A comparative NAIL-MS experiment is quite similar to a  SILAC  experiment but for RNA instead of proteins. First, two populations of the respective cells are cultivated. One of the cell populations is fed with growth medium containing unlabeled nutrients, whereas the second population is fed with growth medium containing stable isotope labeled nutrients. The cells then incorporate the respective isotopologues into their RNA molecules. One of the cell populations serves as a control group whereas the other is subject to the associated research (e.g. KO strain, stress). Upon harvesting of the two cell populations they are mixed and co-processed together to exclude purification-bias. Due to the distinct masses of incorporated nutrients into the nucleosides a differentiation of the two cell populations is possible by mass spectrometry.


Upon initiation of a pulse-chase experiment the medium is switched from medium(1) to medium(2). The two media must only differ in their isotope content. Thereby it is possible to distinguish between RNA molecules already existent before experiment initiation (= RNA molecules grown in medium(1)) and RNA molecules that are newly transcribed after experiment initiation (= RNA molecules grown in medium(2)). This allows the detailed study of modification dynamics  in vivo . The supplementation of labeled methionine in either medium(1) or medium(2) allows the tracing of methylation processes. Other isotopically labeled metabolites potentially allow for further modification analysis.


Altogether NAIL-MS enables the investigation of RNA modification dynamics by mass spectrometry. With this technique, enzymatic demethylation has been observed for several RNA damages inside living bacteria. [ 4 ] [ 7 ]


For the discovery of uncharacterized modifications cells are grown in unlabeled  or   13 C‑labeled  or   15 N‑labeled  or   2 H‑labeled  or   34 S‑labeled medium. Unknown signals occurring during mass spectrometry are then inspected in all differentially labeled cultures. If retention times of unknown compounds with appropriately divergent  m/z  values overlap, a sum formula of the compound can be postulated by calculating the mass differences of the overlapping signal in the differentially labeled cultures. With this method several new RNA modifications could be discovered. This experimental design also was the initial idea that started the concept of NAIL-MS.


NAIL-MS can also be applied to oligonucleotide analysis by mass spectrometry. This is useful when the sequence information is to be retained. [ 11 ]


Nick translation [ 1 ]  (or  head translation ), developed in 1977 by Peter Rigby and Paul Berg, is a  tagging  technique in  molecular biology  in which  DNA polymerase I  is used to replace some of the nucleotides of a  DNA  sequence with their labeled analogues, creating a tagged DNA sequence which can be used as a probe in  fluorescent in situ hybridization  (FISH) or  blotting  techniques. It can also be used for  radiolabeling . [ 2 ]


This process is called "nick translation" because the DNA to be processed is treated with DNAase to produce single-stranded "nicks", where one of the strands is missing nucleotides. This is followed by replacement in nicked sites by  DNA polymerase I , which removes nucleotides from the 3' (downstream) end of a nick with its 3'-5' endonuclease activity and adds new, labeled  dNTPs  from the medium to the 5' end of the nick, moving the nick downstream in the process. [ 3 ]  To radioactively label a DNA fragment for use as a probe in blotting procedures, one of the incorporated nucleotides provided in the reaction is radiolabeled in the  alpha  phosphate position, often using  phosphorus-32 . Similarly, a fluorophore can be attached instead for fluorescent labelling, or an antigen for immunodetection. When DNA polymerase I eventually detaches from the DNA, it leaves another nick in the phosphate backbone.  The nick has "translated" some distance depending on the  processivity  of the polymerase. This nick could be sealed by  DNA ligase , or its 3' hydroxyl group could serve as the template for further DNA polymerase I activity. Proprietary enzyme mixes are available commercially to perform all steps in the procedure in a single incubation.


Nick translation could cause double-stranded DNA breaks, if DNA polymerase I encounters another nick on the opposite strand, resulting in two shorter fragments. This does not influence the performance of the labelled probe in in-situ hybridization.


Nicking Enzyme Amplification Reaction  (NEAR) is a method for  in vitro   DNA  amplification like the  polymerase chain reaction  (PCR).  NEAR is isothermal, replicating DNA at a constant temperature using a  polymerase  (and  nicking enzyme ) to exponentially amplify the DNA at a temperature range of 55 °C to 59 °C.


One disadvantage of PCR is that it consumes time uncoiling the double-stranded DNA with heat into single strands (a process called  denaturation ) . This leads to amplification times typically thirty minutes or more for significant production of amplified products. [ 1 ] [ circular reference ]


Potential advantages of NEAR over PCR are increased speed and lower energy requirements, characteristics that are shared with other isothermal amplification schemes. [ 2 ]  A major disadvantage of NEAR relative to PCR is that production of nonspecific amplification products is a common issue with isothermal amplification reactions. [ 3 ]


The NEAR reaction uses naturally occurring or engineered endonucleases that introduce a strand break on only one strand of a double-stranded DNA cleavage site.  The ability of several of these enzymes to catalyze isothermal DNA amplification was disclosed but not claimed in the patents issued for the enzymes themselves. [ 4 ] [ 5 ]


A  nuclear run-on assay  is conducted to identify the  genes  that are being  transcribed  at a certain time point. Approximately one million  cell nuclei  are isolated and incubated with labeled  nucleotides , and genes in the process of being transcribed are detected by hybridization of extracted RNA to gene specific probes on a  blot . [ 1 ]  Garcia-Martinez et al. (2004)  [ 2 ]  developed a protocol for the yeast  S. cerevisiae  (Genomic run-on, GRO) that allows for the calculation of transcription rates (TRs) for all yeast genes to estimate mRNA stabilities for all yeast mRNAs. [ 3 ]


Alternative microarray methods have recently been developed, mainly PolII RIP-chip:  RNA immunoprecipitation  of RNA polymerase II with phosphorylated C-terminal domain directed antibodies and hybridization on a microarray slide or chip (the word chip in the name stems from "ChIP-chip" where a special  Affymetrix  GeneChip was required). A comparison of methods based on run-on and ChIP-chip has been made in yeast (Pelechano et al., 2009). A general correspondence of both methods has been detected but GRO is more sensitive and quantitative. It has to be considered that run-on only detects elongating RNA polymerases whereas ChIP-chip detects all present RNA polymerases, including backtracked ones.


Attachment of new RNA polymerase to genes is prevented by inclusion of  sarkosyl . Therefore only genes that already have an RNA polymerase will produce labeled transcripts. RNA transcripts that were synthesized before the addition of the label will not be detected as they will lack the label. These run on transcripts can also be detected by purifying labeled transcripts by using antibodies that detect the label and hybridizing these isolated transcripts with gene expression arrays or by next generation sequencing (GRO-Seq). [ 4 ]


Run on assays have been largely supplanted with Global Run on assays that use next generation DNA sequencing as a readout platform. These assays are known as  GRO-Seq  and provide an incredibly detailed view of genes engaged in transcription with quantitative levels of expression. Array based methods for analyzing Global run on (GRO) assays are being replaced with Next Generation Sequencing which eliminates the design of probes against gene sequences. Sequencing will catalog all transcripts produced even if they are not reported in databases. GRO-seq involves the labeling of newly synthesized transcripts with bromouridine (BrU). Cells or nuclei are incubated with BrUTP in the presence of  sarkosyl , which prevents the attachment of RNA polymerase to the DNA. Therefore only RNA polymerase that are already on the DNA before the addition of sarkosyl will produce new transcripts that will be labeled with BrU. The labeled transcripts are captured with anti-BrU antibody labeled beads, converted to cDNAs and then sequenced by Next Generation DNA sequencing. The sequencing reads are then aligned to the genome and number of reads per transcript provide an accurate estimate of the number of transcripts synthesized. [ 5 ]


In molecular biology,  hybridization  (or  hybridisation ) is a phenomenon in which single-stranded deoxyribonucleic acid ( DNA ) or ribonucleic acid ( RNA ) molecules  anneal  to  complementary DNA or RNA . [ 1 ]  Though a double-stranded DNA sequence is generally stable under physiological conditions, changing these conditions in the laboratory (generally by raising the surrounding temperature) will cause the molecules to separate into single strands. These strands are complementary to each other but may also be complementary to other sequences present in their surroundings. Lowering the surrounding temperature allows the single-stranded molecules to anneal or “hybridize” to each other.


DNA replication  and  transcription  of DNA into RNA both rely upon nucleotide hybridization, as do molecular biology techniques including  Southern blots  and  Northern blots , [ 2 ]  the  polymerase chain reaction  (PCR), and most approaches to  DNA sequencing .


Hybridization is a basic property of nucleotide sequences and is taken advantage of in numerous molecular biology techniques. Overall, genetic relatedness of two species can be determined by hybridizing segments of their DNA ( DNA-DNA hybridization ). Due to sequence similarity between closely related organisms, higher temperatures are required to melt such DNA hybrids when compared to more distantly related organisms. A variety of different methods use hybridization to pinpoint the origin of a DNA sample, including the  polymerase chain reaction  (PCR). In another technique, short DNA sequences are hybridized to cellular mRNAs to identify expressed genes. Pharmaceutical drug companies are exploring the use of antisense RNA to bind to undesired mRNA, preventing the ribosome from translating the mRNA into protein. [ 3 ]


Fluorescence in situ hybridization (FISH) is a laboratory method used to detect and locate a DNA sequence, often on a particular  chromosome . [ 4 ]


In the 1960s, researchers Joseph Gall and Mary Lou Pardue found that molecular hybridization could be used to identify the position of DNA sequences  in situ  (i.e., in their natural positions within a chromosome). In 1969, the two scientists published a paper demonstrating that radioactive copies of a ribosomal DNA sequence could be used to detect complementary DNA sequences in the nucleus of a frog egg. [ 5 ]  Since those original observations, many refinements have increased the versatility and sensitivity of the procedure to the extent that in situ hybridization is now considered an essential tool in  cytogenetics .


Nucleic acid methods  are the techniques used to study  nucleic acids :  DNA  and  RNA .


