Cre recombinase consists of 343  amino acids  that form two distinct domains. The  amino terminal  domain encompasses residues 20–129 and this domain contains 5  alpha helical  segments linked by a series of short loops. Helices A & E are involved in the formation of the recombinase tetramer with the C terminus region of helix E known to form contacts with the C terminal domain of adjacent subunits. Helices B & D form direct contacts with the major groove of the loxP DNA. These two helices are thought to make three direct contacts to DNA bases at the loxP site. The  carboxy terminal  domain of the enzyme consists of amino acids 132–341 and it harbours the  active site  of the enzyme. The overall structure of this domain shares a great deal of structural resemblance to the  catalytic domain  of other enzymes of the same family such as λ Integrase and HP1 Integrase. This domain is predominantly helical in structure with 9 distinct helices (F−N). The terminal helix (N) protrudes from the main body of the carboxy domain and this helix is reputed to play a role in mediating interactions with other subunits. Crystal structures demonstrate that this terminal N helix buries its hydrophobic surface into an acceptor pocket of an adjacent Cre subunit. [ 6 ]


The effect of the two-domain structure is to form a C-shaped clamp that grasps the DNA from opposite sides. [ 3 ]


The  active site  of the Cre enzyme consists of the conserved  catalytic triad  residues  Arg  173,  His  289,  Arg  292 as well as the conserved  nucleophilic  residues  Tyr  324 and  Trp  315. Unlike some recombinase enzymes such as Flp recombinase, Cre does not form a shared active site between separate subunits and all the residues that contribute to the active site are found on a single subunit. Consequently, when two Cre molecules bind at a single loxP site two active sites are present. Cre mediated recombination requires the formation of a synapse in which two Cre-LoxP complexes associate to form what is known as the synapse tetramer in which 4 distinct active sites are present. [ 6 ]
 Tyr  324 acts as a  nucleophile  to form a covalent 3’-phosphotyrosine linkage to the DNA substrate. The  scissile  phosphate (phosphate targeted for nucleophilic attack at the cleavage site) is coordinated by the side chains of the 3  amino acid  residues of the  catalytic triad  ( Arg  173,  His  289 &  Trp  315). The  indole   nitrogen  of  tryptophan  315 also forms a  hydrogen bond  to this scissile phosphate. (n.b A  Histidine  occupies this site in other tyrosine recombinase family members and performs the same function). This reaction cleaves the DNA and frees a 5’ hydroxyl group. This process occurs in the active site of two out of the four recombinase subunits present at the synapse tetramer. If the 5’ hydroxyl groups attack the 3’-phosphotyrosine linkage one pair of the DNA strands will exchange to form a  Holliday junction  intermediate. [ 3 ]


Cre recombinase plays important roles in the  life cycle  of the  P1 bacteriophage . Upon infection of a cell the Cre-loxP system is used to cause circularization of the P1 DNA. In addition to this Cre is also used to resolve dimeric lysogenic P1 DNA that forms during the cell division of the phage. [ 7 ]


Inducible Cre activation is achieved using CreER (estrogen receptor) variant, which is only activated after delivery of  tamoxifen . [ 8 ]  This is done through the fusion of a mutated ligand binding domain of the estrogen receptor to the Cre recombinase, resulting in Cre becoming specifically activated by tamoxifen. In the absence of tamoxifen, CreER will result in the shuttling of the mutated recombinase into the cytoplasm. The protein will stay in this location in its inactivated state until tamoxifen is given. Once tamoxifen is introduced, it is metabolized into 4-hydroxytamoxifen, which then binds to the ER and results in the translocation of the CreER into the nucleus, where it is then able to cleave the lox sites. [ 9 ]  Importantly, sometimes fluorescent reporters can be activated in the absence of tamoxifen, due to leakage of a few Cre recombinase molecules into the nucleus which, in combination with very sensitive reporters, results in unintended cell labelling. [ 10 ]  CreER(T2) was developed to minimize tamoxifen-independent recombination and maximize tamoxifen-sensitivity.


In recent years, Cre recombinase has been improved with conversion to preferred mammalian  codons , the removal of reported cryptic  splice sites , an altered  stop codon , and reduced  CpG content  to reduce the risk of epigenetic  silencing  in  mammals . [ 11 ]  A number of mutants with enhanced accuracy have also been identified. [ 12 ]


Cre-Lox recombination  is a  site-specific recombinase technology , used to carry out  deletions ,  insertions ,  translocations  and  inversions  at specific sites in the DNA of cells. It allows the DNA modification to be targeted to a specific cell type or be triggered by a specific external stimulus. It is implemented both in eukaryotic and prokaryotic systems. The Cre-lox recombination system has been particularly useful to help neuroscientists to study the brain in which complex cell types and neural circuits come together to generate cognition and behaviors. NIH Blueprint for Neuroscience Research has created several hundreds of Cre driver mouse lines which are currently used by the worldwide neuroscience community.


An important application of the Cre-lox system is excision of selectable markers in gene replacement. Commonly used gene replacement strategies introduce selectable markers into the genome to facilitate selection of genetic mutations that may cause growth retardation. However, marker expression can have polar effects on the expression of upstream and downstream genes. Removal of selectable markers from the genome by Cre-lox recombination is an elegant and efficient way to circumvent this problem and is therefore widely used in plants, mouse cell lines, yeast, etc. [ 1 ]


The system consists of a single enzyme,  Cre recombinase , that  recombines  a pair of short target sequences called the  Lox  sequences. This system can be implemented without inserting any extra supporting proteins or sequences. The Cre enzyme and the original  Lox  site called the  LoxP  sequence are derived from  bacteriophage   P1 .


Placing Lox sequences appropriately allows genes to be activated, repressed, or exchanged for other genes. At a DNA level many types of manipulations can be carried out. The activity of the Cre enzyme can be controlled so that it is expressed in a particular cell type or triggered by an external stimulus like a chemical signal or a heat shock. These targeted DNA changes are useful in  cell lineage  tracing and when mutants are lethal if expressed globally.


The Cre-Lox system is very similar in action and in usage to the  FLP-FRT recombination  system. [ 2 ]


Cre-Lox recombination is a special type of  site-specific recombination  developed by Dr.  Brian Sauer  and patented by  DuPont  that operated in both mitotic and non-mitotic cells, and was initially used in activating gene expression in mammalian cell lines. [ 3 ] [ 4 ] [ 5 ]   Subsequently, researchers in the laboratory of Dr.  Jamey Marth  demonstrated that Cre-Lox recombination could be used to delete loxP-flanked chromosomal DNA sequences at high efficiency in specific developing T-cells of transgenic animals, with the authors proposing that this approach could be used to define endogenous gene function in specific cell types, indelibly mark progenitors in cell fate determination studies, induce specific chromosomal rearrangements for biological and disease modeling, and determine the roles of early genetic lesions in disease (and phenotype) maintenance. [ 6 ]


Shortly thereafter, researchers in the laboratory of Prof. Klaus Rajewsky reported the production of pluripotent embryonic stem cells bearing a targeted loxP-flanked (floxed) DNA polymerase gene. [ 7 ]   Combining these advances in collaboration, the laboratories of Drs. Marth and Rajewsky reported in 1994 that Cre-lox recombination could be used for conditional gene targeting. [ 8 ]  They observed ≈50% of the DNA polymerase beta gene was deleted in T cells based on DNA blotting. It was unclear whether only one allele in each T-cell or 50% of T cells had 100% deletion in both alleles.  Researchers have since reported more efficient Cre-Lox conditional gene mutagenesis in the developing T cells by the Marth laboratory in 1995. [ 9 ]  Incomplete deletion by  Cre recombinase  is not uncommon in cells when two copies of floxed sequences exist, and allows the formation and study of chimeric tissues.  All cell types tested in mice have been shown to undergo transgenic Cre recombination.


