  In gel electrophoresis, a sample of DNA is first "loaded" onto a slab of agarose gel (literally pipetted into small wells at one end of the slab).  The gel is then subjected to an electric field, which draws the negatively charged DNA across it.  The molecules travel at different rates (and therefore end up at different distances) depending on their net charge (more highly charged particles travel further), and size (smaller particles travel further).  Since none of the four nucleotide bases carry any charge, net charge becomes insignificant and size is the main factor affecting rate of diffusion through the gel.  Net charge in DNA is produced by the  sugar-phosphate backbone .  This is in contrast to proteins, in which there is no "backbone", and net charge is generated by different combinations and numbers of charged  amino acids .


Restriction digest is most commonly used as part of the process of the  molecular cloning  of DNA fragment into a  vector  (such as a  cloning vector  or an  expression vector ).  The vector typically contains a  multiple cloning site  where many restriction site may be found, and a foreign piece of DNA may be inserted into the vector by first cutting the restriction sites in the vector as well the DNA fragment, followed by  ligation  of the DNA fragment into the vector.


Restriction digests are also necessary for performing any of the following analytical techniques:


There are numerous types of restriction enzymes, each of which will cut DNA differently. Most commonly used restriction enzymes are Type II restriction endonuclease (See article on  Restriction enzymes  for examples). There are some that cut a three base pair sequence while others can cut four, six, and even eight. Each enzyme has distinct properties that determine how efficiently it can cut and under what conditions. Most manufacturers that produce such enzymes will often provide a specific buffer solution that contains the unique mix of cations and other components that aid the enzyme in cutting as efficiently as possible. Different restriction enzymes may also have different optimal temperatures under which they function.


Note that for efficient digest of DNA, the restriction site should not be located at the very end of a DNA fragment.  The restriction enzymes may require a minimum number of base pairs between the restriction site and the end of the DNA for the enzyme to work efficiently. [ 2 ]  This number may vary between enzymes, but for most commonly used restriction enzymes around 6–10 base pair is sufficient.


Restriction sites , or  restriction recognition sites , are located on a  DNA  molecule containing specific (4-8 base pairs in length [ 1 ] ) sequences of  nucleotides , which are recognized by  restriction enzymes . These are generally  palindromic sequences [ 2 ]  (because restriction enzymes usually bind as  homodimers ), and a particular restriction enzyme may cut the sequence between two nucleotides within its recognition site, or somewhere nearby.


For example, the common restriction enzyme  EcoRI  recognizes the palindromic sequence GAATTC and cuts between the G and the A on both the top and bottom strands. This leaves an overhang (an end-portion of a  DNA  strand with no attached complement) known as a sticky end [ 2 ]  on each end of AATT. The overhang can then be used to ligate in (see  DNA ligase ) a piece of DNA with a complementary overhang (another EcoRI-cut piece, for example).


Some restriction enzymes cut DNA at a restriction site in a manner which leaves no overhang, called a blunt end. [ 2 ]  Blunt ends are much less likely to be ligated by a DNA ligase because the blunt end doesn't have the overhanging base pair that the enzyme can recognize and match with a complementary pair. [ 3 ]  Sticky ends of DNA however are more likely to successfully bind with the help of a DNA ligase because of the exposed and unpaired nucleotides. For example, a sticky end trailing with AATTG is more likely to bind with a ligase than a blunt end where both the 5' and 3' DNA strands are paired. In the case of the example the AATTG would have a complementary pair of TTAAC which would reduce the functionality of the DNA ligase enzyme. [ 4 ]


Restriction sites can be used for multiple applications in molecular biology such as identifying restriction fragment length polymorphisms ( RFLPs ). Restriction sites are also important consideration to be aware of when designing  plasmids .


Several databases exist for restriction sites and enzymes, of which the largest noncommercial database is REBASE. [ 5 ] [ 6 ]  Recently, it has been shown that statistically significant  nullomers  (i.e. short absent motifs which are highly expected to exist) in virus genomes are restriction sites indicating that viruses have probably got rid of these motifs to facilitate invasion of bacterial hosts. [ 7 ]   Nullomers Database  contains a comprehensive catalogue of minimal absent motifs many of which might potentially be not-yet-known restriction motifs.


R ibo n ucleo p rotein  Networks Analyzed by  M ut a tional  P rofiling  (RNP-MaP)  is a strategy for probing  RNA - protein  networks and protein binding sites at a  nucleotide  resolution. Information about RNP assembly and function can facilitate a better understanding of biological mechanisms. RNP-MaP uses  NHS - diazirine  (SDA), a hetero-bifunctional crosslinker, to freeze RNA-bound proteins in place. Once the RNA-protein  crosslinks  are formed, MaP  reverse transcription [ 1 ]  is then conducted to reversely transcribe the protein-bound RNAs as well as introduce mutations at the site of RNA-protein crosslinks. Sequencing results of the  cDNAs  reveal information about both protein-RNA interaction networks and protein binding sites. [ 2 ]


RNA-MaP involves three major components: [ 2 ]


Long-wavelength UV and SDA reagents are first supplied to living cells to crosslink protein residues with RNA by forming amide bonds between amine groups of lysine (or  arginine ) residues and succinimidyl esters. Next, cells containing crosslinked RNPs are  lysed  and the RNA-bound proteins are digested into peptide adducts. MaP reverse transcription is then performed to label the protein-RNA binding sites through peptide adduct-induced  mutations .  Sequencing  of the mutation-containing cDNA product will reveal the mutation sites (or RNP-MaP sites) and the correlations between the RNP-MaP sites are computationally determined using 3-nucleotide windows. [ 2 ] [ 4 ]


RNP-MaP sites are defined as protein bound nucleotides. SDA and UV treated and UV only treated sample sequence reads are aligned and mutations are counted using ShapeMapper2 software. [ 5 ]  The SDA or RNP-MaP reactivity for a nucleotide is the ratio of the crosslinked (SDA and UV treated) mutational frequency to the un-crosslinked (UV only) mutation frequency. Using differential mutational signatures, RNP-MaP sites are identified based on universal  normalization  factors and thresholds on each RNA nucleotide (U, A, C, and G) derived from analysis of ribonucleoproteins of known structure. [ 2 ]


A nucleotide is identified as a RNP-MaP site if it passes three filters:


Protein-RNA interactions networks are identified using RNP-MaP  correlations  since multiple crosslink sites can be detected for a single RNA molecule. RNP-MaP correlations provide a complementary measure of protein binding to RNA independent of RNP-MaP sites. [ 2 ]  They are identified using a G-test framework known as RingMapper. [ 4 ]


RNP-MaP correlations require a single RNA molecule to form at least two crosslinks and arise from any of three scenarios:


Using RNP-MaP correlations, a network of protein-RNA interaction sites is found and can then be used for functional analysis.