A  nucleic acid test  ( NAT ) is a technique used to detect a particular  nucleic acid  sequence and thus usually to detect and identify a particular species or subspecies of organism, often a  virus  or  bacterium  that acts as a  pathogen  in  blood ,  tissue ,  urine , etc. NATs differ from other tests in that they detect genetic materials ( RNA  or  DNA ) rather than  antigens  or  antibodies . Detection of genetic materials allows an early diagnosis of a disease because the detection of antigens and/or antibodies requires time for them to start appearing in the bloodstream. [ 1 ]  Since the amount of a certain genetic material is usually very small, many NATs include a step that amplifies the genetic material—that is, makes many copies of it. Such NATs are called  nucleic acid amplification tests  ( NAATs ). There are several ways of amplification, including  polymerase chain reaction  (PCR), strand displacement assay (SDA),  transcription mediated assay  (TMA), [ 2 ]  and  loop-mediated isothermal amplification  (LAMP). [ 3 ]


Virtually all nucleic acid amplification methods and detection technologies use the specificity of Watson-Crick  base pairing ; single-stranded probe or  primer  molecules capture  DNA  or  RNA  target molecules of  complementary strands . Therefore, the design of probe strands is highly significant to raise the  sensitivity and specificity  of the detection. However, the  mutants  which form the genetic basis for a variety of human diseases are usually slightly different from the normal nucleic acids. Often, they are only different in a single base, e.g.,  insertions ,  deletions , and  single-nucleotide polymorphisms  (SNPs). In this case, imperfect probe-target binding can easily occur, resulting in  false-positive  outcomes such as mistaking a  strain  that is  commensal  for one that is pathogenic. Much research has been dedicated to achieving single-base specificity.


Nucleic acid  (DNA and RNA) strands with corresponding sequences stick together in pairwise chains, zipping up like Velcro tumbled in a clothes dryer. But each node of the chain is not very sticky, so the double-stranded chain is continuously coming partway unzipped and re-zipping itself under the influence of ambient vibrations (referred to as thermal noise or  Brownian motion ). Longer pairings are more stable. Nucleic acid tests use a "probe" which is a long strand with a short strand stuck to it. The long primer strand has a corresponding (complementary) sequence to a "target" strand from the disease organism being detected. The disease strand sticks tightly to the exposed part of the long primer strand (called the "toehold"), and then little by little, displaces the short "protector" strand from the probe. In the end, the short protector strand is not bound to anything, and the unbound short primer is detectable. The rest of this section gives some history of the research needed to fine-tune this process into a useful test.


In 2012, Yin's research group published a paper about optimizing the specificity of nucleic acid hybridization. [ 4 ]  They introduced a ‘toehold exchange probe (PC)’ which consists of a pre-hybridized complement strand C and a protector strand P. Complement strand is longer than protector strand to have unbound tail in the end, a toehold. Complement is perfectly complementary with the target sequence. When the correct target(X) reacts with the toehold exchange probe(PC), P is released and hybridized product XC is formed. The standard free energy(∆) of the reaction is close to zero. On the other hand, if the toehold exchange probe(PC) reacts with spurious target(S), the reaction forwards, but the standard free energy increases to be less thermodynamically favorable. The standard free energy difference(∆∆) is significant enough to give obvious discrimination in yield. The discrimination factor Q is calculated as, the yield of correct target hybridization divided by the yield of spurious target hybridization. Through the experiments on different toehold exchange probes with 5 correct targets and 55 spurious targets with energetically representative single-base changes (replacements, deletions, and insertions), Yin's group concluded that discrimination factors of these probes were between 3 and 100 + with the median 26. The probes function robustly from 10 °C to 37 °C, from 1 mM to 47 mM, and with nucleic acid concentrations from 1 nM to 5 M. They also figured out the toehold exchange probes work robustly even in RNA detection.


Further researches have been studied thereafter. In 2013, Seelig's group published a paper about fluorescent molecular probes which also utilizes the toehold exchange reaction. [ 5 ]   This enabled the optical detection of correct target and SNP target. They also succeeded in the detection of SNPs in E. coli-derived samples.


In 2015, David's group achieved extremely high (1,000+) selectivity of single-nucleotide variants (SNVs) by introducing the system called ‘competitive compositions’. [ 6 ]   In this system, they constructed a kinetic reaction model of the underlying hybridization processes to predict the optimal parameter values, which vary based on the sequences of SNV and wildtype (WT), on the design architecture of the probe and sink, and on the reagent concentrations and assay conditions. Their model succeeded in a median 890-fold selectivity for 44 cancer-related DNA SNVs, with a minimum of 200, which represents at least a 30-fold improvement over previous hybridization-based assays. In addition, they applied this technology to assay low VAF sequences from human genomic DNA following PCR, as well as directly to synthetic RNA sequences.


Based on the expertise, they developed a new PCR method called Blocker Displacement Amplification (BDA). [ 7 ]  It is a temperature-robust PCR which selectively amplifies all sequence variants within a roughly 20 nt window by 1000-fold over wildtype sequences, allowing easy detection and quantitation of hundreds of potentials variants originally at ≤ 0.1% allele frequency.  BDA achieves similar enrichment performance across anneal temperatures ranging from 56 °C to 64 °C. This temperature robustness facilitates multiplexed enrichment of many different variants across the genome, and furthermore enables the use of inexpensive and portable thermocycling instruments for rare DNA variant detection. BDA has been validated even on sample types including clinical cell-free DNA samples collected from the blood plasma of lung cancer patients.


Oligonucleotide synthesis  is the chemical synthesis of relatively short fragments of  nucleic acids  with defined chemical structure ( sequence ). The technique is extremely useful in current laboratory practice because it provides a rapid and inexpensive access to custom-made  oligonucleotides  of the desired sequence. Whereas  enzymes  synthesize  DNA  and  RNA  only in a  5' to 3' direction , chemical oligonucleotide synthesis does not have this limitation, although it is most often carried out in the opposite, 3' to 5' direction. Currently, the process is implemented as  solid-phase synthesis  using  phosphoramidite  method and phosphoramidite building blocks derived from protected  2'-deoxynucleosides  ( dA ,  dC ,  dG , and  T ),  ribonucleosides  ( A ,  C ,  G , and  U ), or chemically modified nucleosides, e.g.  LNA  or  BNA .


To obtain the desired oligonucleotide, the building blocks are sequentially coupled to the growing oligonucleotide chain in the order required by the sequence of the product (see  Synthetic cycle  below). The process has been fully automated since the late 1970s. Upon the completion of the chain assembly, the product is released from the solid phase to solution, deprotected, and collected. The occurrence of side reactions sets practical limits for the length of synthetic oligonucleotides (up to about 200  nucleotide  residues) because the number of errors accumulates with the length of the oligonucleotide being synthesized. [ 1 ]  Products are often isolated by  high-performance liquid chromatography  (HPLC) to obtain the desired oligonucleotides in high purity. Typically, synthetic oligonucleotides are single-stranded DNA or RNA molecules around 15–25 bases in length.


Oligonucleotides find a variety of applications in molecular biology and medicine. They are most commonly used as  antisense  oligonucleotides,  small interfering RNA ,  primers  for  DNA sequencing  and  amplification ,  probes  for detecting complementary DNA or RNA via molecular  hybridization , tools for the targeted introduction of  mutations  and  restriction sites , and for the  synthesis of artificial genes . An emerging application of Oligonucleotide synthesis is the re-creation of viruses from sequence alone — either harmless, such as  Phi X 174 , or dangerous such as the 1917  influenza  virus or  SARS-CoV-2 .


The evolution of oligonucleotide synthesis saw four major methods of the formation of internucleosidic linkages and has been reviewed in the literature in great detail. [ 2 ] [ 3 ] [ 4 ]


In the early 1950s,  Alexander Todd 's group pioneered H-phosphonate and  phosphate  triester methods of oligonucleotide synthesis. [ 5 ] [ 6 ]  The reaction of compounds  1  and  2  to form H-phosphonate diester  3  is an H-phosphonate coupling in solution while that of compounds  4  and  5  to give  6  is a phosphotriester coupling (see phosphotriester synthesis below).


Thirty years later, this work inspired, independently, two research groups to adopt the H-phosphonate chemistry to the solid-phase synthesis using nucleoside H-phosphonate monoesters  7  as building blocks and pivaloyl chloride, 2,4,6-triisopropylbenzenesulfonyl chloride (TPS-Cl), and other compounds as activators. [ 7 ] [ 8 ]   The practical implementation of H-phosphonate method resulted in a very short and simple synthetic cycle consisting of only two steps, detritylation and coupling (Scheme 2).  Oxidation  of internucleosidic H-phosphonate diester linkages in  8  to phosphodiester linkages in  9  with a solution of  iodine  in aqueous  pyridine  is carried out at the end of the chain assembly rather than as a step in the synthetic cycle. If desired, the oxidation may be carried out under anhydrous conditions. [ 9 ]  Alternatively,  8  can be converted to phosphorothioate  10 [ 10 ] [ 11 ] [ 12 ] [ 13 ]  or phosphoroselenoate  11  (X = Se), [ 14 ]  or oxidized by  CCl 4  in the presence of primary or secondary amines to phosphoramidate analogs  12 . [ 15 ] [ 16 ]  The method is very convenient in that various types of phosphate modifications (phosphate/phosphorothioate/phosphoramidate) may be introduced to the same oligonucleotide for modulation of its properties. [ 17 ] [ 18 ] [ 19 ]


Most often, H-phosphonate building blocks are protected at the 5'-hydroxy group and at the amino group of nucleic bases A, C, and G in the same manner as phosphoramidite building blocks (see below). However, protection at the amino group is not mandatory. [ 9 ] [ 20 ]


In the 1950s,  Har Gobind Khorana  and co-workers developed a  phosphodiester  method where 3'- O -acetylnucleoside-5'- O -phosphate  2  (Scheme 3) was activated with  N , N ' -dicyclohexylcarbodiimide  (DCC) or  4-toluenesulfonyl chloride  (Ts-Cl). The activated species were reacted with a 5'- O -protected nucleoside  1  to give a protected dinucleoside monophosphate  3 . [ 21 ]  Upon the removal of 3'- O -acetyl group using base-catalyzed hydrolysis, further chain elongation was carried out. Following this methodology, sets of tri- and tetradeoxyribonucleotides were synthesized and were enzymatically converted to longer oligonucleotides, which allowed elucidation of the  genetic code . The major limitation of the phosphodiester method consisted in the formation of pyrophosphate oligomers and oligonucleotides branched at the internucleosidic phosphate. The method seems to be a step back from the more selective chemistry described earlier; however, at that time, most phosphate-protecting groups available now had not yet been introduced. The lack of the convenient protection strategy necessitated taking a retreat to a slower and less selective chemistry to achieve the ultimate goal of the study. [ 2 ]


In the 1960s, groups led by R. Letsinger [ 22 ]  and C. Reese [ 23 ]  developed a phosphotriester approach. The defining difference from the phosphodiester approach was the protection of the phosphate moiety in the building block  1  (Scheme 4) and in the product  3  with  2-cyanoethyl  group. This precluded the formation of oligonucleotides branched at the internucleosidic phosphate. The higher selectivity of the method allowed the use of more efficient coupling agents and catalysts, [ 24 ] [ 25 ]  which dramatically reduced the length of the synthesis. The method, initially developed for the solution-phase synthesis, was also implemented on low-cross-linked "popcorn" polystyrene, [ 26 ]  and later on controlled pore glass (CPG, see "Solid support material" below), which initiated a massive research effort in solid-phase synthesis of oligonucleotides and eventually led to the automation of the oligonucleotide chain assembly.