Independently,  Joe Z. Tsien  has pioneered the use of Cre-loxP system for cell type- and region-specific gene manipulation in the adult brain where hundreds of distinct neuron types may exist and nearly all neurons in the adult brain are known to be post-mitotic. [ 10 ]  Tsien and his colleagues demonstrated Cre-mediated recombination can occur in the post-mitotic  pyramidal neurons  in the adult mouse forebrain. [ 11 ]


These developments have led to a widespread use of conditional mutagenesis in biomedical research, spanning many disciplines in which it becomes a powerful platform for determining gene function in specific cell types and at specific developmental times.  In particular, the clear demonstration of its usefulness in precisely defining the complex relationship between specific cells/circuits and behaviors for brain research, [ 12 ]  has promoted the NIH to initiate the NIH Blueprint for Neuroscience Research Cre-driver mouse projects in early 2000. [ 13 ] [ 14 ]   To date, NIH Blueprint for Neuroscience Research Cre projects have created several hundreds of Cre driver mouse lines which are currently used by the worldwide neuroscience community.


Cre-Lox recombination involves the targeting of a specific sequence of  DNA  and splicing it with the help of an enzyme called  Cre recombinase . Cre-Lox recombination is commonly used to circumvent embryonic lethality caused by systemic inactivation of many genes. [ 15 ] [ 16 ]   As of February 2019, Cre–Lox recombination is a powerful tool and is used in transgenic animal modeling to link genotypes to phenotypes. [ 12 ] [ 17 ] [ 18 ]


The Cre-lox system is used as a genetic tool to control site specific  recombination  events in genomic DNA. This system has allowed researchers to manipulate a variety of genetically modified organisms to control gene expression, delete undesired DNA sequences and modify chromosome architecture.


The  Cre  protein is a site-specific DNA recombinase that can catalyse the recombination of DNA between specific sites in a DNA molecule. These sites, known as  loxP  sequences, contain specific binding sites for Cre that surround a directional core sequence where recombination can occur.


When cells that have  loxP  sites in their genome express Cre, a recombination event can occur between the  loxP  sites. Cre recombinase proteins bind to the first and last 13 bp regions of a lox site forming a  dimer . This dimer then binds to a dimer on another lox site to form a  tetramer . Lox sites are directional and the two sites joined by the tetramer are parallel in orientation.  The double stranded DNA is cut at both  loxP  sites by the Cre protein. The strands are then rejoined with  DNA ligase  in a quick and efficient process. The result of recombination depends on the orientation of the  loxP  sites. For two lox sites on the same chromosome arm, inverted  loxP  sites will cause an inversion of the intervening DNA, while a direct repeat of  loxP  sites will cause a deletion event. If  loxP  sites are on different chromosomes it is possible for  translocation  events to be catalysed by Cre induced recombination. Two  plasmids  can be joined using the variant lox sites 71 and 66. [ 19 ]


Cre recombinase can be synthesized by the corresponding gene under the direction of cell-specific promoters, including promoters under the control of doxycycline. An additional level of control can be achieved by using his Cre recombinase, engineered to reversibly activate in the presence of the estrogen analogue 4-hydroxy tamoxifen. This provides the advantage that the Cre recombinase is active for a short time. This prevents non-specific actions of Cre recombinase. The Cre recombinase can recognize cryptic sites in the host genome and induce unauthorized recombination, damaging host DNA. This tool is suitable for deleting antibiotic resistance genes, but above all it allows conditional knockouts that can be induced at specific times in the cell type of choice. Models thus obtained are more likely to mimic the physiological situation. [ 20 ]


The Cre protein (encoded by the locus originally named as "Causes recombination", with "Cyclization recombinase" being found in some references) [ 21 ] [ 22 ]   consists of 4 subunits and two domains: The larger carboxyl ( C-terminal ) domain, and smaller amino ( N-terminal ) domain. The total protein has 343  amino acids . The C domain is similar in structure to the domain in the  Integrase  family of enzymes isolated from  lambda phage . This is also the  catalytic site  of the enzyme.


loxP  (locus of X-over P1) is a site on the bacteriophage P1 consisting of 34  bp . The site includes an asymmetric 8 bp sequence, variable except for the middle two bases, in between two sets of symmetric, 13 bp sequences. The exact sequence is given below; 'N' indicates bases which may vary, and lowercase letters indicate bases that have been mutated from the wild-type. The 13 bp sequences are palindromic but the 8 bp spacer is not, thus giving the loxP sequence a certain direction. Usually loxP sites come in pairs for genetic manipulation. If the two loxP sites are in the same orientation, the floxed sequence (sequence flanked by two loxP sites) is excised; however if the two loxP sites are in the opposite orientation, the floxed sequence is inverted. If there exists a floxed donor sequence, the donor sequence can be swapped with the original sequence. This technique is called  recombinase-mediated cassette exchange  and is a very convenient and time-saving way for genetic manipulation. The caveat, however, is that the recombination reaction can happen backwards, rendering cassette exchange inefficient. In addition, sequence excision can happen  in trans  instead of a  in cis  cassette exchange event. The loxP mutants are created to avoid these problems. [ 23 ]


During genetic recombination, a  Holliday junction  is formed between the two strands of DNA and a double-stranded break in a DNA molecule leaves a 3’OH end exposed. This reaction is aided with the endonuclease activity of an enzyme. 5’ Phosphate ends are usually the substrates for this reaction, thus extended 3’ regions remain. This 3’ OH group is highly unstable, and the strand on which it is present must find its complement. Since homologous recombination occurs after DNA replication, two strands of DNA are available, and thus, the 3’ OH group must pair with its complement, and it does so, with an intact strand on the other duplex. Now, one point of crossover has occurred, which is what is called a Holliday Intermediate.


The 3’OH end is elongated (that is, bases are added) with the help of DNA Polymerase. The pairing of opposite strands is what constitutes the crossing-over or Recombination event, which is common to all living organisms, since the genetic material on one strand of one duplex has paired with one strand of another duplex, and has been elongated by DNA polymerase. Further cleavage of Holliday Intermediates results in formation of Hybrid DNA.


This further cleavage or ‘resolvation’ is done by a special group of enzymes called Resolvases. RuvC is just one of these Resolvases that have been isolated in bacteria and yeast.


For many years, it was thought that when the Holliday junction intermediate was formed, the branch point of the junction (where the strands cross over) would be located at the first cleavage site. Migration of the branch point to the second cleavage site would then somehow trigger the second half of the pathway. This model provided convenient explanation for the strict requirement for homology between recombining sites, since branch migration would stall at a mismatch and would not allow the second strand exchange to occur. In more recent years, however, this view has been challenged, and most of the current models for Int, Xer, and Flp recombination involve only limited branch migration (1–3 base pairs of the Holliday intermediate), coupled to an isomerisation event that is responsible for switching the strand cleavage specificity.


Site-specific recombination (SSR) involves specific sites for the catalyzing action of special enzymes called recombinases. Cre, or cyclic recombinase, is one such enzyme. Site-specific recombination is, thus, the enzyme-mediated cleavage and ligation of two defined deoxynucleotide sequences.


A number of conserved site-specific recombination systems have been described in both prokaryotic and eukaryotic organisms. In general, these systems use one or more proteins and act on unique asymmetric DNA sequences. The products of the recombination event depend on the relative orientation of these asymmetric sequences. Many other proteins apart from the recombinase are involved in regulating the reaction. During site-specific DNA recombination, which brings about genetic rearrangement in processes such as viral integration and excision and chromosomal segregation, these recombinase enzymes recognize specific DNA sequences and catalyse the reciprocal exchange of DNA strands between these sites.


Initiation of site-specific recombination begins with the binding of recombination proteins to their respective DNA targets. A separate recombinase recognizes and binds to each of two recombination sites on two different DNA molecules or within the same DNA strand. At the given specific site on the DNA, the hydroxyl group of the tyrosine in the recombinase attacks a phosphate group in the DNA backbone using a direct  transesterification  mechanism. This reaction links the recombinase protein to the DNA via a phospho-tyrosine linkage. This conserves the energy of the phosphodiester bond, allowing the reaction to be reversed without the involvement of a high-energy cofactor.


Cleavage on the other strand also causes a phospho-tyrosine bond between DNA and the enzyme. At both of the DNA duplexes, the bonding of the phosphate group to tyrosine residues leave a 3’ OH group free in the DNA backbone. In fact, the enzyme-DNA complex is an intermediate stage, which is followed by the ligation of the 3’ OH group of one DNA strand to the 5’ phosphate group of the other DNA strand, which is covalently bonded to the tyrosine residue; that is, the covalent linkage between 5’ end and tyrosine residue is broken. This reaction synthesizes the  Holliday junction  discussed earlier.