CLIP  analyzes protein interactions with RNA by combining UV cross-linking and  immunoprecipitation . CLIP-based techniques are able to map RNA binding protein binding sites of interest on a genome-wide scale. [ 6 ]


There are many CLIP-based methods including:


Quantitative  mass spectrometry  (MS) (or  quantitative proteomics ) can be used to discover RNA-binding proteins (RBPs) bound to RNA. Labeling MS methods involve the differential use of stable isotope labels or chemical tagging of proteins in samples and controls. This is used to obtain enrichment scores and true binding partners through the ratio of labeled peptides. [ 12 ]  Label-free MS methods are able to identify proteins in samples and controls. In order to distinguish true binding partners for nonspecific proteins, analytical tools used alongside spectral count data from non-quantitative MS are used to score the probability of a true RBP-RNA interaction [ 13 ]


RNP-MaP can help reveal functionally important RNA-protein binding networks through binding site density and interconnectivity independent of previous knowledge of interacting proteins. Because of the unbiased nature of the analysis, RNP-MaP is able to detect conserved RNA-protein interactions between species. [ 2 ]


RNP-MaP is also able to facilitate the characterization of functionally critical elements in large non-coding RNAs or even viral RNAs. [ 2 ]


As a standalone technique, RNP-MaP cannot be used to determine protein-RNA binding mechanisms or protein identities. In order to do so, RNP-MaP must be used in conjunction with other techniques such as CLIP and mass spectrometry. [ 2 ]


RNP-MaP requires extremely high  read-depths  for analysis. To identify RNP-MaP sites, 1000x  sequencing coverage  is required, while RNP-MaP correlation sites require 10,000x sequencing coverage. [ 2 ]


There are severe limitations on the ability to characterize RNP-MaP correlations between distant (>500 nucleotides) RNP-MaP sites. This is due to limitations of MaP reverse transcription processivity (500-600 nucleotides) and sequencing instrument clustering (<1,000 nucleotides). [ 2 ]


Ribotyping  is a molecular technique for bacterial identification and characterization that uses information from rRNA-based phylogenetic analyses. [ 1 ]  It is a rapid and specific method widely used in clinical diagnostics and analysis of microbial communities in food, water, and beverages. [ 1 ]


All bacteria have  ribosomal genes , but the exact sequence is unique to each species, serving as a genetic fingerprint. Therefore, sequencing the particular 16S gene and comparing it to a database would yield identification of the particular species. [ 2 ]


Ribotyping involves the digestion of bacterial genomic DNA with specific  restriction enzymes . Each restriction enzyme cuts DNA at a specific nucleotide sequence, resulting in fragments of different lengths. [ 3 ]


Those fragments are then run on a  Gel electrophoresis , where they are separated according to size: the application of electrical field to the gel in which they are suspended causes the movement of DNA fragments (all negatively charged due to the presence of phosphate groups) through a matrix towards the positively charged end of the field. Small fragments move more easily and rapidly through the matrix, reaching a bigger distance from the starting position than larger fragments.


Following the separation in the gel matrix, the DNA fragments are moved onto nylon membranes and hybridized with a labelled 16S or 23S rRNA probe. This way only the fragments coding for such rRNA are visualised and can be analyzed. [ 4 ]   The pattern is then digitized and used to identify the origin of the DNA by a comparison with reference organisms in a computer database. [ 1 ]


Conceptually, ribotyping is similar to probing restriction fragments of chromosomal DNA with  cloned probes  (randomly cloned probes or probes derived from a specific coding sequence such as that of a  virulence factor ). [ 5 ]


RNA extraction  is the purification of  RNA  from biological samples. This procedure is complicated by the ubiquitous presence of  ribonuclease  enzymes in cells and tissues, which can rapidly degrade RNA. [ 1 ]  Several methods are used in  molecular biology  to isolate RNA from samples, the most common of these is  guanidinium thiocyanate-phenol-chloroform extraction . [ 2 ] [ 3 ]  The filter paper based lysis and elution method features high throughput capacity. [ 4 ]


RNA extraction in liquid nitrogen, commonly using a mortar and pestle (or specialized steel devices known as tissue pulverizers) is also useful in preventing ribonuclease activity.


The extraction of RNA in molecular biology experiments is greatly complicated by the presence of ubiquitous and hardy RNases that degrade RNA samples. Certain RNases can be extremely hardy and inactivating them is difficult compared to neutralizing  DNases . In addition to the cellular RNases that are released there are several RNases that are present in the environment. RNases have evolved to have many extracellular functions in various organisms. [ 5 ] [ 6 ] [ 7 ]  For example, RNase 7, a member of the  RNase A  superfamily, is secreted by human skin and serves as a potent antipathogen defence. [ 8 ] [ 9 ]  For these secreted RNases, enzymatic activity may not even be necessary for the RNase's  exapted  function. For example, immune RNases act by destabilizing the cell membranes of bacteria. [ 10 ] [ 11 ]


To counter this, equipment used for RNA extraction is usually cleaned thoroughly, kept separate from common lab equipment and treated with various harsh chemicals that destroy RNases. For the same reason, experimenters take special care not to let their bare skin touch the equipment.


Two-phase wash to solve the ubiquitous contaminant-carryover problem in commercial nucleic-acid extraction kits ; by Erik Jue, Daan Witters & Rustem F. Ismagilov; Nature, Scientific reports, 2020.


RIP-chip (RNA immunoprecipitation chip)  is a  molecular biology  technique which combines RNA  immunoprecipitation  with a  microarray . The purpose of this technique is to identify which  RNA  sequences interact with a particular  RNA binding protein  of interest  in vivo . [ 1 ] [ 2 ] [ 3 ] [ 4 ]  It can also be used to determine relative levels of  gene expression , to identify subsets of RNAs which may be co-regulated, or to identify RNAs that may have related functions. [ 4 ] [ 5 ]  This technique provides insight into the post-transcriptional gene regulation which occurs between RNA and RNA binding proteins. [ 5 ]


The genes fluorescently identified by the chip analysis are the genes whose RNA interacts with the original protein of interest. The strength of the fluorescent signal for a particular gene can indicate how much of that particular RNA was present in the original sample, which indicates the expression level of that gene.