In the 1970s, substantially more reactive P(III) derivatives of nucleosides, 3'- O -chlorophosphites, were successfully used for the formation of internucleosidic linkages. [ 27 ]  This led to the discovery of the  phosphite triester  methodology. The group led by M. Caruthers took the advantage of less aggressive and more selective 1 H -tetrazolidophosphites and implemented the method on solid phase. [ 28 ]  Very shortly after, the workers from the same group further improved the method by using more stable nucleoside phosphoramidites as building blocks. [ 29 ]  The use of 2-cyanoethyl phosphite-protecting group [ 30 ]  in place of a less user-friendly  methyl  group [ 31 ] [ 32 ]  led to the nucleoside phosphoramidites currently used in oligonucleotide synthesis (see Phosphoramidite building blocks below). Many later improvements to the manufacturing of building blocks, oligonucleotide synthesizers, and synthetic protocols made the phosphoramidite chemistry a very reliable and expedient method of choice for the preparation of synthetic oligonucleotides. [ 1 ]


As mentioned above, the naturally occurring nucleotides (nucleoside-3'- or 5'-phosphates) and their phosphodiester analogs are insufficiently reactive to afford an expeditious synthetic preparation of oligonucleotides in high yields. The selectivity and the rate of the formation of internucleosidic linkages is dramatically improved by using 3'- O -( N , N -diisopropyl phosphoramidite) derivatives of nucleosides (nucleoside phosphoramidites) that serve as building blocks in phosphite triester methodology. To prevent undesired side reactions, all other functional groups present in nucleosides have to be rendered unreactive (protected) by attaching  protecting groups . Upon the completion of the oligonucleotide chain assembly, all the protecting groups are removed to yield the desired oligonucleotides. Below, the protecting groups currently used in commercially available [ 33 ] [ 34 ] [ 35 ] [ 36 ]  and most common nucleoside phosphoramidite building blocks are briefly reviewed:


The protection of the exocyclic amino groups has to be orthogonal to that of the 5'-hydroxy group because the latter is removed at the end of each synthetic cycle. The simplest to implement, and hence the most widely used,  strategy is to install a base-labile protection group on the exocyclic amino groups. Most often, two protection schemes are used.


Non-nucleoside phosphoramidites are the phosphoramidite reagents designed to introduce various functionalities at the termini of synthetic oligonucleotides or between nucleotide residues in the middle of the sequence. In order to be introduced inside the sequence, a non-nucleosidic modifier has to possess at least two hydroxy groups, one of which is often protected with the DMT group while the other bears the reactive phosphoramidite moiety.


Non-nucleosidic phosphoramidites are used to introduce desired groups that are not available in natural nucleosides or that can be introduced more readily using simpler chemical designs. A very short selection of commercial phosphoramidite reagents is shown in Scheme for the demonstration of the available structural and functional diversity. These reagents serve for the attachment of 5'-terminal phosphate ( 1 ), [ 52 ]  NH 2  ( 2 ), [ 53 ]  SH ( 3 ), [ 54 ]  aldehydo ( 4 ), [ 55 ]  and carboxylic groups ( 5 ), [ 56 ]  CC triple bonds ( 6 ), [ 57 ]  non-radioactive labels and  quenchers  (exemplified by  6-FAM amidite   7 [ 58 ]  for the attachment of  fluorescein  and dabcyl amidite  8 , [ 59 ]  respectively),  hydrophilic and hydrophobic modifiers (exemplified by  hexaethyleneglycol  amidite  9 [ 60 ] [ 61 ]  and  cholesterol  amidite  10 , [ 62 ]  respectively), and  biotin  amidite  11 . [ 63 ]


Oligonucleotide synthesis is carried out by a stepwise addition of nucleotide residues to the 5'-terminus of the growing chain until the desired sequence is assembled. Each addition is referred to as a synthesis cycle (Scheme 5) and consists of four chemical reactions:


The DMT group is removed with a solution of an acid, such as 2%  trichloroacetic acid  (TCA) or 3%  dichloroacetic acid  (DCA), in an inert solvent ( dichloromethane  or  toluene ). The orange-colored DMT cation formed is washed out; the step results in the solid support-bound oligonucleotide precursor bearing a free 5'-terminal hydroxyl group.
It is worth remembering that conducting detritylation for an extended time or with stronger than recommended solutions of acids leads to  depurination  of solid support-bound oligonucleotide and thus reduces the yield of the desired full-length product. [ citation needed ]


A 0.02–0.2 M solution of nucleoside phosphoramidite (or a mixture of several phosphoramidites) in  acetonitrile  is activated by a 0.2–0.7 M solution of an acidic  azole  catalyst,  1 H -tetrazole , 5-ethylthio-1H-tetrazole, [ 64 ]  2-benzylthiotetrazole, [ 65 ] [ 66 ]  4,5-dicyano imidazole , [ 67 ]  or a number of similar compounds. A more extensive information on the use of various coupling agents in oligonucleotide synthesis can be found in a recent review. [ 68 ]   The mixing is usually very brief and occurs in fluid lines of oligonucleotide synthesizers (see below) while the components are being delivered to the reactors containing solid support. The activated phosphoramidite in 1.5 – 20-fold excess over the support-bound material is then brought in contact with the starting solid support (first coupling) or a support-bound oligonucleotide precursor (following couplings) whose 5'-hydroxy group reacts with the activated phosphoramidite moiety of the incoming nucleoside phosphoramidite to form a phosphite triester linkage. The coupling of 2'-deoxynucleoside phosphoramidites is very rapid and requires, on small scale, about 20 s for its completion. In contrast, sterically hindered 2'- O -protected ribonucleoside phosphoramidites require 5-15 min to be coupled in high yields. [ 47 ] [ 69 ] [ 70 ] [ 71 ]  The reaction is also highly sensitive to the presence of water, particularly when dilute solutions of phosphoramidites are used, and is commonly carried out in anhydrous acetonitrile. Generally, the larger the scale of the synthesis, the lower the excess and the higher the concentration of the phosphoramidites is used. In contrast, the concentration of the activator is primarily determined by its solubility in acetonitrile and is irrespective of the scale of the synthesis. Upon the completion of the coupling, any unbound reagents and by-products are removed by washing.


The capping step is performed by treating the solid support-bound material with a mixture of acetic anhydride and  1-methylimidazole  or, less often,  DMAP  as catalysts and, in the phosphoramidite method, serves two purposes.


The newly formed tricoordinated phosphite triester linkage is not natural and is of limited stability under the conditions of oligonucleotide synthesis. The treatment of the support-bound material with iodine and water in the presence of a weak base (pyridine,  lutidine , or  collidine )  oxidizes  the phosphite triester into a tetracoordinated phosphate triester, a protected precursor of the naturally occurring phosphate diester internucleosidic linkage. Oxidation may be carried out under anhydrous conditions using  tert-Butyl hydroperoxide [ 74 ]  or, more efficiently, (1S)-(+)-(10-camphorsulfonyl)-oxaziridine (CSO). [ 75 ] [ 76 ] [ 77 ]  The step of oxidation may be substituted with a sulfurization step to obtain oligonucleotide phosphorothioates (see  Oligonucleotide phosphorothioates and their synthesis  below). In the latter case, the sulfurization step is best carried out prior to capping.


In solid-phase synthesis, an oligonucleotide being assembled is  covalently  bound, via its 3'-terminal hydroxy group, to a solid support material and remains attached to it over the entire course of the chain assembly. The solid support is contained in columns whose dimensions depend on the scale of synthesis and may vary between 0.05  mL  and several liters. The overwhelming majority of oligonucleotides are synthesized on small scale ranging from 10 n mol  to 1 μmol. More recently, high-throughput oligonucleotide synthesis where the solid support is contained in the wells of multi-well plates (most often, 96 or 384 wells per plate) became a method of choice for parallel synthesis of oligonucleotides on small scale. [ 78 ]  At the end of the chain assembly, the oligonucleotide is released from the solid support and is eluted from the column or the well.


In contrast to organic solid-phase synthesis and  peptide synthesis , the synthesis of oligonucleotides proceeds best on non-swellable or low-swellable solid supports. The two most often used solid-phase materials are controlled pore glass (CPG) and macroporous  polystyrene  (MPPS). [ 79 ]


To make the solid support material suitable for oligonucleotide synthesis, non-nucleosidic linkers or nucleoside succinates are covalently attached to the reactive amino groups in aminopropyl CPG, LCAA CPG, or aminomethyl MPPS. The remaining unreacted amino groups are capped with  acetic anhydride . Typically, three conceptually different groups of solid supports are used.


Oligonucleotide phosphorothioates (OPS) are modified oligonucleotides where one of the oxygen atoms in the phosphate moiety is replaced by sulfur. Only the phosphorothioates having sulfur at a non-bridging position as shown in figure are widely used and are available commercially. The replacement of the non-bridging oxygen with sulfur creates a new center of  chirality  at  phosphorus . In a simple case of a dinucleotide, this results in the formation of a  diastereomeric  pair of S p - and R p -dinucleoside monophosphorothioates whose structures are shown in Figure. In an  n -mer oligonucleotide where all ( n  – 1) internucleosidic linkages are phosphorothioate linkages, the number of diastereomers  m  is calculated as  m  = 2 ( n  – 1) . Being non-natural analogs of nucleic acids, OPS are substantially more stable towards  hydrolysis  by  nucleases , the class of  enzymes  that destroy nucleic acids by breaking the bridging P-O bond of the phosphodiester moiety. This property determines the use of OPS as antisense oligonucleotides in  in vitro  and  in vivo  applications where the extensive exposure to nucleases is inevitable. Similarly, to improve the stability of  siRNA , at least one phosphorothioate linkage is often introduced at the 3'-terminus of both  sense  and antisense strands. In chirally pure OPS, all-Sp diastereomers are more stable to enzymatic degradation than their all-Rp analogs. [ 91 ]  However, the preparation of chirally pure OPS remains a synthetic challenge. [ 13 ] [ 92 ]  In laboratory practice, mixtures of diastereomers of OPS are commonly used.