In this fashion, opposite DNA strands are joined. Subsequent cleavage and rejoining cause DNA strands to exchange their segments. Protein-protein interactions drive and direct strand exchange. Energy is not compromised, since the protein-DNA linkage makes up for the loss of the phosphodiester bond, which occurred during cleavage.


Site-specific recombination is also an important process that viruses, such as bacteriophages, adopt to integrate their genetic material into the infected host. The virus, called a  prophage  in such a state, accomplishes this via integration and excision. The points where the integration and excision reactions occur are called the attachment (att) sites.  An attP site on the phage exchanges segments with an attB site on the bacterial DNA.  Thus, these are site-specific, occurring only at the respective att sites. The  integrase  class of enzymes catalyse this particular reaction.


Two factors have been shown to affect the efficiency of Cre's excision on the lox pair.  First, the nucleotide sequence identity in the spacer region of lox site. Engineered lox variants which differ on the spacer region tend to have varied but generally lower recombination efficiency compared to wildtype loxP, presumably through affecting the formation and resolution of recombination intermediate. [ 26 ]


Another factor is the length of DNA between the lox pair. Increasing the length of DNA leads to decreased efficiency of Cre/lox recombination possibly through regulating the dynamics of the reaction. [ 27 ] [ 28 ] [ 29 ]  Genetic location of the floxed sequence affects recombination efficiency as well probably by influencing the availability of DNA by  Cre recombinase . [ 29 ]  The choice of Cre driver is also important as low expression of  Cre recombinase  tends to result in non-parallel recombination. Non-parallel recombination is especially problematic in a  fate mapping  scenario where one recombination event is designed to manipulate the gene under study and the other recombination event is necessary for activating a reporter gene (usually encoding a fluorescent protein) for  cell lineage  tracing. [ 29 ]  Failure to activate both recombination events simultaneously confounds the interpretation of cell  fate mapping  results.


Inducible Cre activation is achieved using CreER (estrogen receptor) variant, which is only activated after delivery of  tamoxifen . [ 30 ]  This is done through the fusion of a mutated ligand binding domain of the estrogen receptor to the Cre recombinase, resulting in Cre becoming specifically activated by tamoxifen. In the absence of tamoxifen, CreER will result in the shuttling of the mutated recombinase into the cytoplasm. The protein will stay in this location in its inactivated state until tamoxifen is given. Once tamoxifen is introduced, it is metabolized into 4-hydroxytamoxifen, which then binds to the ER and results in the translocation of the CreER into the nucleus, where it is then able to cleave the lox sites. [ 31 ]  Importantly, sometimes fluorescent reporters can be activated in the absence of tamoxifen, due to leakage of a few Cre recombinase molecules into the nucleus which, in combination with very sensitive reporters, results in unintended cell labelling. [ 32 ]  CreER(T2) was developed to minimize tamoxifen-independent recombination and maximize tamoxifen-sensitivity.


Cells alter their phenotype in response to numerous environmental stimuli and can lose the expression of genes typically used to mark their identity, making it difficult to research the contribution of certain cell types to disease. Therefore, researchers often use transgenic mice expressing CreER t2  recombinase induced by tamoxifen administration, under the control of a promoter of a gene that marks the specific cell type of interest, with a Cre-dependent fluorescent protein reporter. The Cre recombinase is fused to a mutant form of the oestrogen receptor, which binds the synthetic oestrogen 4-hydroxytamoxifen instead of its natural ligand 17β-estradiol. CreER(T2) resides within the cytoplasm and can only translocate to the nucleus following tamoxifen administration, allowing tight temporal control of recombination. The fluorescent reporter cassette  will contain a promoter to permit high expression of the fluorescent transgene reporter (e.g. a CAG promoter) and a loxP flanked stop cassette, ensuring the expression of the transgene is Cre-recombinase dependent and the reporter sequence. Upon Cre driven recombination, the stop cassette is excised, allowing reporter genes to express specifically in cells in which the Cre expression is being driven by the cell-specific marker promoter. Since removal of the stop cassette is permanent, the reporter genes are expressed in all the progeny produced by the initial cells where the Cre was once activated. Such conditional lineage tracing has proved to be extremely useful to efficiently and specifically identify vascular smooth muscle cells (VSMCs) and VSMC-derived cells and has been used to test effects on VSMC and VSMC-derived cells  in vivo. [ 33 ] [ 34 ] [ 35 ] [ 36 ] [ 37 ] [ 38 ]


The  P1 phage  is a  temperate  phage that causes either a  lysogenic  or  lytic cycle  when it infects a bacterium. In its lytic state, once its viral genome is injected into the host cell, viral proteins are produced, virions are assembled, and the host cell is lysed to release the phages, continuing the cycle. In the lysogenic cycle the phage genome replicates with the rest of the bacterial genome and is transmitted to daughter cells at each subsequent cell division. It can transition to the lytic cycle by a later event such as UV radiation or starvation.


Phages like the  lambda phage  use their site specific recombinases to integrate their DNA into the host genome during lysogeny. P1 phage DNA on the other hand, exists as a  plasmid  in the host. The Cre-lox system serves several functions in the phage: it circularizes the phage DNA into a plasmid, separates interlinked plasmid rings so they are passed to both daughter bacteria equally and may help maintain copy numbers through an alternative means of replication. [ 39 ]


The P1 phage DNA when released into the host from the virion is in the form of a linear double stranded DNA molecule. The Cre enzyme targets loxP sites at the ends of this molecule and cyclises the genome. This can also take place in the absence of the Cre lox system [ 40 ]  with the help of other bacterial and viral proteins. The P1 plasmid is relatively large (≈90Kbp) and hence exists in a low copy number - usually one per cell. If the two daughter plasmids get interlinked one of the daughter cells of the host will lose the plasmid. The Cre-lox recombination system prevents these situations by unlinking the rings of DNA by carrying out two recombination events (linked rings -> single fused ring -> two unlinked rings). It is also proposed that rolling circle replication followed by recombination will allow the plasmid to increase its copy number when certain regulators (repA) are limiting. [ 39 ]


A classical strategy for generating gene deletion variants is based on double cross-integration of non-replicating vectors into the genome. Furthermore, recombination systems such as Cre-lox are widely used, mostly in eukaryotes. The versatile properties of Cre recombinase make it ideal for use in many genetic engineering strategies. As such, the Cre lox system has been used in a wide variety of eukaryotes, including plants. [ 41 ]


Multiple variants of loxP, [ 42 ]  in particular lox2272 and loxN, have been used by researchers with the combination of different Cre actions (transient or constitutive) to create a " Brainbow " system that allows multi-colouring of mice's brain with four fluorescent proteins.


Another report using two lox variants pair but through regulating the length of DNA in one pair results in stochastic gene activation with regulated level of sparseness. [ 27 ]


In  genetics ,  crosslinking of DNA  occurs when various exogenous or endogenous agents react with two  nucleotides  of  DNA , forming a covalent linkage between them. This crosslink can occur within the same strand (intrastrand) or between opposite strands of double-stranded DNA (interstrand). These adducts interfere with cellular metabolism, such as  DNA replication  and  transcription , triggering  cell death . These crosslinks can, however, be  repaired  through excision or recombination pathways.


DNA crosslinking also has useful merit in chemotherapy and targeting cancerous cells for  apoptosis , [ 1 ]  as well as in understanding how proteins interact with DNA.


Many characterized crosslinking agents have two independently reactive groups within the same molecule, each of which is able to bind with a nucleotide residue of DNA. These agents are separated based upon their source of origin and labeled either as exogenous or endogenous. Exogenous crosslinking agents are chemicals and compounds, both natural and synthetic, that stem from environmental exposures such as pharmaceuticals and cigarette smoke or automotive exhaust. Endogenous crosslinking agents are compounds and metabolites that are introduced from cellular or biochemical pathways within a cell or organism.