Previous techniques aiming to understand protein-RNA interactions included RNA  Electrophoretic Mobility Shift Assays  and UV-crosslinking followed by RT-PCR, [ 9 ]  however such selective analysis cannot be used when the bound RNAs are not yet known. [ 1 ]  To resolve this, RIP-chip combines RNA immunoprecipitation to isolate RNA molecules interacting with specific proteins with a microarray which can elucidate the identity of the RNAs participating in this interaction. [ 5 ] [ 1 ]  Alternatives to RIP-chip include:


Saturation mutagenesis , or  site saturation mutagenesis (SSM) , or simply  site saturation , is a  random mutagenesis  technique used in  protein engineering , in which a single  codon  or set of codons is substituted with all possible  amino acids  at the position. [ 1 ]  There are many variants of the site saturation technique, from  paired site saturation  (saturating two positions in every mutant in the library) to  scanning site saturation  (performing a site saturation at every site in the protein, resulting in a library of size [20^(number of residues in the protein)] that contains every possible  point mutant  of the protein).


Saturation mutagenesis is commonly achieved by  site-directed mutagenesis  PCR with a randomised codon in the primers (e.g.  SeSaM ) [ 2 ]  or by  artificial gene synthesis , with a mixture of synthesis  nucleotides  used at the codons to be randomised. [ 3 ]


Different degenerate codons can be used to encode sets of amino acids. [ 1 ]  Because some amino acids are encoded by more codons than others, the exact ratio of amino acids cannot be equal. Additionally, it is usual to use degenerate codons that minimise stop codons (which are generally not desired). Consequently, the fully randomised 'NNN' is not ideal, and alternative, more restricted degenerate codons are used. 'NNK' and 'NNS' have the benefit of encoding all 20 amino acids, but still encode a stop codon 3% of the time. Alternative codons such as 'NDT', 'DBK' avoid stop codons entirely, and encode a minimal set of amino acids that still encompass all the main biophysical types (anionic, cationic, aliphatic hydrophobic, aromatic hydrophobic, hydrophilic, small). [ 1 ]  In the case there is no restriction to use a single degenerate codon only, it is possible to reduce the bias considerably. [ 4 ] [ 5 ]  Several computational tools were developed to allow high level of control over the degenerate codons and their corresponding amino acids. [ 6 ] [ 7 ] [ 8 ]


Saturation mutagenesis is commonly used to generate variants for  directed evolution . [ 9 ] [ 10 ]


A  selectable marker  is a  gene  introduced into  cells , especially  bacteria  or cells in  culture , which confers one or more traits suitable for  artificial selection . They are a type of  reporter gene  used in laboratory  microbiology ,  molecular biology , and  genetic engineering  to indicate the success of a  transfection  or  transformation  or other procedure meant to introduce foreign  DNA  into a cell. Selectable markers are often  antibiotic resistance  genes: bacteria subjected to a procedure by which exogenous DNA containing an antibiotic resistance gene (usually alongside other  genes of interest ) has been introduced are grown on a medium containing an  antibiotic , such that only those bacterial cells which have successfully taken up and  expressed  the introduced genetic material, including the gene which confers antibiotic resistance, can survive and produce  colonies . The genes encoding resistance to antibiotics such as  ampicillin ,  chloramphenicol ,  tetracycline ,  kanamycin , etc., are all widely used as selectable markers for  molecular cloning  and other genetic engineering techniques in  E. coli .


Selectable markers allow scientists to separate non-recombinant organisms (those which do not contain the selectable marker) from recombinant organisms (those which do); that is, a  recombinant DNA  molecule such as a  plasmid   expression vector  is introduced into  bacterial  cells, and some bacteria are successfully transformed while some remain non-transformed. Antibiotics such as  ampicillin , at sufficient concentrations, are toxic to most bacteria, which ordinarily lack resistance to them; when cultured on a  nutrient medium  containing ampicillin, bacteria lacking ampicillin resistance fail to divide and eventually die. The position is later noted on nitrocellulose paper and separated out to move them to a nutrient medium for mass production of the required product. An alternative to a selectable marker is a screenable marker, another type of reporter gene which allows the researcher to distinguish between wanted and unwanted cells or colonies, such as between blue and white colonies in  blue–white screening . These wanted or unwanted cells are simply non-transformed cells that were unable to take up the screenable gene during the experiment. [ citation needed ]


For molecular biology research, different types of markers may be used based on the selection sought. These include:


Examples of selectable markers include:


In the future, alternative marker technologies will need to be used more often to, at the least, assuage concerns about their persistence into the final product. It is also possible that markers will be replaced entirely by future techniques which use removable markers, and others which do not use markers at all, instead relying on  co-transformation ,  homologous recombination , and  recombinase-mediated excision . [ 6 ]


In  pathology ,  silver staining  is the use of  silver  to selectively alter the appearance of a target in  microscopy  of  histological sections ; in  temperature gradient gel electrophoresis ; and in  polyacrylamide gels .


In traditional  stained glass ,  silver stain  is a technique to produce yellow to orange or brown shades (or green on a blue glass base), by adding a mixture containing silver compounds (notably  silver nitrate ), and firing lightly. It was introduced soon after 1800, and is the "stain" in the term "stained glass". Silver compounds [ 1 ]  are mixed with binding substances, applied to the surface of glass, and then fired in a furnace or kiln. [ 2 ] [ 3 ] [ 4 ]


Camillo Golgi  perfected silver staining for the study of the  nervous system . Although the exact chemical mechanism by which this occurs is unknown, [ 5 ]   Golgi's method  stains a limited number of cells at random in their entirety. [ 6 ]


Silver staining was introduced by Kerenyi and Gallyas as a sensitive procedure to detect trace amounts of proteins in  gels . [ 7 ]  The technique has been extended to the study of other biological  macromolecules  that have been separated in a variety of supports. [ 8 ]


Classical  Coomassie brilliant blue  staining can usually detect a 50 ng protein band; silver staining increases the sensitivity typically 50 times.


Many variables can influence the color intensity and every protein has its own staining characteristics; clean glassware, pure reagents, and water of highest purity are the key points to successful staining. [ 9 ]


Some cells are  argentaffin . These  reduce  silver solution to metallic silver after  formalin   fixation . Other cells are  argyrophilic . These reduce silver solution to metallic silver after being exposed to the stain that contains a  reductant , for example  hydroquinone  or formalin.


Silver nitrate  forms insoluble  silver phosphate  with  phosphate  ions; this method is known as the  Von Kossa Stain . When subjected to a reducing agent, usually  hydroquinone , it forms black elementary silver. This is used for study of formation of  calcium phosphate  particles during bone growth.