Synthesis of OPS is very similar to that of natural oligonucleotides. The difference is that the oxidation step is replaced by sulfur transfer reaction (sulfurization) and that the capping step is performed after the sulfurization. Of many reported reagents capable of the efficient sulfur transfer, only three are commercially available:


In the past, oligonucleotide synthesis was carried out manually in solution or on solid phase. The solid phase synthesis was implemented using, as containers for the solid phase, miniature glass columns similar in their shape to low-pressure chromatography columns or syringes equipped with porous filters. [ 101 ]
Currently, solid-phase oligonucleotide synthesis is carried out automatically using computer-controlled instruments (oligonucleotide synthesizers) and is technically implemented in column, multi-well plate, and array formats. The column format is best suited for research and large scale applications where a high-throughput is not required. [ 102 ]  Multi-well plate format is designed specifically for high-throughput synthesis on small scale to satisfy the growing demand of industry and academia for synthetic oligonucleotides. [ 103 ]


Large scale oligonucleotide synthesizers were often developed by augmenting the capabilities of a preexisting instrument platform. One of the first mid scale synthesizers appeared in the late 1980s, manufactured by the Biosearch company in Novato, CA (The 8800). This platform was originally designed as a peptide synthesizer and made use of a fluidized bed reactor essential for accommodating the swelling characteristics of polystyrene supports used in the Merrifield methodology. Oligonucleotide synthesis involved the use of CPG (controlled pore glass) which is a rigid support and is more suited for column reactors as described above. The scale of the 8800 was limited to the flow rate required to fluidize the support. Some novel reactor designs as well as higher than normal pressures enabled the 8800 to achieve scales that would prepare 1 mmol of oligonucleotide. In the mid 1990s several companies developed platforms that were based on semi-preparative and preparative liquid chromatographs. These systems were well suited for a column reactor approach. In most cases all that was required was to augment the number of fluids that could be delivered to the column. Oligo synthesis requires a minimum of 10 and liquid chromatographs usually accommodate 4. This was an easy design task and some semi-automatic strategies worked without any modifications to the preexisting LC equipment. PerSeptive Biosystems as well as Pharmacia (GE) were two of several companies that developed synthesizers out of liquid chromatographs. Genomic Technologies, Inc. [ 104 ]  was one of the few companies to develop a large scale oligonucleotide synthesizer that was, from the ground up, an oligonucleotide synthesizer. The initial platform called the VLSS for very large scale synthesizer utilized large Pharmacia liquid chromatograph columns as reactors and could synthesize up to 75 mmol of material. Many oligonucleotide synthesis factories designed and manufactured their own custom platforms and little is known due to the designs being proprietary. The VLSS design continued to be refined and is continued in the QMaster synthesizer [ 105 ]  which is a scaled down platform providing milligram to gram amounts of synthetic oligonucleotide.


The current practices of synthesis of chemically modified oligonucleotides on large scale have been recently reviewed. [ 106 ]


One may visualize an oligonucleotide microarray as a miniature multi-well plate where physical dividers between the wells (plastic walls) are intentionally removed. With respect to the chemistry, synthesis of oligonucleotide microarrays is different from the conventional oligonucleotide synthesis in two respects:


After the completion of the chain assembly, the solid support-bound oligonucleotide is fully protected:


To furnish a functional oligonucleotide, all the protecting groups have to be removed. The N-acyl base protection and the 2-cyanoethyl phosphate protection may be, and is often removed simultaneously by treatment with inorganic bases or amines. However, the applicability of this method is limited by the fact that the cleavage of 2-cyanoethyl phosphate protection gives rise to  acrylonitrile  as a side product. Under the strong basic conditions required for the removal of N-acyl protection, acrylonitrile is capable of alkylation of nucleic bases, primarily, at the N3-position of thymine and uracil residues to give the respective N3-(2-cyanoethyl) adducts via  Michael reaction . The formation of these side products may be avoided by treating the solid support-bound oligonucleotides with solutions of bases in an organic solvent, for instance, with 50%  triethylamine  in  acetonitrile [ 109 ]  or 10%  diethylamine  in acetonitrile. [ 110 ]  This treatment is strongly recommended for medium- and large scale preparations and is optional for syntheses on small scale where the concentration of acrylonitrile generated in the deprotection mixture is low.


Regardless of whether the phosphate protecting groups were removed first, the solid support-bound oligonucleotides are deprotected using one of the two general approaches.


As with any other organic compound, it is prudent to characterize synthetic oligonucleotides upon their preparation. In more complex cases (research and large scale syntheses) oligonucleotides are characterized after their deprotection and after purification. Although the ultimate approach to the characterization is  sequencing , a relatively inexpensive and routine procedure, the considerations of the cost reduction preclude its use in routine manufacturing of oligonucleotides. In day-by-day practice, it is sufficient to obtain the  molecular mass  of an oligonucleotide by recording its  mass spectrum . Two methods are currently widely used for characterization of oligonucleotides:  electrospray mass spectrometry  (ESI MS) and  matrix-assisted laser desorption/ionization   time-of-flight mass spectrometry  ( MALDI-TOF ). To obtain informative spectra, it is very important to exchange all metal ions that might be present in the sample for ammonium or trialkylammonium [ e.c.  triethylammonium, (C 2 H 5 ) 3 NH + ] ions prior to submitting a sample to the analysis by either of the methods.


DNA paternity testing  is the use of  DNA profiles  to determine whether an individual is the  biological  parent of another individual. Paternity testing can be especially important when the rights and duties of the father are in issue and a child's  paternity  is in doubt. Tests can also determine the likelihood of someone being a biological grandparent. Though  genetic  testing is the most reliable standard, older methods also exist, including  ABO blood group typing , analysis of various other  proteins  and  enzymes , or using  human leukocyte antigen   antigens . The current techniques for paternity testing are using  polymerase chain reaction  (PCR) and  restriction fragment length polymorphism  (RFLP). Paternity testing can now also be performed while the woman is still pregnant from a blood draw. [ 1 ] [ 2 ]


DNA testing is currently the most advanced and accurate technology to determine parentage. In a DNA paternity test, the result (called the 'probability of parentage) [ 3 ] [ failed verification ]  is 0% when the alleged parent is not biologically related to the child, and the probability of parentage is typically 99.99% when the alleged parent is biologically related to the child. However, while almost all individuals have a single and distinct set of genes, rare individuals, known as " chimeras ", have at least two different sets of genes, which can result in a false negative result if their reproductive tissue has a different genetic make-up from the tissue sampled for the test. [ 4 ]


The DNA test is performed by collecting buccal (cheek) cells found on the inside of a person's cheek using a buccal or cheek  swab . These swabs have wooden or plastic stick handles with a cotton on synthetic tip. The collector rubs the inside of a person's cheek to collect as many buccal cells as possible, which are then sent to a laboratory for testing. Samples from the alleged father or mother and the child would be needed.


It is possible to determine who the biological father of the fetus is while the woman is still pregnant through procedures called  chorionic villus sampling  or  amniocentesis . Chorionic villus sampling retrieves placental tissue in either a transcervical or transabdominal manner. Amniocentesis retrieves amniotic fluid by inserting a needle through the pregnant mother's abdominal wall. These procedures are highly accurate because they are taking a sample directly from the fetus; however, there is a small risk for the woman to miscarry and lose the pregnancy as a result. Both CVS and amniocentesis require the pregnant woman to visit a genetic specialist known as a maternal-fetal medicine specialist who will perform the procedure.


Advances in  genetic testing  have led to the ability to identify the biological father while the woman is still pregnant. There is a small amount of fetal DNA ( cffDNA ) present in the mother's blood during pregnancy. This allows for accurate fetal DNA paternity testing during pregnancy from a blood draw with no risk of miscarriage. Studies have shown that cffDNA can first be observed as early as seven weeks gestation, and the amount of cffDNA increases as the pregnancy progresses. [ 5 ] [ 6 ]


The  DNA  of an individual is the same in every somatic (nonreproductive)  cell .  Sexual reproduction  brings the DNA of both parents together to create a unique combination of genetic material in a new cell, so the genetic material of an individual is derived from the genetic material of each parent in equal amounts; this genetic material is known as the nuclear  genome  of the individual, because it is found in the  nucleus .


Comparing the DNA sequence of one person to that of another can prove if one of them was derived from the other, but DNA paternity tests are not currently 100% accurate. Specific sequences are examined to see if they were copied verbatim from one individual's genome; if so, then the genetic material of one individual could have been derived from that of the other (i.e. one is the parent of the other). This is called Autosomal DNA testing. It is currently the gold standard in paternity testing as it allows a comparison of the child’s DNA to that of the mother and alleged father. The genetic contribution to the child from the mother can be evaluated, resulting in possible genotypes for the true father. If the alleged father cannot be excluded as the true father, then statistical calculations can be conducted to determine how likely the alleged father is the true father compared to if another random man was the true father. [ 7 ]


Besides nuclear DNA,  mitochondria  also have their own genetic material called  mitochondrial DNA . Mitochondrial DNA comes only from the mother, without any shuffling. Proving a relationship based on comparison of the mitochondrial genome is much easier than that based on the nuclear genome. However, testing the mitochondrial genome can prove only if two individuals are related by  common descent  through maternal lines only from a common ancestor and is, thus, of limited value (i.e., it could not be used to test for paternity).


In testing the paternity of a male child, comparison of the  Y chromosome  can be used, since it is passed directly from father to son. However, similar to mitochondrial DNA, the Y chromosome is passed through the paternal line. This means that two brothers share the Y chromosome of their father. Therefore if one brother is the suspected father, his biological brother could also be the father based on Y chromosomal data alone. This is true with any male related to the suspected father on the paternal line. For this reason autosomal DNA testing would be a more precise paternity testing method.  [ 8 ]


In the US, the  AABB  has regulations for DNA paternity and family relationship testing, but AABB accreditation is not required. DNA test results are legally admissible if the collection and the processing follows a chain of custody. Similarly in Canada, the SCC has regulations on DNA paternity and relationship testing, but this accreditation, while recommended, is not required.


The Paternity Testing Commission of the  International Society for Forensic Genetics  has taken up the task of establishing the biostatistical recommendations in accordance with the  ISO/IEC 17025  standards. [ 9 ]  Bio-statistical evaluations of paternity should be based on a likelihood ratio principle - yielding the  Paternity Index , PI. The recommendations provide guidance on concepts of genetic hypotheses and calculation concerns needed to produce valid PIs, as well as on specific issues related to  population genetics .