Interstrand;  5'-GNG


Crosslinked DNA is repaired in cells by a combination of enzymes and other factors from the  nucleotide excision repair  (NER) pathway,  homologous recombination , and the  base excision repair  (BER) pathway. To repair interstrand crosslinks in eukaryotes, a 3’ flap endonuclease from the NER,  XPF-ERCC1 , is recruited to the crosslinked DNA, where it assists in ‘unhooking’ the DNA by cleaving the 3’ strand at the crosslink site. The 5’ strand is then cleaved, either by  XPF-ERCC1  or another  endonuclease , forming a  double-strand break  (DSB), which can then be repaired by the  homologous recombination  pathway. [ 17 ]


The repair of interstrand crosslinks requires both  nucleotide excision repair  and  homologous recombinational  repair. [ 18 ] .  The  Werner syndrome  (WRN) helicase and the  BRCA1  protein collaborate as part of a cellular response to repair DNA interstrand crosslinks. [ 18 ]   In vitro , BRCA1 interacts directly with WRN to stimulate WRN helicase activity. [ 18 ]


DNA crosslinks generally cause loss of overlapping sequence information from the two strands of DNA.  Therefore, accurate repair of the damage depends on retrieving the lost information from an undamaged  homologous chromosome  in the same cell.  Retrieval can occur by pairing with a  sister chromatid  produced during a preceding round of replication.  In a  diploid cell  retrieval may also occur by pairing with a non-sister  homologous chromosome , as occurs especially during  meiosis . [ citation needed ]   Once pairing has occurred, the crosslink can be removed and correct information introduced into the damaged chromosome by homologous recombination.


Cleavage of the bond between a deoxyribose sugar in DNA's sugar-phosphate backbone and its associated nucleobase leaves an abasic site in double stranded DNA. These abasic sites are often generated as an intermediate and then restored in base excision repair. However, if these sites are allowed to persist, they can inhibit DNA replication and transcription. [ 19 ]  Abasic sites can react with amine groups on proteins to form DNA-protein crosslinks or with exocyclic amines of other nucleobases to form interstrand crosslinks. To prevent interstrand or DNA-protein crosslinks, enzymes from the BER pathway tightly bind the abasic site and sequester it from nearby reactive groups, as demonstrated in human alkyladenine DNA glycosylase (AAG) and  E. coli  3-methyladenine DNA glycosylase II (AlkA). [ 20 ]   in vitro  evidence demonstrated that the Interstand Cross-Links induced by abasic site (DOB-ICL) is a replication-blocking and highly miscoding lesion. Compared to several other TLS pols examined, pol η is likely to contribute to the TLS-mediated repair of the DOB-ICL  in vivo . [ 21 ]  By using O 6 -2'-deoxyguanosine-butylene-O 6 -2'-deoxyguanosine (O6-dG-C4-O6-dG)  DNA lesions which is a chemically stable structure, the bypassing activity of several DNA polymerases had been investigated and the results demonstrated that pol η exhibited the highest bypass activity; however, 70% of the bypass products were mutagenic containing substitutions or deletions. The increase in the size of unhooked repair intermediates elevates the frequency of deletion mutation.  [ 22 ]


Treatment of  E. coli  with  psoralen -plus-UV light ( PUVA ) produces interstrand crosslinks in the cells’ DNA.  Cole et al. [ 23 ]   and Sinden and Cole [ 24 ]  presented evidence that a  homologous recombinational  repair process requiring the products of genes  uvrA ,  uvrB , and  recA  can remove these crosslinks in  E. coli . This process appears to be quite efficient.  Even though one or two unrepaired crosslinks are sufficient to inactivate a cell, a wild-type bacterial cell can repair and therefore recover from 53 to 71 psoralen crosslinks.   Eukaryotic  yeast cells are also inactivated by one remaining crosslink, but wild type yeast cells can recover from 120 to 200 crosslinks. [ 25 ]


DNA-protein crosslinking can be caused by a variety of chemical and physical agents, including transition metals, ionizing radiation, and endogenous aldehydes, in addition to  chemotherapeutic agents . [ 26 ]
Similar to DNA crosslinking, DNA-protein crosslinks are lesions in cells that are frequently damaged by UV radiation. The UV's effect can lead to reactive interactions and cause DNA and the proteins that are in contact with it to crosslink. These crosslinks are very bulky and complex lesions. They primarily occur in areas of the chromosomes that are undergoing DNA replication and interfere with cellular processes.


The advancement in structure-identification methods has progressed, and the addition in the ability to measure interactions between DNA and protein is a requirement to fully understand the biochemical processes. The structure of DNA-protein complexes can be mapped by  photocrosslinking , which is the photoinduced formation of a covalent bond between two macromolecules or between two different parts of one macromolecule. The methodology involves covalently linking a DNA-binding motif of the target sequence-specific DNA-binding protein with a photoactivatable crosslinking agent capable of reacting with DNA nucleotides when exposed to UV. This method provides information on the interaction between the DNA and protein in the crosslink. [ 27 ]


DNA repair pathways can result in the formation of  tumor cells .  Cancer  treatments have been engineered using DNA cross-linking agents to interact with nitrogenous bases of DNA to block DNA replication. These cross-linking agents have the ability to act as single-agent therapies by targeting and destroying specific nucleotides in cancerous cells. This result is stopping the cycle and growth of cancer cells; because it inhibits specific DNA repair pathways, this approach has a potential advantage in having fewer side effects. [ 28 ]


In humans, the leading cause of cancer deaths worldwide is lung cancer, including  non small cell lung carcinoma  (NSCLC) which accounts for 85% of all lung cancer cases in the United States. [ 29 ]   Individuals with NSCLC are often treated with therapeutic platinum compounds (e.g. cisplatin, carboplatin or oxaliplatin) (see  Lung cancer chemotherapy ) that cause interstrand DNA crosslinks. Among individuals with NSLC, low expression of the breast cancer 1 gene ( BRCA1 )  in the primary tumor has correlated with improved survival after platinum-containing chemotherapy. [ 30 ] [ 31 ]   This correlation implies that low BRCA1 in the cancer, and the consequent low level of DNA repair, causes vulnerability of the cancer to treatment by the DNA crosslinking agents.  High BRCA1 may protect cancer cells by acting in the  homologous recombinational  repair pathway that removes the damages in DNA introduced by the platinum drugs. The level of  BRCA1  expression is potentially an important tool for tailoring chemotherapy in lung cancer management. [ 30 ] [ 31 ]


Clinical chemotherapeutics can induce enzymatic and non-enzymatic DNA-protein crosslinks. An example of this induction is with platinum derivatives, such as cisplatin and oxaliplatin. They create non-enzymatic DNA-protein crosslinks through non-specific crosslinking of chromatin-interacting proteins to DNA. Crosslinking is also possible in other therapeutic agents by either stabilizing covalent DNA–protein reaction intermediates or by creating a pseudosubstrate, which traps the enzyme on DNA. Camptothecin derivatives, such as irinotecan and topotecan, target and trap specific DNA  topoisomerase  1 (TOP1) by intercalating within the enzyme–DNA interface. Because the toxicity of these drugs depends on TOP1 trapping, cellular sensitivity to these compounds depends directly on TOP1 expression levels. As a result, the function of these drugs is to serve as enzyme poisons rather than inhibitors. This can be applied to treat tumor cells by utilizing TOP 2 enzyme poisons. [ 32 ]


In molecular biology  ctRNA  (counter-transcribed  RNA ) is a plasmid encoded  noncoding RNA  that binds to the  mRNA  of repB and causes  translational  inhibition. [ 1 ]
ctRNA is encoded by plasmids and functions in  rolling circle replication  to maintain a low copy number. In  Corynebacterium glutamicum , it achieves this by  antisense  pairing with the mRNA of RepB, a  replication  initiation protein. [ 1 ]
In  Enterococcus faecium  the plasmid pJB01 contains three open reading frames, copA, repB, and repC. The pJB01 ctRNA is coded on the opposite strand from the {{chem name|copA}/repB intergenic region and partially overlaps an atypical ribosome binding site for repB. [ 2 ]


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In genetics and cell biology, repression is a mechanism often used to decrease or inhibit the expression of a gene. Removal of repression is called  derepression . This mechanism may occur at different stages in the expression of a gene, with the result of increasing the overall RNA or protein products. Dysregulation of derepression mechanisms can result in altered gene expression patterns, which may lead to negative phenotypic consequences such as disease.