Silver staining aids the visualization of targets of interest, namely intracellular and extracellular cellular components such as  DNA  and  proteins , such as type III  collagen  and  reticulin  fibres by the deposition of metallic silver particles on the targets of interest. [ 10 ]


Pseudomonas , [ 11 ]   Legionella ,  Leptospira ,  H. pylori ,  Bartonella  and  Treponema , and fungi such as  Pneumocystis ,  Cryptococcus , and  Candida  are organisms that are stained with silver. [ citation needed ]


Silver staining is used in  karyotyping . Silver nitrate stains the  nucleolar organization region  (NOR)-associated protein, producing a dark region wherein the silver is deposited and denoting the activity of  rRNA  genes within the NOR. Human  chromosomes  13, 14, 15, 21, and 22 have NORs, which increase the silver stain activity by at least 50 times. [ citation needed ]


Silver staining is used to stain gels. The silver stain of proteins in  Agarose gels  was developed in 1973 by Kerenyi and Gallyas. [ 12 ]  Later it was adapted to  polyacrylamide gels  used in  SDS-PAGE , [ 13 ] [ 14 ] [ 15 ] [ 16 ] [ 17 ]  and also for staining DNA or RNA. [ 18 ]  The  glycosylations  of  glycoproteins  and  polysaccharides  can be oxidised by a 1-hour pre-treatment with 0.1%  periodic acid  at 4 °C, which improves the binding of silver ions and the staining result. [ 19 ]


First, the proteins are denatured in the gel by a fixative solution of 10%  acetic acid  and 30%  ethanol  and precipitated, at the same time the  detergent  (mostly  SDS ) is extracted. The  diffusion  of the proteins is thus significantly reduced. After repeated washing with water, the gel is incubated in a  silver nitrate  solution. Silver ions bind to negatively charged side chains of the proteins. Excess silver ions are then washed off with water. In the final development step, the silver ions are reduced to elemental silver by addition of alkaline  formaldehyde . This stains the sites where proteins are present, brown to black.


The intensity of the staining depends on the  primary structure  of the protein. Furthermore, the cleanliness of the vessels used and the purity of the  reagents  influence the silver stain. [ 20 ]  Common artifacts in silver stained gels are bands of  keratin  in the ranges of 54–57 kDa and 65–68 kDa [ 21 ]  as a contamination of the sample prior to the electrophoresis.


There are several silver stains incorporating  methenamine , including:


Single-base extension (SBE)  is a method for determining the identity of a  nucleotide  base at a specific position along a  nucleic acid . The method is used to identify a  single-nucleotide polymorphism  (SNP).


In the method, an  oligonucleotide  primer hybridizes to a complementary region along the nucleic acid to form a duplex, with the primer’s terminal 3’-end directly adjacent to the nucleotide base to be identified. Using a DNA polymerase, the oligonucleotide primer is enzymatically extended by a single base in the presence of all four nucleotide terminators; the nucleotide terminator complementary to the base in the template being interrogated is incorporated and identified. The presence of all four terminators suppresses misincorporation of non-complementary nucleotides. Many approaches can be taken for determining the identity of an incorporated terminator, including fluorescence labeling, mass labeling for mass spectrometry, isotope labeling, and tagging the base with a hapten and detecting chromogenically with an anti-hapten antibody-enzyme conjugate (e.g., via an ELISA format).


The method was invented by Philip Goelet, Michael Knapp, Richard Douglas and Stephen Anderson while working at the company Molecular Tool. This approach was designed for high-throughput SNP genotyping and was originally called "Genetic Bit Analysis" (GBA).  Illumina, Inc. utilizes this method in their Infinium technology ( http://www.illumina.com/technology/beadarray-technology/infinium-hd-assay.html ) to measure  DNA methylation  levels in the human genome.


Site-directed mutagenesis  is a  molecular biology  method that is used to make specific and intentional mutating changes to the  DNA sequence  of a  gene  and any  gene products . Also called  site-specific mutagenesis  or  oligonucleotide-directed mutagenesis , it is used for investigating the structure and biological activity of  DNA ,  RNA , and  protein  molecules, and for  protein engineering .


Site-directed mutagenesis is one of the most important laboratory techniques for creating  DNA libraries  by introducing mutations into DNA sequences. There are numerous methods for achieving site-directed mutagenesis, but with decreasing costs of  oligonucleotide synthesis ,  artificial gene synthesis  is now occasionally used as an alternative to site-directed mutagenesis. Since 2013, the development of the  CRISPR /Cas9 technology, based on a prokaryotic viral defense system, has also allowed for the  editing of the genome , and mutagenesis may be performed  in vivo  with relative ease. [ 1 ]


Early attempts at  mutagenesis  using radiation or chemical mutagens were non-site-specific, generating random mutations. [ 2 ]   Analogs of nucleotides and other chemicals were later used to generate localized  point mutations , [ 3 ]  examples of such chemicals are  aminopurine , [ 4 ]   nitrosoguanidine , [ 5 ]  and  bisulfite . [ 6 ]   Site-directed mutagenesis was achieved in 1974 in the laboratory of  Charles Weissmann  using a nucleotide analogue N 4 -hydroxycytidine, which induces  transition  of GC to AT. [ 7 ] [ 8 ]  These methods of mutagenesis, however, are limited by the kind of mutation they can achieve, and they are not as specific as later site-directed mutagenesis methods.


In 1971,  Clyde Hutchison  and Marshall Edgell showed that it is possible to produce mutants with small fragments of  phage ϕX174  and  restriction nucleases . [ 9 ] [ 10 ]  Hutchison later produced with his collaborator  Michael Smith  in 1978 a more flexible approach to site-directed mutagenesis by using  oligonucleotides  in a primer extension method with DNA polymerase. [ 11 ]   For his part in the development of this process, Michael Smith later shared the  Nobel Prize in Chemistry  in October 1993 with  Kary B. Mullis , who invented  polymerase chain reaction .


The basic procedure requires the  synthesis  of a short DNA primer.  This synthetic primer contains the desired mutation and is complementary to the template DNA around the mutation site so it can  hybridize  with the DNA in the gene of interest.  The mutation may be a single base change (a  point mutation ), multiple base changes,  deletion , or  insertion .  The single-strand primer is then extended using a  DNA polymerase , which copies the rest of the gene. The gene thus copied contains the mutated site, and is then introduced into a host cell in a vector and  cloned . Finally, mutants are selected by  DNA sequencing  to check that they contain the desired mutation.


The original method using single-primer extension was inefficient due to a low yield of mutants.  This resulting mixture contains both the original unmutated template as well as the mutant strand, producing a mixed population of mutant and non-mutant progenies. Furthermore, the template used is  methylated  while the mutant strand is unmethylated, and the mutants may be counter-selected due to presence of  mismatch repair  system that favors the methylated template DNA, resulting in fewer mutants.  Many approaches have since been developed to improve the efficiency of mutagenesis.


A large number of methods are available to effect site-directed mutagenesis, [ 12 ]  although most of them have rarely been used in laboratories since the early 2000s, as newer techniques allow for simpler and easier ways of introducing site-specific mutation into genes.