The first form of any kind of parental testing was blood typing, or matching  blood types  between the child and alleged parent, which became available in the 1920s, after scientists recognized that blood types, which had been discovered in the early 1900s, were genetically inherited. Under this form of testing, the blood types of the child and parents are compared, and it can be determined whether there is any possibility of a parental link. For example, two O blood type parents can produce a child only with an O blood type, and two parents with a B blood type can produce a child with either a B or an O blood type. This often led to inconclusive results, as 30% of the entire population can be excluded from being the possible parent under this form of testing. [ 10 ]  In the 1930s,  serological  testing, which tests certain proteins in the blood, became available, with a 40% exclusion rate. [ 11 ]


In the 1960s, accurate genetic paternity testing became a possibility when  HLA  typing was developed, which compares the genetic fingerprints on white blood cells between the child and alleged parent. [ 12 ]  HLA tests could be done with 80% accuracy but could not distinguish between close relatives. [ 13 ]  Genetic parental testing technology advanced further with the isolation of the first  restriction enzyme  in 1970. Highly accurate DNA parental testing became available in the 1980s with the development of  RFLP . In the 1990s,  PCR  became the standard method for DNA parental testing: a simpler, faster, and more accurate method of testing than RFLP, it has an exclusion rate of 99.99% or higher. [ 13 ]


The DNA parentage test that follows strict  chain of custody  can generate legally admissible results that are used for child support, inheritance, social welfare benefits, immigration, or adoption purposes. To satisfy the chain-of-custody legal requirements, all tested parties have to be properly identified and their specimens collected by a third-party professional who is not related to any of the tested parties and has no interest in the outcome of the test.


The  quantum of evidence  needed is  clear and convincing evidence : that is, more evidence than an ordinary case in  civil litigation , but less than beyond a  reasonable doubt  required to convict a defendant in a criminal case.


In recent years, immigration authorities in various countries, such as the United States, United Kingdom, Canada, Australia, France, and others, may accept DNA parentage test results from immigration petitioners and beneficiaries in a family-based immigration case when primary documents that prove biological relationship are missing or inadequate.


In the U.S., immigration applicants bear the responsibility of arranging and paying for DNA testing. The U.S. immigration authorities require that the DNA test, if pursued, be performed by one of the laboratories accredited by the  AABB  (formerly American Association of Blood Banks). Similarly, in Canada, the laboratory needs to be accredited by the  Standards Council of Canada .


Although paternity tests are more common than maternity tests, there may be circumstances in which the biological mother of the child is unclear: examples include cases of an  adopted  child attempting to reunify with his or her biological mother, potential  hospital mix-ups , and  in vitro fertilization  where the laboratory may have implanted an unrelated embryo inside the mother.


Other factors, such as new laws regarding reproductive technologies using donated eggs and sperm and surrogate mothers, can also mean that the female giving birth is not necessarily the legal mother of the child. For example, in Canada, the federal  Human Assisted Reproduction Act  provides for the use of hired surrogate mothers. The legal mother of the child may be the egg donor. Similar laws are in place in the United Kingdom and Australia.


In Brazil in 2019, two male identical twins were ordered to both pay maintenance for a child fathered by one of them, because the father could not be identified with DNA. [ 14 ]


Peace-of-mind parentage tests are widely available on the internet. For a parentage test (paternity or maternity) to be admissible for legal purposes, such as for changing a birth certificate, Family Law Court proceedings, visa/citizenship applications or child support claims, the process must comply with the  Family Law Regulations 1984  (Cth). [ 15 ]  Further, the laboratory processing the samples must be accredited by the National Association of Testing Authorities (NATA). [ 16 ]


Personal paternity-testing kits are available. The Standards Council of Canada regulates paternity testing in Canada whereby laboratories are ISO 17025-approved. In Canada, only a handful of labs have this approval, and it is recommended that testing is performed in these labs. Courts also have the power to order paternity tests during divorce cases. [ 17 ]


In  China , paternity testing is legally available to fathers who suspect their child is not theirs. Chinese law also requires a paternity test for any child born outside the  one-child policy  for the child to be eligible for a  hukou , or family registration record. Family tie formed by adoption can also only be confirmed by a paternity test. A large number of Chinese citizens seek paternity testing each year, and this has given rise to many unlicensed illegal testing centers being set up. [ 18 ]


DNA paternity testing is solely performed on decision of a judge in case of a judiciary procedure in order either to establish or contest paternity or to obtain or deny child support. [ 19 ]  Non consensual private DNA paternity testing is illegal, including through laboratories in other countries, and is punishable by up to a year in prison and a €15,000 fine. [ 20 ]  The  French Council of State  has described the law's purpose as upholding the "French regime of filiation" and preserving "the peace of families." [ 21 ]


Under the Gene Diagnostics Act of 2009, secret paternity testing is illegal. Any paternity testing must be conducted by a licensed physician or by an expert with a university degree in science and special education in parentage testing, and the laboratory carrying out genetic testing must be accredited according to  ISO/IEC 17025 . Full informed consent of both parents is required, and prenatal paternity testing is prohibited, with the exception of sexual abuse and rape cases. Any genetic testing done without the other parent's consent is punishable with a €5,000 fine. [ 22 ]  Due to an amendment of the civil law  section 1598a  in 2005, any man who contests paternity no longer automatically severs legal rights and obligations to the child. [ 23 ] [ 24 ]


A paternity test with any legal standing must be ordered by a family court. Though parents have access to "peace of mind" parental tests through overseas laboratories, family courts are under no obligation to accept them as evidence. It is also illegal to take genetic material for a parental test from a minor over 16 years of age without the minor's consent. Family courts have the power to order paternity tests against the will of the father in divorce and child support cases, as well as in other cases such as determining heirs and settling the question involving the population registry. A man seeking to prove that he is not the father of the child registered as his is entitled to a paternity test, even if the mother and natural guardian object. Paternity tests are not ordered when it is believed it could lead to the murder of the mother, and until 2007, were not ordered when there was a chance that the child of a married woman could have been fathered by a man other than her husband, thereby making the child a  mamzer  under Jewish law. [ 25 ] [ 26 ] [ 27 ]


DNA paternity testing for personal knowledge is legal, and home test kits are available by mail from representatives of AABB- and ISO 17025-certified laboratories. [ 28 ]  DNA Paternity Testing for official purposes, such as sustento (child support) and inheritance disputes, must follow the Rule on DNA Evidence A.M. No. 06-11-5-SC, which was promulgated by the Philippine Supreme Court on October 15, 2007. [ 29 ]  Tests are sometimes ordered by courts when proof of paternity is required.


In  Spain , peace-of-mind paternity tests are a "big business," partly due to the French ban on paternity testing, with many genetic testing companies being based in Spain. [ 30 ] [ 31 ]


In the  United Kingdom , there were no restrictions on paternity tests until the  Human Tissue Act 2004  came into force in September 2006. Section 45 states that it is an offence to possess without appropriate consent any human bodily material with the intent of analysing its DNA. Legally declared fathers have access to paternity-testing services under the new regulations, provided the putative parental DNA being tested is their own. Tests are sometimes ordered by courts when proof of paternity is required. In the UK, the  Ministry of Justice  accredits bodies that can conduct this testing. The  Department of Health  produced a voluntary code of practice on genetic paternity testing in 2001. This document is currently under review, and responsibility for it has been transferred to the  Human Tissue Authority .
In the 2018 case of  Anderson V Spencer  the Court of Appeal permitted for the very first time DNA samples taken from a Deceased person to be used for paternity testing.


In the  United States , paternity testing is fully legal, and fathers may test their children without the consent or knowledge of the mother. Paternity testing take-home kits are readily available for purchase, though their results are not admissible in court and are for personal knowledge only.


Only a court-ordered paternity test may be used as evidence in court proceedings. If parental testing is being submitted for legal purposes, including immigration, testing must be ordered through a lab that has  AABB  accreditation for relationship DNA testing. [ 32 ]


The legal implications of a parentage result test vary by state and according to whether the putative parents are unmarried or married. If a parentage test does not meet forensic standards for the state in question, a court-ordered test may be required for the results of the test to be admissible for legal purposes. For unmarried parents, if a parent is currently receiving child support or custody, but DNA testing later proves that the man is not the father, support automatically stops. However, in many states, this testing must be performed during a narrow window of time, if a voluntary acknowledgement of parentage form has already been signed by the putative father; otherwise, the results of the test may be disregarded by law, and in many cases, a man may be required to pay child support, though the child is biologically unrelated. In a few states, if the mother is receiving the support, then that alleged father has the right to file a lawsuit to get back any money that he lost from paying support. As of 2011, in most states, unwed parents confronted with a voluntary acknowledgement of parentage form are informed of the possibility and right to request a DNA paternity test. If testing is refused by the mother, the father may not be required to sign the  birth certificate  or the voluntary acknowledgement of parentage form for the child. For wedded putative parents, the husband of the mother is presumed to be the father of the child. But, in most states, this presumption can be overturned by the application of a forensic paternity test; in many states, the time for overturning this presumption may be limited to the first few years of the child's life.


Reverse paternity determination is the ability to establish the biological father when the father of that person is not available. The test uses the STR  alleles  in  mother and her child, other children and brothers of the alleged father, and deduction of genetic constitution of the father by the basis of genetic laws, all to create a rough amalgamation. This can compare the father's DNA when a direct sample of the father's DNA is unavailable. An episode of  Solved  shows this test being used to know if a blood sample matches with the victim of a kidnapping.


Genetic:


In  molecular phylogenetics , relationships among individuals are determined using character traits, such as  DNA ,  RNA  or  protein , which may be obtained using a variety of  sequencing  technologies. High-throughput  next-generation sequencing  has become a popular  technique in transcriptomics , which represent a snapshot of gene expression. In  eukaryotes , making phylogenetic inferences using  RNA  is complicated by  alternative splicing , which produces multiple  transcripts  from a single  gene . As such, a variety of approaches may be used to improve  phylogenetic inference using transcriptomic data  obtained from  RNA-Seq  and processed using  computational phylogenetics .


There have been several  transcriptomics technologies  used to gather sequence information on  transcriptomes . However the most widely used is  RNA-Seq .


RNA reads may be obtained using a variety of RNA-seq methods.


There are a number of  public databases  that contain freely available RNA-Seq data.


RNA-Seq data may be directly assembled into  transcripts  using  sequence assembly .
Two main categories of  sequence assembly  are often distinguished:


Both methods attempt to generate biologically representative isoform-level constructs from RNA-seq data and generally attempt to associate isoforms with a gene-level construct. However, proper identification of gene-level constructs may be complicated by recent  duplications ,  paralogs ,  alternative splicing  or  gene fusions . These complications may also cause downstream issues during ortholog inference. When selecting or generating sequence data, it is also vital to consider the tissue type, developmental stage and environmental conditions of the organisms. Since the  transcriptome  represents a snapshot of  gene expression , minor changes to these conditions may significantly affect which transcripts are expressed. This may detrimentally affect downstream ortholog detection. [ 1 ]


RNA may also be acquired from public databases, such as  GenBank ,  RefSeq ,  1000 Plants (1KP)  and  1KITE . Public databases potentially offer curated sequences which can improve inference quality and avoid the computational overhead associated with  sequence assembly .