Transcription  can be repressed in a variety of ways, and therefore can be derepressed in different ways as well. A common mechanism is  allosteric regulation . This is when a  substrate  binds a  repressor protein  and causes it to undergo a  conformational change . If the repressor is bound upstream of a gene, such as in an operator sequence, then it would be repressing the gene's expression. This conformational change would take away the repressor’s ability to bind DNA, thus removing its repressive effect on transcription. [ 1 ]


Another form of transcriptional derepression uses  chromatin remodeling  complexes. For transcription to occur,  RNA polymerase  needs to have access to the  promoter  sequence of the gene or it cannot bind the DNA. Sometimes these sequences are wrapped around  nucleosomes  or are in condensed  heterochromatin  regions, and are therefore inaccessible. Through different chromatin remodeling mechanisms these promoter sequences can become accessible to the RNA polymerase, and transcription becomes derepressed. [ 2 ]


Transcriptional derepression may also occur at the level of  transcription factor  activation. Certain families of transcription factors are non-functional on their own because their  active domains  are blocked by another part of the protein. [ 3 ]  Substrate binding to this second, regulatory domain causes a conformational change in the protein to allows access to the active domain. [ 3 ]  This lets the transcription factor bind to DNA and serve its function, thus derepressing the transcription factor.


Derepression of  translation  increases protein production without altering the levels of  mRNA  in the cell.  miRNAs  are a common mechanism of translation repression, binding to the mRNA through complementary base pairing to silence them. [ 4 ]  Certain  RNA binding proteins  have been shown to target  untranslated regions  of the mRNAs and upregulate the translation initiation rates by alleviating the repressive miRNA effects. [ 5 ]


An example is the  auxin  mediated derepression of the auxin response factor family of transcription factors in plants. These auxin response factors are repressed by Aux/IAA repressors. In the presence of auxin, these Aux/AII proteins undergo  ubiquitination  and are then degraded. [ 6 ] [ 7 ]  This derepresses the auxin response factors so they may carry out their functions in the cell.


Alzheimer’s  is a  neurodegenerative  disease involving progressive memory loss and other declines in brain function. One common cause of familial Alzheimer’s is mutation in the  PSEN1  gene. [ 8 ]  This gene encodes a protein that cleaves certain intracellular peptides which, once free in the  cytoplasm , promote  CBP degradation. Mutations in  PSEN1  decrease its production or ability to cleave proteins. This derepresses the CBP proteins, and allows them to perform their function of upregulating transcription of their target genes. [ 8 ]


Rett syndrome is a  neurodevelopmental disorder  involving deterioration of learned language and motor skills,  autism , and seizures starting in infancy. Many cases of Rett syndrome are associated with mutations in  MECP2 , a gene encoding a transcriptional repressor. [ 8 ]  Mutations in this gene decrease the levels of MeCP2 binding to different promoter sequences, resulting in their overall derepression. The increased expression of these MeCP2 regulated genes in  neurons  contribute to the Rett syndrome phenotype. [ 8 ] [ 9 ]


This syndrome is associated with increased susceptibility to tumors and growth abnormalities in children. A common cause of this syndrome is a mutation in an  imprint control region  near the  Igf2  gene . [ 9 ]  This imprint control region is normally bound by an  insulator  on the maternal  allele , which represses an  enhancer  from acting on the  Igf2  gene. This insulator is absent on the paternal allele and allows it access to the gene. Mutations in this imprint control region inhibit the insulator from binding, which derepresses enhancer activity on the maternal Igf2 gene. This abnormal derepression and increase in gene expression can result in Beckwith-Wiedemann syndrome. [ 9 ]


The  Dische test , or  Dische reaction , is used to distinguish  DNA  from  RNA . It was invented by  Zacharias Dische .


Dische's diphenylamine reagent consists of  diphenylamine , glacial acetic acid, sulfuric acid, and ethanol. [ 1 ]


When heated with DNA, it turns blue in the presence of DNA. A more intense blue color indicates a greater concentration of DNA.


The acid converts deoxyribose to a molecule that binds with diphenylamine to form a blue substance. The reagent does not interact with RNA, so can be used to distinguish DNA from RNA. [ 2 ] [ 3 ]


In  molecular genetics , a  DNA adduct  is a segment of  DNA  bound to a  cancer-causing chemical . This process could lead to the development of cancerous cells, or  carcinogenesis . DNA adducts in scientific experiments are used as  biomarkers  of exposure. They are especially useful in quantifying an organism's exposure to a carcinogen. [ 1 ]  The presence of such an adduct indicates prior exposure to a potential carcinogen, but it does not necessarily indicate the presence of cancer in the subject animal.


DNA adducts are researched in laboratory settings. A typical experimental design for studying DNA adducts is to induce them with known  carcinogens . A scientific journal will often incorporate the name of the carcinogen with their experimental design. For example, the term "DMBA-DNA adduct" in a scientific journal refers to a piece of DNA that has DMBA ( 7,12-dimethylbenz(a)anthracene ) attached to it.   [ 2 ]


Several diseases, including cancer, develop from mutated DNA. These mutations are caused by  carcinogens  through external and internal factors. Carcinogens are chemical or physical agents that cause DNA damage, which may later develop into cancer. They can initiate mutagenesis in DNA by interfering with the  replication  process. [ 3 ]  These interactions typically cause chemical adducts to form in the cell. This allows for DNA adducts to serve as biomarkers of exposure to carcinogens from the environment. They are attractive biomarkers because they are stable, abundant, and easily characterizable. Exposure to them can directly or indirectly cause DNA damage. In the direct case, a carcinogen can bind to DNA and cause it to distort or become cross-linked. Although  DNA repair happens  under normal circumstances, sometimes the DNA will not repair itself. This could be the start of a mutation, or  mutagenesis . Repeated mutations can lead to  carcinogenesis  – the beginnings of cancer. [ 4 ]


The presence of endogenous carcinogens contributes to levels of DNA adducts in a patient. This can bias the quantification of carcinogens that are from environmental exposure. Ongoing research on DNA adducts seeks to overcome these complications. It is the hope that in future medical practices DNA adducts may serve to guide therapeutic treatments that are more targeted and effective. [ 5 ]


Adduct formation is determined by the structures of reactive chemicals, the movement(s) of  electrophiles , and the capacity of the compounds to bind with DNA, potentially driving adduct formation to specific  nucleophilic  sites. The N3 and N7 locations (nucleotide positioning) of guanine and adenine are believed to be the most nucleophilic, and hence, they form adducts selectively over  exocyclic  oxygen atoms. The generation of DNA adducts is also influenced by certain  steric factors . Guanine's N7 position is exposed in the major groove of  double-helical DNA , making it more suitable for adduction than when compared to adenine's N3 position, which is orientated in the minor groove. [ 6 ]


Many compounds require enzyme metabolic activation to become  mutagenic  and cause DNA damage. Furthermore, reactive intermediates can be produced in the body as a result of  oxidative stress , thus harming the DNA. Some chemical carcinogens, metabolites, as well as  endogenous  compounds generated by inflammatory processes cause oxidative stress. This can result in the formation of a  reactive oxygen species  (ROS) or  reactive nitrogen species  (RNS). ROS and RNS are known to cause DNA damage via oxidative processes. Figure 2 shows each of the reactive sites for the nucleic acids involved in adduction and damage, with each form of transfer distinguished by arrow color. These positions are of interest to researchers studying DNA adduct formation. Research has indicated that many different chemicals may change human DNA and that lifestyle and host characteristics can impact the extent of DNA damage. Humans are constantly exposed to a diverse combination of potentially dangerous substances that might cause DNA damage. [ 6 ]


32P-postlabeling assay:


Liquid chromatography–mass spectrometry (LC–MS):


Fluorescence labeling:


Enzyme linked immunosorbent assay (ELISA):