In 1985,  Thomas Kunkel  introduced a technique that reduces the need to select for the mutants. [ 13 ]  The DNA fragment to be mutated is inserted into a  phagemid  such as  M13mp18/19  and is then transformed into an  E. coli  strain deficient in two enzymes,  dUTPase  ( dut ) and  uracil deglycosidase  ( udg ). Both enzymes are part of a  DNA repair  pathway that protects the bacterial chromosome from mutations by the spontaneous deamination of dCTP to dUTP. The dUTPase deficiency prevents the breakdown of dUTP, resulting in a high level of dUTP in the cell. The uracil deglycosidase deficiency prevents the removal of uracil from newly synthesized DNA. As the double-mutant  E. coli  replicates the phage DNA, its enzymatic machinery may, therefore, misincorporate dUTP instead of dTTP, resulting in single-strand DNA that contains some uracils (ssUDNA). The ssUDNA is  extracted  from the bacteriophage that is released into the medium, and then used as template for mutagenesis.  An  oligonucleotide  containing the desired mutation is used for primer extension. The heteroduplex DNA, that forms, consists of one parental non-mutated strand containing dUTP and a mutated strand containing dTTP.  The DNA is then transformed into an  E. coli  strain carrying the wildtype  dut  and  udg  genes. Here, the uracil-containing parental DNA strand is degraded, so that nearly all of the resulting DNA consists of the mutated strand.


Unlike other methods,  cassette mutagenesis  need not involve primer extension using DNA polymerase.  In this method, a fragment of DNA is synthesized, and then inserted into a plasmid. [ 14 ]  It involves the cleavage by a  restriction enzyme  at a site in the plasmid and subsequent  ligation  of a pair of complementary oligonucleotides containing the mutation in the gene of interest to the plasmid. Usually, the restriction enzymes that cut at the plasmid and the oligonucleotide are the same, permitting sticky ends of the plasmid and insert to ligate to one another.  This method can generate mutants at close to 100% efficiency, but is limited by the availability of suitable restriction sites flanking the site that is to be mutated.


The limitation of restriction sites in cassette mutagenesis may be overcome using  polymerase chain reaction  with  oligonucleotide  " primers ", such that a larger fragment may be generated, covering two convenient restriction sites.  The exponential amplification in PCR produces a fragment containing the desired mutation in sufficient quantity to be separated from the original, unmutated plasmid by  gel electrophoresis , which may then be inserted in the original context using standard recombinant molecular biology techniques.  There are many variations of the same technique.  The simplest method places the mutation site toward one of the ends of the fragment whereby one of two oligonucleotides used for generating the fragment contains the mutation.  This involves a single step of PCR, but still has the inherent problem of requiring a suitable restriction site near the mutation site unless a very long primer is used.  Other variations, therefore, employ three or four oligonucleotides, two of which may be non-mutagenic oligonucleotides that cover two convenient restriction sites and generate a fragment that can be digested and ligated into a plasmid, whereas the mutagenic oligonucleotide may be complementary to a location within that fragment well away from any convenient restriction site.  These methods require multiple steps of PCR so that the final fragment to be ligated can contain the desired mutation. The design process for generating a fragment with the desired mutation and relevant restriction sites can be cumbersome. Software tools like SDM-Assist [ 15 ]  can simplify the process.


For plasmid manipulations, other site-directed mutagenesis techniques have been supplanted largely by techniques that are highly efficient but relatively simple, easy to use, and commercially available as a kit.  An example of these techniques is the "Quikchange" method, [ 16 ]  wherein a pair of complementary mutagenic primers are used to amplify the entire plasmid in a  thermocycling  reaction using a high-fidelity non-strand-displacing DNA polymerase such as  Pfu  polymerase . The reaction generates a  nicked , circular DNA.   The template DNA must be eliminated by enzymatic digestion with a  restriction enzyme  such as  Dpn I , which is specific for methylated DNA. All DNA produced from most  Escherichia coli  strains would be methylated; the template plasmid that is biosynthesized in  E. coli  will, therefore, be digested, while the mutated plasmid, which is generated  in vitro  and is therefore unmethylated, would be left undigested. Note that, in these double-strand plasmid mutagenesis methods, while the thermocycling reaction may be used, the DNA is not exponentially amplified if the two primers are designed such that they bind symmetrically to the same region around the mutagenesis site, as described in the original protocol. In this case the amplification is linear, and it is therefore inaccurate to describe the procedure as a PCR, since there is no chain reaction. However, if the primers are designed to bind in an offset manner such that mutagenesis site is close to the 5' end of both primers, the 3' region of the primers can bind also to the amplified products and thus exponential product formation is observed. The name "Quikchange" originates from the registered trademark "QuikChange mutagenesis" of   Stratagene , now  Agilent Technologies  , for site directed mutagenesis kits. The method was developed by scientists working at Stratagene. [ 16 ]


Note that  Pfu  polymerase can become strand-displacing at higher extension temperature (≥70 °C) which can result in the failure of the experiment, therefore the extension reaction should be performed at the recommended temperature of 68 °C.  In some applications, this method has been observed to lead to insertion of multiple copies of primers. [ 17 ]  A variation of this method, called SPRINP, prevents this artifact and has been used in different types of site directed mutagenesis. [ 17 ]


Other techniques such as scanning mutagenesis of oligo-directed targets (SMOOT) can semi-randomly combine mutagenic oligonucleotides in plasmid mutagenesis. [ 18 ]  This technique can create plasmid mutagenesis libraries ranging from single mutations to comprehensive codon mutagenesis across an entire gene.


Since 2013, the development of  CRISPR -Cas9 technology has allowed for the efficient introduction of various mutations into the genome of a wide variety of organisms. The method does not require a transposon insertion site, leaves no marker, and its efficiency and simplicity has made it the preferred method for  genome editing . [ 21 ] [ 22 ]


Site-directed mutagenesis is used to generate mutations that may produce a  rationally designed  protein that has improved or special properties (i.e.protein engineering).