Orthology  or  paralogy  inference requires an assessment of  sequence homology , usually via  sequence alignment .  Phylogenetic analyses  and  sequence alignment  are often considered jointly, as phylogenetic analyses using  DNA  or  RNA  require sequence alignment and alignments themselves often represent some hypothesis of  homology . As proper ortholog identification is pivotal to phylogenetic analyses, there are a variety of methods available to infer  orthologs  and  paralogs . [ 2 ]


These methods are generally distinguished as either graph-based algorithms or tree-based algorithms. Some examples of graph-based methods include InParanoid, [ 3 ]  MultiParanoid, [ 4 ]  OrthoMCL, [ 5 ]  HomoloGene [ 6 ]  and OMA. [ 7 ]  Tree-based algorithms include programs such as OrthologID or RIO. [ 8 ] [ 2 ]


A variety of  BLAST  methods are often used to detect  orthologs  between  species  as a part of graph-based algorithms, such as MegaBLAST, BLASTALL, or other forms of all-versus-all BLAST and may be  nucleotide - or  protein -based  alignments . [ 9 ] [ 10 ]  RevTrans [ 11 ]  will even use protein data to inform DNA alignments, which can be beneficial for resolving more distant phylogenetic relationships. These approaches often assume that best-reciprocal-hits passing some threshold metric(s), such as identity, E-value, or percent alignment, represent  orthologs  and may be confounded by  incomplete lineage sorting . [ 12 ] [ 13 ]


It is important to note that orthology relationships in public databases typically represent gene-level orthology and do not provide information concerning conserved  alternative splice variants .


Databases that contain and/or detect orthologous relationships include:


As  eukaryotic transcription  is a complex process by which multiple  transcripts  may be generated from a single  gene  through  alternative splicing  with variable  expression , the utilization of RNA is more complicated than DNA. However,  transcriptomes  are cheaper to sequence than complete genomes and may be obtained without the use of a pre-existing  reference genome . [ 1 ]


It is not uncommon to  translate  RNA sequence into protein sequence when using transcriptomic data, especially when analyzing highly diverged taxa. This is an intuitive step as many (but not all) transcripts are expected to code for  protein isoforms . Potential benefits include the reduction of mutational biases and a reduced number of characters, which may speed analyses. However, this reduction in characters may also result in the loss of potentially informative characters. [ 1 ]


There are a number of  tools available for multiple sequence alignment . All of which possess their own strengths and weaknesses and may be specialized for distinct sequence types (DNA, RNA or protein). As such, a splice-aware aligner may be ideal for aligning RNA sequences, whereas an aligner that considers  protein structure  or  residue substitution rates  may be preferable for translated RNA sequence data.


Using RNA for phylogenetic analysis comes with its own unique set of strengths and weaknesses.


A  plasmid preparation  is a method of  DNA extraction  and purification for  plasmid DNA . It is an important step in many  molecular biology  experiments and is essential for the successful use of plasmids in research and  biotechnology . [ 1 ] [ 2 ]  Many methods have been developed to purify plasmid  DNA  from  bacteria . [ 1 ] [ 3 ]  During the purification procedure, the plasmid DNA is often separated from contaminating proteins and genomic DNA.


These methods invariably involve three steps: growth of the bacterial culture, harvesting and  lysis  of the bacteria, and purification of the plasmid DNA. [ 4 ]  Purification of plasmids is central to  molecular cloning . A purified plasmid can be used for many standard applications, such as  sequencing  and  transfections  into cells.


Plasmids  are almost always purified from liquid  bacteria cultures , usually  E. coli , which have been  transformed  and isolated. [ 5 ] [ 6 ]  Virtually all plasmid vectors in common use encode one or more  antibiotic resistance  genes as a  selectable marker , for example a gene encoding ampicillin or kanamycin resistance, which allows bacteria that have been successfully transformed to multiply uninhibited. [ 7 ] [ 8 ] [ 9 ]  Bacteria that have not taken up the plasmid vector are assumed to lack the resistance gene, and thus only colonies representing successful transformations are expected to grow. [ 5 ] [ 9 ] [ 10 ]
Bacteria are grown under favourable conditions.


There are several methods for cell lysis, including alkaline lysis, mechanical lysis, and enzymatic lysis. [ 11 ] [ 12 ] [ 13 ] [ 14 ]


The most common method is alkaline lysis, which involves the use of a high concentration of a basic solution, such as  sodium hydroxide , to lyse the bacterial cells. [ 15 ] [ 16 ] [ 17 ]  When bacteria are lysed under  alkaline  conditions (pH 12.0–12.5) both chromosomal DNA and  protein  are  denatured ; the plasmid DNA however, remains stable. [ 16 ] [ 17 ]  Some scientists reduce the concentration of NaOH used to 0.1M in order to reduce the occurrence of  ssDNA . After the addition of  acetate -containing neutralization  buffer   to lower the pH to around 7, the large and less  supercoiled   chromosomal  DNA and proteins  form large complexes and precipitate; but the small bacterial DNA plasmids stay in solution. [ 17 ] [ 14 ]


Mechanical lysis involves the use of physical force, such as grinding or  sonication , to break down bacterial cells and release the plasmid DNA. There are several different mechanical lysis methods that can be used, including  French press ,  bead-beating , and  ultrasonication . [ 11 ] [ 12 ] [ 13 ] [ 14 ]


Enzymatic lysis, also called  Lysozyme  lysis, involves the use of  enzymes  to digest the cell wall and release the plasmid DNA. [ 11 ]  The most commonly used enzyme for this purpose is lysozyme, which breaks down the  peptidoglycan  in the cell wall of  Gram-positive bacteria .  Lysozyme  is usually added to the bacterial culture, followed by heating and/or shaking the culture to release the plasmid DNA. [ 11 ] [ 12 ] [ 13 ] [ 14 ]


Plasmid preparation can be divided into five main categories based on the scale of the preparation: minipreparation, midipreparation, maxipreparation, megapreparation, and gigapreparation. The choice of which method to use will depend on the amount of plasmid DNA required, as well as the specific application for which it will be used. [ 18 ] [ 19 ]


Kits are available from varying manufacturers to purify plasmid DNA, which are named by size of bacterial culture and corresponding plasmid yield. In increasing order they are: miniprep, midiprep, maxiprep, megaprep, and gigaprep. The plasmid DNA yield will vary depending on the plasmid copy number, type and size, the  bacterial strain , the growth conditions, and the kit. [ 2 ]


Minipreparation of plasmid DNA is a rapid, small-scale isolation of  plasmid   DNA  from  bacteria . [ 20 ] [ 21 ]  Commonly used miniprep methods include  alkaline lysis  and  spin-column  based kits. [ 3 ] [ 22 ]  It is based on the  alkaline lysis  method. The extracted plasmid DNA resulting from performing a miniprep is itself often called a "miniprep".
Minipreps are used in the process of  molecular cloning  to analyze bacterial  clones . A typical plasmid DNA yield of a miniprep is 5 to 50 μg depending on the cell strain.
Miniprep of a large number of plasmids can also be done conveniently on filter paper by lysing the cell and eluting the plasmid on to filter paper. [ 21 ]


The starting E. coli culture volume is 15-25 mL of  Lysogeny broth  (LB) and the expected DNA yield is 100-350 μg.


The starting E. coli culture volume is 100-200 mL of LB and the expected DNA yield is 500-850 μg.


The starting E. coli culture volume is 500 mL – 2.5 L of LB and the expected DNA yield is 1.5-2.5 mg.


The starting E. coli culture volume is 2.5-5 L of LB and the expected DNA yield is 7.5–10 mg.


It is important to consider the downstream applications of the plasmid DNA when choosing a purification method. For example, if the plasmid is to be used for  transfection  or  electroporation , a purification method that results in high purity and low endotoxin levels is desirable. Similarly, if the plasmid is to be used for sequencing or PCR, a purification method that results in high yield and minimal contaminants is desirable. [ 2 ]   However, multiple methods of nucleic acid purification exist. [ 23 ] [ 24 ] [ 25 ]  All work on the principle of generating conditions where either only the nucleic acid precipitates, or only other  biomolecules  precipitate, allowing the nucleic acid to be separated. [ 15 ] [ 23 ]


In high-throughput DNA extraction workflows, laboratory equipment such as  96 well plate template  can be utilized to efficiently process multiple samples in parallel. These templates allow for the automation of extraction protocols, significantly increasing the throughput of plasmid DNA isolation while maintaining consistency across large sample sets. When used in combination with automated liquid handling systems, a 96 well plate template helps streamline the process of extracting plasmid DNA from bacterial cultures, ensuring uniformity and reducing manual errors during the purification steps.


Ethanol precipitation  is a widely used method for purifying and concentrating  nucleic acids , including plasmid DNA. [ 26 ]  The basic principle of this method is that nucleic acids are insoluble in ethanol or isopropanol but soluble in water. Therefore, it works by using  ethanol  as an  antisolvent  of DNA, causing it to precipitate out of solution and then it can be collected by  centrifugation . The soluble fraction is discarded to remove other biomolecules. [ 27 ]


Spin column-based nucleic acid purification   is a method of purifying DNA, RNA or plasmid from a sample using a spin column filter. [ 25 ]  The method is based on the principle of selectively binding nucleic acids to a solid matrix in the spin column, while other contaminants, such as  proteins  and salts, are washed away. The conditions are then changed to elute the purified nucleic acid off the column using a suitable elution buffer. [ 25 ]


The basic principle of the  phenol-chloroform extraction  is that DNA and RNA are relatively insoluble in phenol and chloroform, while other cellular components are relatively soluble in these solvents. The addition of a  phenol / chloroform  mixture will dissolve protein and lipid contaminants, leaving the nucleic acids in the aqueous phase. It also denatures proteins, like  DNase , which is especially important if the plasmids are to be used for  enzyme  digestion. Otherwise, smearing may occur in enzyme restricted form of plasmid DNA. [ 24 ]


In beads-based extraction, addition of a mixture containing magnetic beads commonly made of  iron  ions binds to plasmid DNA, separating them from unwanted compounds by a magnetic rod or stand. [ 25 ]  The plasmid-bound beads are then released by removal of the magnetic field and extracted in an elution solution for down-stream experiments such as  transformation  or  restriction digestion . This form of miniprep can also be automated, which increases the conveniency while reducing mechanical error.