Human consumption of more than 2.5–3.5 oz (70–100 g) of  red meat  (beef, lamb or pork) a day increases the risk of  colon cancer , but eating chicken does not have this risk. [ 21 ] [ 22 ]  The increased risk of colon cancer from red meat may be due to higher increases in DNA adducts from digestion of red meat. When rats were fed either beef or chicken, three types of DNA adducts in colon tissue were significantly higher after consumption of beef than after consumption of chicken. [ 23 ]  These adducts were a type of methyl-cytosine (possibly N3-methyl-cytosine), an adduct of two malondialdehyde molecules with guanine, and carboxyl-adenine. [ 24 ]


Human exposure to tobacco smoke has been associated with an increased risk of lung cancer. Tobacco smoke can impose great risk to DNA, with chemicals such as  formaldehyde  and  acetaldehyde  reacting directly with DNA to form adducts. In addition, there are other tobacco-specific carcinogens to consider in humans that are activated metabolically, such as  nicotine-derived nitrosamine ketone  (NNK) and  N'-nitrosonornicotine  (NNN). These carcinogens end up forming adducts when reacted with DNA, with those being called pyridyl oxobutyl (POB) adducts. [ 25 ]


Further analysis has been conducted on the topic, determining that  1,3-Butadiene  (BD) is a human carcinogen that is found in cigarette smoke among other  synthetic polymer  industries. Tests were conducted to understand the differences in the level of urinary BD-DNA adducts among various ethnic groups – white, Japanese American, and Native Hawaiian. It was determined that Japanese American smokers exhibited heightened levels of urinary BD-induced  guanine  adducts than white and Native Hawaiian individuals, while there were no differences in outcome by ethnicity among non-smokers. Understanding the  epigenetic  and genetic factors driving these differences in urinary BD-DNA adduct presence is the next step for this research, serving as a link between sociology and the life sciences. [ 26 ]


Particulate matter  (PM), broadly known as air pollution, is considered a  group 1 carcinogen  by the  International Agency for Research on Cancer ; while it is unclear if a direct link between cancer and PM exposure exists, it is likely that PM exposure leads to some degree of cell damage. Upon further investigation, it was determined that PM exposure causes  oxidative stress  – creating reactive oxygen species, forming DNA adducts, and inducing  double-strand breaks  (DSBs). In regards to DNA adduct formation, this analysis was conducted after looking at  leukocytes  from residents of heavily-populated cities (e.g. pollution, long-term traffic); a common component of PMs,  polycyclic aromatic hydrocarbon  (PAH), was one of the many molecules considered to be highly correlated with the presence of DNA bulky  lesions  in these individuals. These findings support the theory that DNA adduct presence indicates a level of carcinogenic activity. [ 27 ]


DNA adenine methyltransferase identification , often abbreviated  DamID , [ 1 ]  is a  molecular biology  protocol used to map the binding sites of  DNA - and  chromatin -binding  proteins  in  eukaryotes . DamID identifies binding sites by expressing the proposed DNA-binding protein as a  fusion protein  with  DNA methyltransferase . Binding of the protein of interest to DNA localizes the methyltransferase in the region of the binding site.  Adenine  methylation does not occur naturally in eukaryotes and therefore adenine methylation in any region can be concluded to have been caused by the fusion protein, implying the region is located near a binding site. DamID is an alternate method to  ChIP-on-chip  or  ChIP-seq . [ 2 ]


N6-methyladenine (m6A) is the product of the addition of a methyl group (CH 3 ) at position 6 of the adenine. This modified nucleotide is absent from the vast majority of eukaryotes, with the exception of  C. elegans , [ 3 ]  but is widespread in bacterial genomes, [ 4 ]  as part of the  restriction modification  or  DNA repair  systems. In  Escherichia coli , adenine methylation is catalyzed by the adenine methyltransferase  Dam  (DNA adenine methyltransferase), which catalyses adenine methylation exclusively in the palindromic sequence GATC. Ectopic expression of Dam in eukaryotic cells leads to methylation of adenine in GATC sequences without any other noticeable side effect.


Based on this, DamID consists in fusing Dam to a protein of interest (usually a protein that interacts with DNA such as  transcription factors ) or a chromatin component. The protein of interest thus targets Dam to its cognate  in vivo  binding site, resulting in the methylation of neighboring GATCs. The presence of m6A, coinciding with the binding sites of the proteins of interest, is revealed by  methyl PCR .


In this assay the genome is digested by  DpnI , which cuts only methylated GATCs. Double-stranded adapters with a known sequence are then ligated to the ends generated by DpnI. Ligation products are then digested by  DpnII . This enzyme cuts non-methylated GATCs, ensuring that only fragments flanked by  consecutive  methylated GATCs are amplified in the subsequent PCR. A PCR with primers matching the adaptors is then carried out, leading to the specific amplification of genomic fragments flanked by methylated GATCs.


Chromatin Immuno-Precipitation , or (ChIP), is an alternative method to assay protein binding at specific loci of the genome. Unlike ChIP, DamID does not require a specific  antibody  against the protein of interest. On the one hand, this allows to map proteins for which no such antibody is available. On the other hand, this makes it impossible to specifically map  posttranslationally modified  proteins.


Another fundamental difference is that ChIP assays where the protein of interests  is  at a given time, whereas DamID assays where it  has been . The reason is that m6A stays in the DNA after the Dam fusion protein goes away. For proteins that are either bound or unbound on their target sites this does not change the big picture. However, this can lead to strong differences in the case of proteins that slide along the DNA ( e.g.  RNA polymerase).


Depending on how the experiment is carried out, DamID can be subject to plasmid methylation biases. Because plasmids are usually amplified in  E. coli  where Dam is naturally expressed, they are methylated on every GATC. In transient  transfection  experiments, the DNA of those plasmids is recovered along with the DNA of the transfected cells, meaning that fragments of the plasmid are amplified in the methyl PCR. Every sequence of the genome that shares  homology  or identity with the plasmid may thus appear to be bound by the protein of interest. In particular, this is true of the  open reading frame  of the protein of interest, which is present in both the plasmid and the genome. In  microarray  experiments, this bias can be used to ensure that the proper material was hybridized. In stable cell lines or fully transgenic animals, this bias is not observed as no plasmid DNA is recovered.


Apoptotic cells degrade their DNA in a characteristic nucleosome ladder pattern. This generates DNA fragments that can be ligated and amplified during the DamID procedure (van Steensel laboratory, unpublished observations). The influence of these nucleosomal fragments on the binding profile of a protein is not known.


The resolution of DamID is a function of the availability of GATC sequences in the genome. A protein can only be mapped within two consecutive GATC sites. The median spacing between GATC fragments is 205 bp in Drosophila (FlyBase release 5), 260 in mouse (Mm9), and 460 in human (HG19). A modified protocol (DamIP), which combines  immunoprecipitation  of m6A with a Dam variant with less specific target site recognition, may be used to obtain higher resolution data. [ 5 ]


A major advantage of DamID over ChIP seq is that profiling of protein binding sites can be assayed in a particular cell type  in vivo  without requiring the physical separation of a subpopulation of cells. This allows for investigation into developmental or physiological processes in animal models.


The targeted DamID (TaDa) approach uses the phenomenon of ribosome reinitiation to express Dam-fusion proteins at appropriately low levels for DamID (i.e. Dam is non-saturating, thus avoiding toxicity). This construct can be combined with cell-type specific promoters resulting in tissue-specific methylation. [ 6 ] [ 7 ]  This approach can be used to assay transcription factor binding in a cell type of interest or alternatively, dam can be fused to  Pol II  subunits to determine binding of RNA polymerase and thus infer cell-specific gene expression. Targeted DamID has been demonstrated in  Drosophila  and mouse [ 8 ] [ 9 ]  cells.


Cell-specific DamID can also be achieved using  recombination  mediated excision of a transcriptional  terminator  cassette upstream of the Dam-fusion protein. [ 10 ]  The terminator cassette is flanked by FRT recombination sites which can be removed when combined with tissue specific expression of  FLP recombinase . Upon removal of the cassette, the Dam-fusion is expressed at low levels under the control of a basal promoter.


As well as detection of standard protein-DNA interactions, DamID can be used to investigate other aspects of chromatin biology.