Investigative tools  – specific mutations in DNA allow the function and properties of a DNA sequence or a protein to be investigated in a rational approach. Furthermore, single amino-acid changes by site-directed mutagenesis in proteins can help understand the importance of post-translational modifications. For instance changing a particular serine (phosphoacceptor) to an alanine (phospho-non-acceptor) in a substrate protein blocks the attachment of a phosphate group, thereby allows the phosphorylation to be investigated. This approach has been used to uncover the phosphorylation of the protein  CBP  by the kinase  HIPK2 [ 23 ]  Another comprehensive approach is site  saturation mutagenesis  where one  codon  or a set of codons may be substituted with all possible  amino acids  at the specific positions. [ 24 ]


Commercial applications  – Proteins may be engineered to produce mutant forms that are tailored for a specific application.  For example, commonly used laundry detergents may contain  subtilisin , whose wild-type form has a methionine that can be oxidized by bleach, significantly reducing the activity the protein in the process. [ 25 ]  This methionine may be replaced by alanine or other residues, making it resistant to oxidation thereby keeping the protein active in the presence of bleach. [ 26 ]


As the cost of DNA oligonucleotides synthesis falls,  artificial synthesis of a complete gene  is now a viable method for introducing mutation into gene. This method allows for extensive mutagenesis over multiples sites, including the complete redesign of the codon usage of gene to optimise it for a particular organism. [ 27 ]


Site-specific recombination , also known as  conservative site-specific recombination , is a type of  genetic recombination  in which  DNA  strand exchange takes place between segments possessing at least a certain degree of  sequence homology . [ 1 ] [ 2 ] [ 3 ]   Enzymes  known as site-specific  recombinases  (SSRs) perform rearrangements of DNA segments by recognizing and binding to short, specific DNA sequences (sites), at which they cleave the DNA backbone, exchange the two DNA helices involved, and rejoin the DNA strands. In some cases the presence of a recombinase enzyme and the recombination sites is sufficient for the reaction to proceed; in other systems a number of accessory proteins and/or accessory sites are required. Many different  genome modification strategies , among these  recombinase-mediated cassette exchange  (RMCE), an advanced approach for the targeted introduction of transcription units into predetermined genomic loci, rely on SSRs.


Site-specific recombination systems are highly specific, fast, and efficient, even when faced with complex  eukaryotic  genomes. [ 4 ]  They are employed naturally in a variety of cellular processes, including bacterial genome  replication , differentiation and  pathogenesis , and movement of  mobile genetic elements . [ 5 ]  For the same reasons, they present a potential basis for the development of  genetic engineering  tools. [ 6 ]


Recombination sites are typically between 30 and 200  nucleotides  in length and consist of two motifs with a partial inverted-repeat symmetry, to which the recombinase binds, and which flank a central crossover sequence at which the recombination takes place. The pairs of sites between which the recombination occurs are usually identical, but there are exceptions (e.g.  attP and attB of λ integrase ). [ 7 ]


Based on amino acid sequence homologies and mechanistic relatedness, most site-specific recombinases are grouped into one of two families: the  tyrosine (Tyr) recombinase family  or  serine (Ser) recombinase  family. The names stem from the conserved nucleophilic amino acid residue present in each class of recombinase which is used to attack the DNA and which becomes covalently linked to it during strand exchange. The earliest identified members of the serine recombinase family were known as  resolvases  or  DNA invertases , while the founding member of the tyrosine recombinases,  lambda phage  integrase (using attP/B recognition sites), differs from the now well-known enzymes such as  Cre  (from the  P1 phage ) and  FLP  (from the yeast  Saccharomyces cerevisiae ). Famous serine recombinases include enzymes such as  gamma-delta resolvase  (from the Tn 1000   transposon ),  Tn3 resolvase  (from the Tn3 transposon), and  φ C31 integrase  (from the  φ C31 phage). [ 8 ]  There are several classes of serine recombinases, consisting of the small serine recombinase, the ISXc5 resolvase, the serine transposase, and the large serine recombinase. [ 9 ]


Although the individual members of the two recombinase families can perform reactions with the same practical outcomes, the families are unrelated to each other, having different protein structures and reaction mechanisms. Unlike tyrosine recombinases, serine recombinases are highly modular, as was first hinted by biochemical studies [ 10 ]  and later shown by crystallographic structures. [ 11 ] [ 12 ]  Knowledge of these protein structures could prove useful when attempting to re-engineer recombinase proteins as tools for genetic manipulation.


Recombination between two DNA sites begins by the recognition and binding of these sites – one site on each of two separate double-stranded DNA molecules, or at least two distant segments of the same molecule – by the recombinase enzyme. This is followed by  synapsis , i.e. bringing the sites together to form the synaptic complex. It is within this synaptic complex that the strand exchange takes place, as the DNA is cleaved and rejoined by controlled  transesterification  reactions. During strand exchange, each double-stranded DNA molecule is cut at a fixed point within the crossover region of the recognition site, releasing a  deoxyribose   hydroxyl group , while the recombinase enzyme forms a transient  covalent bond  to a DNA backbone  phosphate . This  phosphodiester bond  between the hydroxyl group of the  nucleophilic   serine  or  tyrosine  residue conserves the energy that was expended in cleaving the DNA. Energy stored in this bond is subsequently used for the rejoining of the DNA to the corresponding deoxyribose hydroxyl group on the other DNA molecule. The entire reaction therefore proceeds without the need for external energy-rich  cofactors  such as  ATP .


Although the basic chemical reaction is the same for both tyrosine and serine recombinases, there are some differences between them. [ 13 ]  Tyrosine recombinases, such as  Cre  or  FLP , cleave one DNA strand at a time at points that are staggered by 6–8bp, linking the 3' end of the strand to the hydroxyl group of the tyrosine nucleophile (Fig. 1). [ 14 ]  Strand exchange then proceeds via a crossed strand intermediate analogous to the  Holliday junction  in which only one pair of strands has been exchanged. [ 15 ] [ 16 ]


The mechanism and control of serine recombinases is much less well understood. This group of enzymes was only discovered in the mid-1990s and is still relatively small. The now classical members  gamma-delta  and  Tn3 resolvase , but also new additions like φC31-, Bxb1-, and R4 integrases, cut all four DNA strands simultaneously at points that are staggered by 2 bp (Fig. 2). [ 17 ]  During cleavage, a protein–DNA bond is formed via a  transesterification  reaction, in which a  phosphodiester bond  is replaced by a phosphoserine bond between a 5' phosphate at the cleavage site and the hydroxyl group of the conserved serine residue (S10 in resolvase). [ 18 ] [ 19 ]


It is still not entirely clear how the strand exchange occurs after the DNA has been cleaved. However, it has been shown that the strands are exchanged while covalently linked to the protein, with a resulting net rotation of 180°. [ 20 ] [ 21 ]  The most quoted (but not the only) model accounting for these facts is the "subunit rotation model" (Fig. 2). [ 13 ] [ 22 ]  Independent of the model, DNA duplexes are situated outside of the protein complex, and large movement of the protein is needed to achieve the strand exchange. In this case the recombination sites are slightly asymmetric, which allows the enzyme to tell apart the left and right ends of the site. When generating products, left ends are always joined to the right ends of their partner sites, and vice versa. This causes different recombination hybrid sites to be reconstituted in the recombination products. Joining of left ends to left or right to right is avoided due to the asymmetric "overlap" sequence between the staggered points of top and bottom strand exchange, which is in stark contrast to the mechanism employed by tyrosine recombinases. [ 13 ]