Polony sequencing  is an inexpensive but highly accurate  multiplex sequencing technique  that can be used to “read” millions of immobilized DNA sequences in parallel. This technique was first developed by  Dr. George Church 's group at  Harvard Medical School . Unlike other sequencing techniques,  Polony  sequencing technology is an open platform with freely downloadable, open source software and protocols. Also, the hardware of this technique can be easily set up with a commonly available  epifluorescence microscopy  and a computer-controlled flowcell/fluidics system.  Polony sequencing is generally performed on  paired-end tags  library that each molecule of DNA template is of 135 bp in length with two 17–18 bp paired genomic tags separated and flanked by common sequences. The current read length of this technique is 26 bases per amplicon and 13 bases per tag, leaving a gap of 4–5 bases in each tag.


The protocol of Polony sequencing can be broken into three main parts, which are the paired end-tag library construction, template amplification and DNA sequencing.


This protocol begins by randomly shearing the tested genomic DNA into a tight size distribution. The sheared DNA molecules are then subjected for the end repair and A-tailed treatment. The end repair treatment converts any damaged or incompatible protruding ends of DNA to 5’-phosphorylated and blunt-ended DNA, enabling immediate blunt-end ligation, while the A-tailing treatment adds an A to the 3’ end of the sheared DNA. DNA molecules with a length of 1 kb are selected by loading on the 6% TBE PAGE gel. In the next step, the DNA molecules are circularized with T-tailed 30 bp long synthetic oligonucleotides (T30), which contains two outward-facing MmeI recognition sites, and the resulting circularized DNA undergoes  rolling circle replication . The amplified circularized DNA molecules are then digested with MmeI (type IIs restriction endonuclease) which will cuts at a distance from its recognition site, releasing the T30 fragment flanked by 17–18 bp tags (≈70 bp in length). The paired-tag molecules need to be end-repaired prior to the ligation of ePCR (emulsion PCR) primer oligonucleotides (FDV2 and RDV2) to their both ends. The resulting 135 bp library molecules are size-selected and  nick translated . Lastly, amplify the 135 bp paired end-tag library molecules with  PCR  to increase the amount of library material and eliminate extraneous ligation products in a single step. The resulted DNA template consists of a 44 bp FDV sequence, a 17–18 bp proximal tag, the T30 sequence, a 17-18 bp distal tag, and a 25 bp RDV sequence.


The mono-sized, paramagnetic  streptavidin –coated beads are pre-loaded with dual biotin forward primer.  Streptavidin  has a very strong affinity for  biotin , thus the forward primer will bind firmly on the surface of the beads. Next, an aqueous phase is prepared with the pre-loaded beads, PCR mixture, forward and reverse primers, and the paired end-tag library. This is mixed and vortexed with an oil phase to create the emulsion. Ideally, each droplet of water in the oil emulsion has one bead and one molecule of template DNA, permitting millions of non-interacting amplification within a milliliter-scale volume by performing PCR.


After amplification, the emulsion from preceding step is broken using isopropanol and detergent buffer (10 mM Tris pH 7.5, 1 mM EDTA pH 8.0, 100 mM NaCl, 1% (v/v) Triton X‐100, 1% (w/v) SDS), following with a series of vortexing, centrifuging, and magnetic separation. The resulted solution is a suspension of empty, clonal and non-clonal beads, which arise from emulsion droplets that initially have zero, one or multiple DNA template molecules, respectively. The amplified bead could be enriched in the following step.


The enrichment of amplified beads is achieved through hybridization to a larger, low density, non-magnetic  polystyrene  beads that pre-loaded with a biotinylated capture oligonucleotides (DNA sequence that complementary to ePCR amplicon sequence). The mixture is then centrifuged to separate the amplified and capture beads complex from the unamplified beads. The amplified, capture bead complex has a lower density and thus will remain in the supernatant while the unamplified beads form a pellet. The supernatant is recovered and treated with NaOH which will break the complex. The paramagnetic amplified beads are separated from the non-magnetic capture beads by magnetic separation. This enrichment protocol is capable in enriching five times of amplified beads.


The purpose of bead capping is to attach a “capping” oligonucleotide to the 3’ end of both unextended forward ePCR primers and the RDV segment of template DNA. The cap that being use is an amino group that prevents fluorescent probes from ligating to these ends and at the same time, helping the subsequent coupling of template DNA to the aminosilanated flow cell coverslip.


First, the coverslips are washed and aminosilane-treated, enabling the subsequent covalent coupling of template DNA on it and eliminating any fluorescent contamination. The amplified, enriched beads are mixed with  acrylamide  and poured into a shallow mold formed by a  Teflon -masked microscope slide. Immediately, place the aminosilane-treated coverslip on top of the acrylamide gel and allow to polymerize for 45 minutes. Next, invert the slide/coverslip stack and remove the microscope slide from gel. The silane-treated coverslips will bind covalently to the gel while the Teflon on the surface of microscope slide will enable the better removal of slide from the acrylamide gel. The coverslips then bonded to the flow cell body and any unattached beads will be removed.


The biochemistry of Polony sequencing mainly relies on the  discriminatory capacities of ligases  and polymerases. First, a series of anchor primers are flowed through the cells and hybridize to the synthetic oligonucleotide sequences at the immediate 3’ or 5’ end of the 17-18 bp proximal or distal genomic DNA tags. Next, an enzymatic ligation reaction of the anchor primer to a population of  degenenerate   nonamers  that are labeled with fluorescent dyes is performed.
Differentially labeled nonamers:


The fluorophore-tagged nonamers  anneal  with differential success to the tag sequences according to a strategy similar to that of  degenerate primers , but instead of submission to polymerases, nonamers are selectively ligated onto adjoining DNA- the anchor primer.  The fixation of the fluorophore molecule provides a fluorescent signal that indicates whether there is an A, C, G, or T at the query position on the genomic DNA tag. After four-colour imaging, the anchor primer/nonamer complexes are stripped off and a new cycle is begun by replacing the anchor primer. A new mixture of the fluorescently tagged nonamers is introduced, for which the query position is shifted one base further into the genomic DNA tag.


Seven bases from the 5’ to 3’ direction and six bases from the 3’ end could be queried in this fashion. The ultimate result is a read length of 26 bases per run (13 bases from each of the paired tags) with a 4-base to 5-base gap in the middle of each tag.


The polony sequencing generates millions of 26 reads per run and this information needed to be normalized and converted to sequence. These can be done by the software that has been developed by Church Lab. All of the software is free and could be downloaded from the website. [ 1 ]


The sequencing instrument used in this technique could be set up by the commonly available fluorescence microscope and a computer controlled flowcell. The required instruments cost around US$130,000 in 2005. [ citation needed ]  A dedicated polony sequencing machine,  Polonator , was developed in 2009 and sold at US$170,000 by  Dover . [ 2 ] [ 3 ]  It and featured open source software, reagents, and protocols and was intended for use in the  Personal Genome Project . [ 4 ]


Polony sequencing allows for a high throughput and high consensus accuracies of DNA sequencing based on a commonly available, inexpensive instrument. Also, it is a very flexible technique that enables variable application including  BAC  (bacterial artificial chromosome) and bacterial genome resequencing, as well as SAGE (serial analysis of gene expression) tag and barcode sequencing. Furthermore, the polony sequencing technique is emphasized as an open system that shares everything including the software that have been developed, protocol and reagents.


However, although the raw data acquisition could be achieved as high as 786 gigabits but only 1 bit of information out of 10,000 bits collected is useful. Another challenge of this technique is the uniformity of the relative amplification of individual targets. The non-uniform amplification could lower the efficiency of sequencing and posted as the biggest obstacle in this technique.


Polony sequencing is a development of the  polony  technology from the late 1990s and 2000s. [ 5 ]  Methods were developed in 2003 to sequence  in situ  polonies using single-base extension which could achieve 5-6 bases reads. [ 6 ]  By 2005, these early attempts had been overhauled to develop the existing polony sequencing technology. [ 7 ]  The highly parallel sequencing-by-ligation concepts of polony sequencing has contributed the basis for later sequencing technologies such as  ABI Solid Sequencing .


Polymerase cycling assembly  (or  PCA , also known as  Assembly PCR ) is a method for the assembly of large  DNA   oligonucleotides  from shorter fragments.  The process uses the same technology as  PCR , but takes advantage of DNA hybridization and annealing as well as  DNA polymerase  to amplify a complete sequence of DNA in a precise order based on the single stranded oligonucleotides used in the process.  It thus allows for the production of  synthetic genes  and even entire synthetic  genomes .


Much like how primers are designed such that there is a forward primer and a reverse primer capable of allowing DNA polymerase to fill the entire template sequence, PCA uses the same technology but with multiple oligonucleotides. While in PCR the customary size of oligonucleotides used is 18 base pairs, in PCA lengths of up to 50 are used to ensure uniqueness and correct hybridization.


Each oligonucleotide is designed to be either part of the top or bottom strand of the target sequence. As well as the basic requirement of having to be able to tile the entire target sequence, these oligonucleotides must also have the usual properties of similar melting temperatures, hairpin free, and not too GC rich to avoid the same complications as PCR.


During the polymerase cycles, the oligonucleotides anneal to complementary fragments and then are filled in by polymerase. Each cycle thus increases the length of various fragments randomly depending on which oligonucleotides find each other. It is critical that there is complementarity between all the fragments in some way or a final complete sequence will not be produced as polymerase requires a template to follow.


After this initial construction phase, additional primers encompassing both ends are added to perform a regular PCR reaction, amplifying the target sequence away from all the shorter incomplete fragments. A gel purification can then be used to identify and isolate the complete sequence.


A typical reaction consists of oligonucleotides ~50 base pairs long each overlapping by about 20 base pairs. The reaction  with all the oligonucleotides is then carried out for ~30 cycles followed by an additional 23 cycles with the end primers. [ 1 ]


A modification of this method,  Gibson assembly , described by Gibson et al. allows for single-step isothermal assembly of DNA with up to several hundreds  kb . By using T5 exonuclease to 'chew back' complementary ends, an overlap of about 40bp can be created. The reaction takes place at 50 °C, a temperature where the T5 exonuclease is unstable. After a short timestep it is degraded, the overlaps can anneal and be ligated. [ 2 ]   Cambridge University   IGEM  team made a video describing the process. [ 3 ]  Ligation independent cloning (LIC) is a new variant of the method for compiling several DNA pieces together and needing only exonuclease enzyme for the reaction.