This method can be used to detect co-binding of two factors to the same genomic  locus . The Dam methylase may be expressed in two halves which are fused to different proteins of interest. When both proteins bind to the same region of DNA, the Dam enzyme is reconstituted and is able to methylate the surrounding GATC sites. [ 11 ]


Due to the high activity of the enzyme, expression of untethered Dam results in methylation of all regions of accessible chromatin. [ 12 ] [ 13 ]  This approach can be used as an alternative to  ATAC-seq  or  DNAse-seq . When combined with cell-type specific DamID methods, expression of untethered Dam can be used to identify cell-type specific promoter or enhancer regions.


A DamID variant known as RNA-DamID can be used to detect interactions between RNA molecules and DNA. [ 14 ]  This method relies on the expression of a Dam-MCP fusion protein which is able to bind to an RNA that has been modified with  MS2  stem-loops. Binding of the Dam-fusion protein to the RNA results in detectable methylation at sites of RNA binding to the genome.


DNA sequences distal to a protein binding site may be brought into physical proximity through looping of chromosomes. For example, such interactions mediate  enhancer  and  promoter  function. These interactions can be detected through the action of Dam methylation. If Dam is targeted to a specific known DNA locus, distal sites brought into proximity due to the 3D configuration of the DNA will also be methylated and can be detected as in conventional DamID. [ 15 ]


DamID is usually performed on around 10,000 cells, [ 16 ]  (although it has been demonstrated with fewer [ 6 ] ). This means that the data obtained represents the average binding, or probability of a binding event across that cell population. A DamID protocol for single cells has also been developed and applied to human cells. [ 17 ]  Single cell approaches can highlight the heterogeneity of chromatin associations between cells.


The first isolation of  deoxyribonucleic acid  (DNA) was done in 1869 by  Friedrich Miescher . [ 1 ]   DNA extraction  is the process of isolating DNA from the cells of an organism isolated from a sample, typically a biological sample such as blood, saliva, or tissue. It involves breaking open the cells, removing proteins and other contaminants, and purifying the DNA so that it is free of other cellular components. The purified DNA can then be used for downstream applications such as  PCR , [ 2 ]   sequencing , or  cloning . Currently, it is a routine procedure in  molecular biology  or  forensic  analyses.


This process can be done in several ways, depending on the type of the sample and the downstream application, [ 3 ]  the most common methods are: mechanical, chemical and enzymatic lysis, precipitation, purification, and concentration. The specific method used to extract the DNA, such as phenol-chloroform extraction, alcohol precipitation, or silica-based purification. [ 4 ]


For the chemical method, many different kits are used for extraction, and selecting the correct one will save time on kit optimization and extraction procedures.  PCR  sensitivity detection is considered to show the variation between the commercial kits. [ 5 ]


There are many different methods for extracting DNA, but some common steps include:


Some variations on these steps may be used depending on the specific DNA extraction protocol. Additionally, some kits are commercially available that include reagents and protocols specifically tailored to a specific type of sample. [ 6 ]


DNA extraction is frequently a preliminary step in many diagnostic procedures used to identify environmental viruses and bacteria and diagnose illnesses and hereditary diseases. These methods consist of, but are not limited to:


Fluorescence In Situ Hybridization (FISH) technique was developed in the 1980s. The basic idea is to use a nucleic acid probe to hybridize nuclear DNA from either interphase cells or metaphase chromosomes attached to a microscopic slide. It is a molecular method used, among other things, to recognize and count particular bacterial groupings. [ 1 ]


To recognize, define, and quantify the geographical and temporal patterns in marine bacterioplankton communities, researchers employ a technique called terminal restriction fragment length polymorphism (T-RFLP).


Sequencing: Whole or partial genomes and other chromosomal components, ended for comparison with previously published sequences.


Cellular and  histone  proteins bound to the DNA can be removed either by adding a  protease  or having precipitated the proteins with  sodium  or  ammonium acetate  or  extracted them with a phenol-chloroform  mixture before the DNA precipitation.


After isolation, the DNA is dissolved in a slightly alkaline buffer, usually in a  TE buffer , or in  ultra-pure water .


The most common chemicals used for DNA extraction include:


Some of the most common DNA extraction methods include  organic extraction ,  Chelex extraction , and  solid phase extraction . [ 8 ]  These methods consistently yield isolated DNA, but they differ in both the quality and the quantity of DNA yielded. When selecting a DNA extraction method, there are multiple factors to consider, including cost, time, safety, and risk of contamination.


Organic extraction  involves the addition of incubation in multiple different chemical solutions; [ 8 ]  including a  lysis  step, a phenol-chloroform extraction, an  ethanol precipitation , and washing steps. Organic extraction is often used in laboratories because it is cheap, and it yields large quantities of pure DNA. Though it is easy, there are many steps involved, and it takes longer than other methods. It also involves the unfavorable use of the toxic chemicals  phenol  and  chloroform , and there is an increased risk of contamination due to transferring the DNA between multiple tubes. [ 9 ]  Several protocols based on organic extraction of DNA were effectively developed decades ago, [ 10 ]  though improved and more practical versions of these protocols have also been developed and published in the last years. [ 11 ]


The chelex extraction  method involves adding the Chelex resin to the sample, boiling the solution, then vortexing and centrifuging it. The cellular materials bind to the Chelex beads, while the DNA is available in the  supernatant . [ 9 ]  The Chelex method is much faster and simpler than organic extraction, and it only requires one tube, which decreases the risk of DNA contamination. Unfortunately, Chelex extraction does not yield as much quantity and the DNA yielded is single-stranded, which means it can only be used for  PCR -based analyses and not for  RFLP . [ 9 ]


Solid phase extraction  such as using a  spin-column-based extraction  method takes advantage of the fact that DNA binds to  silica . The sample containing DNA is added to a column containing a silica gel or silica beads and  chaotropic  salts. The chaotropic salts disrupt the hydrogen bonding between strands and facilitate the binding of the DNA to silica by causing the nucleic acids to become hydrophobic. This exposes the phosphate residues so they are available for adsorption. [ 12 ]  The DNA binds to the silica, while the rest of the solution is washed out using ethanol to remove chaotropic salts and other unnecessary constituents. [ 8 ]  The DNA can then be rehydrated with aqueous low-salt solutions allowing for  elution  of the DNA from the beads.


This method yields high-quality, largely double-stranded DNA which can be used for both  PCR  and  RFLP  analysis. This procedure can be automated [ 9 ]  and has a high throughput, although lower than the  phenol-chloroform method . This is a one-step method i.e. the entire procedure is completed in one tube. This lowers the risk of contamination making it very useful for the forensic extraction of DNA. Multiple solid-phase extraction commercial kits are manufactured and marketed by different companies; the only problem is that they are more expensive than organic extraction or Chelex extraction.


Specific techniques must be chosen for the isolation of DNA from some samples. Typical samples with complicated DNA isolation are:


Extrachromosomal DNA is generally easy to isolate, especially  plasmids  may be easily isolated by cell lysis followed by precipitation of proteins, which traps chromosomal DNA in insoluble fraction and after centrifugation, plasmid DNA can be purified from soluble fraction.


A Hirt DNA Extraction is an isolation of all  extrachromosomal DNA  in a mammalian cell. The Hirt extraction process gets rid of the high molecular weight  nuclear DNA , leaving only low molecular weight  mitochondrial DNA  and any viral  episomes  present in the cell.


A  diphenylamine (DPA) indicator  will confirm the presence of DNA. This procedure involves chemical hydrolysis of DNA: when heated (e.g. ≥95 °C) in acid, the reaction requires a  deoxyribose sugar  and therefore is specific for DNA. Under these conditions, the 2-deoxyribose is converted to w-hydroxylevulinyl aldehyde, which reacts with the compound, diphenylamine, to produce a blue-colored compound. DNA concentration can be determined by  measuring the intensity of absorbance of the solution at the 600 nm with a  spectrophotometer  and comparing to a  standard curve  of known DNA concentrations.


Measuring the intensity of absorbance of the DNA solution at wavelengths  260 nm and 280 nm  is used as a measure of DNA purity. DNA can be quantified by cutting the DNA with a  restriction enzyme , running it on an agarose  gel , staining with  ethidium bromide (EtBr)  or a different stain and comparing the intensity of the DNA with a DNA marker of known concentration.


Using the  Southern blot  technique, this quantified DNA can be isolated and examined further using  PCR  and  RFLP  analysis. These procedures allow differentiation of the repeated sequences within the genome. It is these techniques which  forensic  scientists use for comparison, identification, and analysis.