The reaction catalysed by Cre-recombinase, for instance, may lead to excision of the DNA segment flanked by the two sites (Fig. 3A), but may also lead to integration or inversion of the orientation of the flanked DNA segment (Fig. 3B). What the outcome of the reaction will be is dictated mainly by the relative locations and orientations of the sites that are to be recombined, but also by the innate specificity of the site-specific system in question. Excisions and inversions occur if the recombination takes place between two sites that are found on the same molecule (intramolecular recombination), and if the sites are in the same (direct repeat) or in an opposite orientation (inverted repeat), respectively. Insertions, on the other hand, take place if the recombination occurs on sites that are situated on two different DNA molecules (intermolecular recombination), provided that at least one of these molecules is circular. Most site-specific systems are highly specialised, catalysing only one of these different types of reaction, and have evolved to ignore the sites that are in the "wrong" orientation.


SNV calling from NGS data  is any of a range of methods for identifying the existence of  single nucleotide variants  (SNVs) from the results of  next generation sequencing  (NGS) experiments. These are computational techniques, and are in contrast to special experimental methods based on known population-wide single nucleotide polymorphisms (see  SNP genotyping ). Due to the increasing abundance of NGS data, these techniques are becoming increasingly popular for performing SNP genotyping, with a wide variety of algorithms designed for specific experimental designs and applications. [ 1 ]
In addition to the usual application domain of SNP genotyping, these techniques have been successfully adapted to identify rare SNPs within a population, [ 2 ]
as well as detecting  somatic  SNVs within an individual using multiple tissue samples. [ 3 ]


Most NGS based methods for SNV detection are designed to detect  germline  variations in the individual's genome. These are the mutations that an individual biologically inherits from their parents, and are the usual type of variants searched for when performing such analysis (except for certain specific applications where  somatic mutations  are sought). Very often, the searched for variants occur with some (possibly rare) frequency, throughout the population, in which case they may be referred to as  single nucleotide polymorphisms  (SNPs). Technically the term SNP only refers to these kinds of variations, however in practice they are often used synonymously with SNV in the literature on variant calling. In addition, since the detection of germline SNVs requires determining the individual's genotype at each locus, the phrase "SNP genotyping" may also be used to refer to this process. However this phrase may also refer to wet-lab experimental procedures for classifying genotypes at a set of known SNP locations.


The usual process of such techniques are based around: [ 1 ]


The usual output of these procedures is a  VCF  file.


In an ideal error free world with high read  coverage , the task of variant calling from the results of a NGS data alignment would be simple; at each  locus  (position on the genome) the number of occurrences of each distinct nucleotide among the reads aligned at that position can be counted, and the true genotype would be obvious; either  AA  if all nucleotides match allele  A ,  BB  if they match allele  B , or  AB  if there is a mixture. However, when working with real NGS data this sort of naive approach is not used, as it cannot account for the noise in the input data. [ 4 ]
The nucleotide counts used for base calling contain errors and bias, both due do the sequenced reads themselves, and the alignment process. This issue can be mitigated to some extent by sequencing to a greater depth of read coverage, however this is often expensive, and many practical studies require making inferences on low coverage data. [ 1 ]


Probabilistic methods aim to overcome the above issue, by producing robust estimates of the probabilities of each of the possible genotypes, taking into account noise, as well as other available prior information that can be used to improve estimates. A genotype can then be predicted based on these probabilities, often according to the  MAP  estimate.


Probabilistic methods for variant calling are based on  Bayes' theorem . In the context of variant calling, Bayes' Theorem defines the probability of each genotype being the true genotype given the observed data, in terms of the prior probabilities of each possible genotype, and the probability distribution of the data given each possible genotype. The formula is:


In the above equation:


Given the above framework, different software solutions for detecting SNVs vary based on how they calculate the prior probabilities

 P
 (
 G
 )

 {\displaystyle P(G)}

 , the error model used to model the probabilities

 P
 (
 D
 ∣
 G
 )

 {\displaystyle P(D\mid G)}

 , and the partitioning of the overall genotypes into separate sub-genotypes, whose probabilities can be individually estimated in this framework. [ 5 ]


The calculation of prior probabilities depends on available data from the genome being studied, and the type of analysis being performed. For studies where good reference data containing frequencies of known mutations is available (for example, in studying human genome data), these known frequencies of genotypes in the population can be used to estimate priors. Given population wide allele frequencies, prior genotype probabilities can be calculated at each locus according to the  Hardy–Weinberg equilibrium . [ 6 ]
In the absence of such data, constant priors can be used, independent of the locus. These can be set using heuristically chosen values, possibly informed by the kind of variations being sought by the study. Alternatively, supervised machine-learning procedures have been investigated that seek to learn optimal prior values for individuals in a sample, using supplied NGS data from these individuals. [ 4 ]


The error model used in creating a probabilistic method for variant calling is the basis for calculating the

 P
 (
 D
 ∣
 G
 )

 {\displaystyle P(D\mid G)}

  term used in Bayes' theorem. If the data was assumed to be error free, then the distribution of observed nucleotide counts at each locus would follow a  Binomial Distribution , with 100% of nucleotides matching the A or B allele respectively in the  AA  and  BB  cases, and a 50% chance of each nucleotide matching either  A  or  B  in the  AB  case. However, in presence of noise in the read data this assumption is violated, and the

 P
 (
 D
 ∣
 G
 )

 {\displaystyle P(D\mid G)}

  values need to account for the possibility that erroneous nucleotides are present in the aligned reads at each locus.