Pulsed-field gel electrophoresis  ( PFGE ) is a technique used for the separation of large  DNA   molecules  by applying an  electric field  that periodically changes direction to a  gel  matrix. [ 1 ] [ 2 ]  Unlike standard  agarose gel electrophoresis , which can separate DNA fragments of up to 50 kb, PFGE resolves fragments up to 10 Mb. [ 1 ]  This allows for the direct analysis of genomic DNA. [ 2 ]


In 1984, David C. Schwartz and  Charles Cantor  published the first successful application of alternating electric fields for the separation of large DNA molecules. [ 3 ] [ 4 ]  This technique, which they named PFGE, resulted in the development of several variations, including Orthogonal Field Alternation Gel Electrophoresis (OFAGE), Transverse Alternating Field Electrophoresis (TAFE), Field-Inversion Gel Electrophoresis (FIGE), and Clamped Homogeneous Electric Fields (CHEF), among others. [ 3 ]


The procedure for PFGE is similar to that of standard  agarose gel electrophoresis , with the main exception being the application of the electric current. Generally, in PFGE electrophoresis chambers, the voltage periodically switches between three directions: one along the central axis, and two at a 60 degree angle along each side. [ 5 ]  The application of the voltage can change depending on the variation of PFGE used. [ 6 ] [ 7 ]


PFGE may be used for  genotyping  or  genetic fingerprinting . It has commonly been considered a  gold standard  in  epidemiological  studies of pathogenic organisms for several decades. For instance, subtyping bacterial isolates with this method has made it easier to discriminate among strains of  Listeria monocytogenes ,  Lactococcus garvieae [ 8 ]  and some clinical isolates of  Bacillus cereus   [ 9 ]  group isolated from diseases aquatic organisms and thus to link environmental or food isolates with clinical infections. It is now in the process of being superseded by next generation sequencing methods. [ 10 ]


The  Punnett square  is a square diagram that is used to predict the genotypes of a particular cross or breeding experiment. It is named after  Reginald C. Punnett , who devised the approach in 1905. [ 3 ] [ 4 ] [ 5 ] [ 6 ] [ 7 ] [ 8 ]  The diagram is used by  biologists  to determine the  probability  of an offspring having a particular  genotype . The Punnett square is a tabular summary of possible combinations of maternal  alleles  with paternal alleles. [ 9 ]  These tables can be used to examine the genotypical outcome probabilities of the offspring of a single trait (allele), or when crossing multiple traits from the parents.


The Punnett square is a visual representation of  Mendelian inheritance , a fundamental concept in genetics discovered by  Gregor Mendel . [ 10 ]  For multiple traits, using the "forked-line method" is typically much easier than the Punnett square. Phenotypes may be predicted with at least better-than-chance accuracy using a Punnett square, but the phenotype that may appear in the presence of a given genotype can in some instances be influenced by many other factors, as when  polygenic inheritance  and/or  epigenetics  are at work.


Zygosity  refers to the grade of similarity between the  alleles  that determine one specific  trait  in an  organism . In its simplest form, a pair of alleles can be either  homozygous  or  heterozygous . Homozygosity, with  homo  relating to  same  while  zygous  pertains to a  zygote , is seen when a combination of either two  dominant  or two  recessive  alleles code for the same trait. Recessive are always lowercase letters. For example, using 'A' as the representative character for each allele, a  homozygous dominant  pair's  genotype  would be depicted as 'AA', while  homozygous recessive  is shown as 'aa'. Heterozygosity, with  hetero  associated with  different , can only be 'Aa' (the capital letter is always presented first by convention). The  phenotype  of a homozygous dominant pair is 'A', or  dominant , while the opposite is true for homozygous  recessive . Heterozygous pairs always have a dominant phenotype. [ 11 ]  To a lesser degree,  hemizygosity [ 12 ]  and nullizygosity [ 13 ]  can also be seen in gene pairs.


"Mono-" means "one"; this cross indicates that the examination of a single trait. This could mean (for example) eye color. Each genetic locus is always represented by two letters. So in the case of eye color, say "B = Brown eyes" and "b = green eyes".
In this example, both parents have the  genotype   Bb . For the example of eye color, this would mean they both have brown eyes. They can produce  gametes  that contain either the  B  or the  b  allele. (It is conventional in  genetics  to use capital letters to indicate  dominant  alleles and lower-case letters to indicate  recessive  alleles.) The probability of an individual offspring's having the genotype  BB  is 25%,  Bb  is 50%, and  bb  is 25%. The ratio of the phenotypes is 3:1, typical for a  monohybrid cross . When assessing phenotype from this,  "3" of the offspring have "Brown" eyes and only one offspring has "green" eyes. (3 are "B?" and 1 is "bb")


The way in which the  B  and  b  alleles interact with each other to affect the appearance of the offspring depends on how the gene products ( proteins ) interact (see  Mendelian inheritance ). This can include lethal effects and  epistasis  (where one allele masks another, regardless of dominant or recessive status).


More complicated crosses can be made by looking at two or more genes. The Punnett square works, however, only if the genes are  independent  of each other, which means that having a particular allele of gene "A" does not alter the probability of possessing an allele of gene "B". This is equivalent to stating that the genes are not  linked , so that the two genes do not tend to sort together during meiosis.


The following example illustrates a  dihybrid cross  between two double-heterozygote pea plants.  R  represents the dominant allele for shape (round), while  r  represents the recessive allele (wrinkled).  A  represents the dominant allele for color (yellow), while  a  represents the recessive allele (green). If each plant has the genotype  RrAa , and since the alleles for shape and color genes are independent, then they can produce four types of gametes with all possible combinations:  RA ,  Ra ,  rA , and  ra .


Since dominant traits mask recessive traits (assuming no epistasis), there are nine combinations that have the phenotype round yellow, three that are round green, three that are wrinkled yellow, and one that is wrinkled green. The ratio 9:3:3:1 is the expected outcome when crossing two double-heterozygous parents with unlinked genes. Any other ratio indicates that something else has occurred (such as lethal alleles, epistasis, linked genes, etc.).


The forked-line method (also known as the tree method and the branching system) can also solve dihybrid and multi-hybrid crosses. A problem is converted to a series of monohybrid crosses, and the results are combined in a tree. However, a tree produces the same result as a Punnett square in less time and with more clarity. The example below assesses another double-heterozygote cross using RrYy x RrYy. As stated above, the phenotypic ratio is expected to be 9:3:3:1 if crossing unlinked genes from two double-heterozygotes. The genotypic ratio was obtained in the diagram below, this diagram will have more branches than if only analyzing for phenotypic ratio.


Rapid amplification of cDNA ends  ( RACE ) is a technique used in  molecular biology  to obtain the full length sequence of an  RNA  transcript found within a cell. RACE results in the production of a  cDNA  copy of the RNA sequence of interest, produced through  reverse transcription , followed by  PCR  amplification of the cDNA copies (see  RT-PCR ). The amplified cDNA copies are then sequenced and, if long enough, should map to a unique genomic region. RACE is commonly followed up by cloning before sequencing of what was originally individual RNA molecules. A more high-throughput alternative which is useful for identification of novel transcript structures, is to sequence the RACE-products by next generation sequencing technologies.


RACE can provide the sequence of an RNA transcript from a small known sequence within the transcript to the  5' end  (5' RACE-PCR) or  3' end  (3' RACE-PCR) of the RNA. This technique is sometimes called  one-sided PCR  or  anchored PCR .


The first step in RACE is to use reverse transcription to produce a  cDNA  copy of a region of the RNA transcript.  In this process, an unknown end portion of a transcript is copied using a known sequence from the center of the transcript. The copied region is bounded by the known sequence, at either the 5' or 3' end.


The protocols for 5' or 3' RACES differ slightly.  5' RACE-PCR begins using mRNA as a template for a first round of cDNA synthesis (or  reverse transcription ) reaction using an  anti-sense  (reverse)  oligonucleotide  primer that recognizes a known sequence in the middle of the gene of interest; the  primer  is called a  gene specific primer  (GSP). The primer binds to the mRNA, and the enzyme reverse transcriptase adds base pairs to the 3' end of the primer to generate a specific single-stranded cDNA product; this is the reverse complement of the mRNA. Following cDNA synthesis, the  enzyme   terminal deoxynucleotidyl transferase  (TdT) is used to add a string of identical  nucleotides , known as a homopolymeric tail, to the 3' end of the cDNA. (There are some other ways to add the 3'-terminal sequence for the first strand of the de novo cDNA synthesis which are much more efficient than homopolymeric tailing, but the sense of the method remains the same).
 PCR  is then carried out, which uses a second anti-sense gene specific primer (GSP2) that binds to the known sequence, and a sense (forward) universal primer (UP) that binds the homopolymeric tail added to the 3' ends of the cDNAs to amplify a cDNA product from the 5' end.


3' RACE-PCR uses the natural  polyA tail  that exists at the 3' end of all eukaryotic mRNAs for priming during reverse transcription, so this method does not require the addition of nucleotides by TdT.  cDNAs are generated using an  Oligo-dT -adaptor primer (a primer with a short sequence of deoxy-thymine nucleotides) that complements the  polyA  stretch and adds a special adaptor sequence to the 5' end of each cDNA.  PCR is then used to amplify 3' cDNA from a known region using a sense GSP, and an anti-sense primer complementary to the adaptor sequence.


The  cDNA  molecules generated by RACE can be sequenced using  high-throughput sequencing  technologies (also called, RACE-seq). High-throughput sequencing characterization of RACE fragments is highly time-efficient, more sensitive, less costly and technically feasible compared to traditional characterization of RACE fragments with  molecular cloning  followed by  Sanger sequencing  of a few clones.


RACE can be used to amplify unknown 5' (5'-RACE) or 3' (3'-RACE) parts of RNA molecules where part of the RNA sequence is known and targeted by a gene-specific primer. Combined with high-throughput sequencing for characterization of these amplified RACE products, it is possible to apply the approach to characterize any types of coding or non-coding RNA-molecules.


The idea of combining RACE with high-throughput sequencing was first introduced in 2009 as Deep-RACE to perform mapping of  Transcription start sites  (TSS) of 17 genes in a single cell-line. [ 1 ]  For example, In a study from 2014 to accurately map cleavage sites of target RNA directed by synthetic  siRNAs , the approach was first named RACE-seq. [ 2 ]  Further, the methodology was used to characterize full-length unknown parts of novel transcripts and  fusion transcripts  in  colorectal cancer . [ 3 ]
In another study aiming to characterize unknown transcript structures of  lncRNAs , RACE was used in combination with semi-long  454 sequencing . [ 4 ]


Recombinant DNA  ( rDNA ) molecules are  DNA  molecules formed by laboratory methods of  genetic recombination  (such as  molecular cloning ) that bring together genetic material from multiple sources, creating  sequences  that would not otherwise be found in the  genome .


Recombinant DNA is the general name for a piece of DNA that has been created by combining two or more fragments from different sources. Recombinant DNA is possible because DNA molecules from all organisms share the same chemical structure, differing only in the  nucleotide  sequence. Recombinant DNA molecules are sometimes called  chimeric DNA  because they can be made of material from two different species like the mythical  chimera . rDNA technology uses  palindromic sequences  and leads to the production of  sticky and blunt ends .