A simple error model is to introduce a small error to the data probability term in the homozygous cases, allowing a small constant probability that nucleotides which don't match the  A  allele are observed in the  AA  case, and respectively a small constant probability that nucleotides not matching the  B  allele are observed in the  BB  case. However more sophisticated procedures are available which attempt to more realistically replicate the actual error patterns observed in real data in calculating the conditional data probabilities. For instance, estimations of read quality (measured as  Phred  quality scores) have been incorporated in these calculations, taking into account the expected error rate in each individual read at a locus. [ 7 ]
Another technique that has successfully been incorporated into error models is base quality recalibration, where separate error rates are calculated – based on prior known information about error patterns – for each possible nucleotide substitution. Research shows that each possible nucleotide substitution is not equally likely to show up as an error in sequencing data, and so base quality recalibration has been applied to improve error probability estimates. [ 6 ]


In the above discussion, it has been assumed that the genotype probabilities at each locus are calculated independently; that is, the entire genotype is partitioned into independent genotypes at each locus, whose probabilities are calculated independently. However, due to  linkage disequilibrium  the genotypes of nearby loci are in general not independent. As a result, partitioning the overall genotype instead into a sequence of overlapping  haplotypes  allows these correlations to be modelled, resulting in more precise probability estimates through the incorporation of population-wide haplotype frequencies in the prior. The use of haplotypes to improve variant detection accuracy has been applied successfully, for instance in the  1000 Genomes Project . [ 8 ]


As an alternative to probabilistic methods,  heuristic  methods exist for performing variant calling on NGS data. Instead of modelling the distribution of the observed data and using Bayesian statistics to calculate genotype probabilities, variant calls are made based on a variety of heuristic factors, such as minimum allele counts, read quality cut-offs, bounds on read depth, etc. Although they have been relatively unpopular in practice in comparison to probabilistic methods, in practice due to their use of bounds and cut-offs they can be robust to outlying data that violate the assumptions of probabilistic models. [ 9 ]


An important part of the design of variant calling methods using NGS data is the DNA sequence used as a reference to which the NGS reads are aligned. In human genetics studies, high quality references are available, from sources such as the  HapMap project , [ 10 ]
which can substantially improve the accuracy of the variant calls made by variant calling algorithms. As a bonus, such references can be a source of prior genotype probabilities for Bayesian-based analysis. However, in the absence of such a high quality reference, experimentally obtained reads can first be  assembled  in order to create a reference sequence for alignment. [ 1 ]


Various methods exist for filtering data in variant calling experiments, in order to remove sources of error/bias. This can involve the removal of suspicious reads before performing alignment and/or filtering of the list of variants returned by the variant calling algorithm.


Depending on the sequencing platform used, various biases may exist within the set of sequenced reads. For instance, strand bias can occur, where there is a highly unequal distribution of forward vs reverse directions in the reads aligned in some neighborhood. Additionally, there may occur an unusually high duplication of some reads (for instance due to bias in  PCR ). Such biases can result in dubious variant calls – for instance if a fragment containing a PCR error at some locus is over amplified due to PCR bias, that locus will have a high count of the false allele, and may be called as a SNV – and so analysis pipelines frequently filter calls based on these biases. [ 1 ]


In addition to methods that align reads from individual sample(s) to a reference genome in order to detect  germline  genetic variants, reads from multiple tissue samples within a single individual can be aligned and compared in order to detect somatic variants. These variants correspond to  mutations  that have occurred  de novo  within groups of  somatic cells  within an individual (that is, they are not present within the individual's germline cells). This form of analysis has been frequently applied to the study of  cancer , where many studies are designed around investigating the profile of somatic mutations within cancerous tissues. Such investigations have resulted in diagnostic tools that have seen clinical application, and are used to improve scientific understanding of the disease, for instance by the discovery of new cancer-related genes, identification of involved  gene regulatory networks  and  metabolic pathways , and by informing models of how tumors grow and evolve. [ 11 ]


Until recently, software tools for carrying out this form of analysis have been heavily underdeveloped, and were based on the same algorithms used to detect germline variations. Such procedures are not optimized for this task, because they do not adequately model the statistical correlation between the genotypes present in multiple tissue samples from the same individual. [ 3 ]


More recent investigations have resulted in the development of software tools especially optimized for the detection of somatic mutations from multiple tissue samples. Probabilistic techniques have been developed that pool allele counts from all tissue samples at each locus, and using statistical models for the likelihoods of joint-genotypes for all the tissues, and the distribution of allele counts given the genotype, are able to calculate relatively robust probabilities of somatic mutations at each locus using all available data. [ 3 ] [ 12 ]
In addition there has recently been some investigation in  machine learning  based techniques for performing this analysis. [ 13 ] [ 14 ] [ 15 ] [ 16 ]


In 2021, the Sequencing Quality Control Phase 2 Consortium [ 17 ]  has published a number of studies that investigated the effects of sample preparations, sequencing library kits, sequencing platforms, and bioinformatics workflows on the accuracy of somatic SNV detection [ 18 ]
based on a pair of tumor-normal cell lines that the Consortium has established as the reference samples, data, and call sets. [ 19 ]


DNA ends  refer to the properties of the ends of linear  DNA  molecules, which in  molecular biology  are described as "sticky" or "blunt" based on the shape of the complementary strands at the terminus. In  sticky ends , one strand is longer than the other (typically by at least a few nucleotides), such that the longer strand has bases which are left unpaired. In  blunt ends , both strands are of equal length – i.e. they end at the same base position, leaving no unpaired bases on either strand.


The concept is used in  molecular biology , in  cloning , or when subcloning  insert DNA  into  vector DNA . Such ends may be generated by  restriction enzymes  that break the molecule's phosphodiester backbone at specific locations, which themselves belong to a larger class of enzymes called  exonucleases  and  endonucleases . A restriction enzyme that cuts the backbones of both strands at non-adjacent locations leaves a staggered cut, generating two overlapping sticky ends, while an enzyme that makes a straight cut (at locations directly across from each other on both strands) generates two blunt ends. [ 1 ]


A single-stranded non-circular DNA molecule has two non-identical ends,  the 3' end and the 5' end  (usually pronounced "three prime end" and "five prime end").  The numbers refer to the numbering of carbon atoms in the  deoxyribose , which is a sugar forming an important part of the backbone of the DNA molecule. In the backbone of DNA the 5' carbon of one deoxyribose is linked to the 3' carbon of another by a phosphodiester bond linkage. [ 2 ]


When a molecule of DNA is double stranded, as DNA usually is, the two strands run in opposite directions. Therefore, one end of the molecule will have the 3' end of strand 1 and the 5' end of strand 2, and vice versa in the other end. [ 2 ]  However, the fact that the molecule is two stranded allows numerous different variations.


The simplest DNA end of a double stranded molecule is called a  blunt end . Blunt ends are also known as non-cohesive ends. In a blunt-ended molecule, both strands terminate in a  base pair . Blunt ends are not always desired in biotechnology since when using a  DNA ligase  to join two molecules into one, the yield is significantly lower with blunt ends. [ 3 ]  When performing subcloning, it also has the disadvantage of potentially inserting the insert DNA in the opposite orientation desired. On the other hand, blunt ends are always compatible with each other. Here is an example of a small piece of blunt-ended DNA:


Non-blunt ends are created by various  overhangs . An overhang is a stretch of unpaired  nucleotides  in the end of a DNA molecule. These unpaired nucleotides can be in either strand, creating either 3' or 5' overhangs. [ 3 ]  These overhangs are in most cases palindromic.