In this method, plant  nuclei  are isolated by physically grinding tissues and reconstituting the intact nuclei in a unique Nuclear Isolation Buffer (NIB). The plastid DNAs are released from organelles and eliminated with an osmotic buffer by washing and centrifugation. The purified nuclei are then lysed and further cleaned by organic extraction, and the genomic DNA is precipitated with a high concentration of CTAB. The highly pure, high molecular weight  gDNA  is extracted from the nuclei, dissolved in a high pH buffer, allowing for stable long-term storage. [ 14 ]


DNA storage is an important aspect of DNA extraction projects as it ensures the integrity and stability of the extracted DNA for downstream applications. [ 15 ]


One common method of DNA storage is ethanol precipitation, which involves adding ethanol and a salt, such as sodium chloride or potassium acetate, to the extracted DNA to precipitate it out of solution. The DNA is then pelleted by centrifugation and washed with 70% ethanol to remove any remaining contaminants. The DNA pellet is then air-dried and resuspended in a buffer, such as Tris-EDTA (TE) buffer, for storage.


Another method is freezing the DNA in a buffer such as TE buffer, or in a cryoprotectant such as glycerol or DMSO, at -20 or -80 degrees Celsius. This method preserves the integrity of the DNA and slows down the activity of any enzymes that may degrade it.


It's important to note that the choice of storage buffer and conditions will depend on the downstream application for which the DNA is intended. For example, if the DNA is to be used for PCR, it may be stored in TE buffer at 4 degrees Celsius, while if it is to be used for long-term storage or shipping, it may be stored in ethanol at -20 degrees Celsius. The extracted DNA should be regularly checked for its quality and integrity, such as by running a gel electrophoresis or spectrophotometry. The storage conditions should be also noted and controlled, such as the temperature and humidity.


It's also important to consider the long-term stability of the DNA and the potential for degradation over time. The extracted DNA should be stored for as short a time as possible, and the conditions for storage should be chosen to minimize the risk of degradation.


In general, the extracted DNA should be stored under the best possible conditions to ensure its stability and integrity for downstream applications.


There are several quality control techniques used to ensure the quality of extracted DNA, including: [ 16 ]


DNA ligase  is a type of enzyme that facilitates the joining of  DNA  strands together by catalyzing the formation of a  phosphodiester bond . It plays a role in repairing single-strand breaks in duplex  DNA  in living organisms, but some forms (such as  DNA ligase IV ) may specifically repair double-strand breaks (i.e. a break in both  complementary  strands of DNA). Single-strand breaks are repaired by DNA ligase using the complementary strand of the double helix as a template, [ 1 ]  with DNA ligase creating the final phosphodiester bond to fully repair the DNA.


DNA ligase is used in both  DNA repair  and  DNA replication  (see  Mammalian ligases ).  In addition, DNA ligase has extensive use in  molecular biology  laboratories for  recombinant DNA  experiments (see  Research applications ). Purified DNA ligase is used in gene cloning to join DNA molecules together to form  recombinant DNA .


The mechanism of DNA ligase is to form two  covalent   phosphodiester bonds  between  3' hydroxyl ends  of one  nucleotide  ("acceptor"), with the  5' phosphate end  of another ("donor"). Two ATP molecules are consumed for each phosphodiester bond formed.  [ citation needed ] AMP is required for the ligase reaction, which proceeds in four steps:


Ligase will also work with  blunt ends , although higher enzyme concentrations and different reaction conditions are required.


The  E. coli  DNA ligase is encoded by the  lig  gene.  DNA ligase in  E. coli , as well as most prokaryotes, uses energy gained by cleaving  nicotinamide adenine dinucleotide  (NAD) to create the phosphodiester bond. [ 3 ]   It does not ligate blunt-ended DNA except under conditions of molecular crowding with  polyethylene glycol , and cannot join RNA to DNA efficiently. [ citation needed ]


The activity of E. coli DNA ligase can be enhanced by  DNA polymerase  at the right concentrations. Enhancement only works when the concentrations of the DNA polymerase 1 are much lower than the DNA fragments to be ligated. When the concentrations of Pol I DNA polymerases are higher, it has an adverse effect on E. coli DNA ligase [ 4 ]


The DNA ligase from  bacteriophage T4  (a  bacteriophage  that infects  Escherichia coli  bacteria). The T4 ligase is the most-commonly used in laboratory research. [ 5 ]   It can ligate either  cohesive or blunt ends  of DNA, oligonucleotides, as well as RNA and RNA-DNA hybrids, but not single-stranded nucleic acids. It can also ligate  blunt-ended DNA  with much greater efficiency than  E. coli  DNA ligase.  Unlike  E. coli  DNA ligase, T4 DNA ligase cannot utilize NAD and it has an absolute requirement for ATP as a cofactor. Some engineering has been done to improve the  in vitro  activity of T4 DNA ligase; one successful approach, for example, tested T4 DNA ligase fused to several alternative DNA binding proteins and found that the constructs with either p50 or  NF-kB  as fusion partners were over 160% more active in blunt-end ligations for cloning purposes than wild type T4 DNA ligase. [ 6 ]   A typical reaction for inserting a fragment into a plasmid vector would use about 0.01 (sticky ends) to 1 (blunt ends) units of ligase.  The optimal incubation temperature for T4 DNA ligase is 16 °C. [ citation needed ]


Bacteriophage T4  ligase  mutants  have increased sensitivity to both  UV  irradiation [ 7 ] [ 8 ]  and the alkylating agent  methyl methanesulfonate [ 9 ]  indicating that DNA ligase is employed in the  repair  of the  DNA damages  caused by these agents.


In mammals, there are four specific types of ligase.


DNA ligase from  eukaryotes  and some microbes uses  adenosine triphosphate  (ATP) rather than NAD. [ 3 ]


Derived from a thermophilic bacterium, the enzyme is stable and active at much higher temperatures than conventional DNA ligases. Its half-life is 48 hours at 65 °C and greater than 1 hour at 95 °C. Ampligase DNA Ligase has been shown to be active for at least 500 thermal cycles (94 °C/80 °C) or 16 hours of cycling. 10  This exceptional thermostability permits extremely high hybridization stringency and ligation specificity. [ 11 ]


There are at least three different units used to measure the activity of DNA ligase: [ 12 ]


DNA ligases have become indispensable tools in modern molecular biology research for generating  recombinant DNA  sequences.  For example, DNA ligases are used with  restriction enzymes  to insert DNA fragments, often  genes , into  plasmids .


Controlling the optimal temperature is a vital aspect of performing efficient recombination experiments involving the ligation of cohesive-ended fragments.  Most experiments use T4 DNA Ligase (isolated from  bacteriophage T4 ), which is most active at 37 °C. [ 13 ]   However, for optimal ligation efficiency with cohesive-ended fragments ("sticky ends"), the optimal enzyme temperature needs to be balanced with the  melting temperature  T m  of the sticky ends being ligated, [ 14 ]  the  homologous pairing  of the sticky ends will not be stable because the high temperature disrupts  hydrogen bonding . A ligation reaction is most efficient when the sticky ends are already stably annealed, and disruption of the annealing ends would therefore result in low ligation efficiency. The shorter the  overhang , the lower the T m .


Since blunt-ended DNA fragments have no cohesive ends to anneal, the melting temperature is not a factor to consider within the normal temperature range of the ligation reaction. The limiting factor in blunt end ligation is not the activity of the ligase but rather the number of alignments between DNA fragment ends that occur.  The most efficient ligation temperature for blunt-ended DNA would therefore be the temperature at which the greatest number of alignments can occur.  The majority of blunt-ended ligations are carried out at 14-25 °C overnight.  The absence of stably annealed ends also means that the ligation efficiency is lowered, requiring a higher ligase concentration to be used. [ 14 ]


A novel use of DNA ligase can be seen in the field of nano chemistry, specifically in DNA origami.  DNA based self-assembly principles have proven useful for organizing nanoscale objects, such as biomolecules, nanomachines, nanoelectronic and photonic component. Assembly of such nano structure requires the creation of an intricate mesh of DNA molecules. Although DNA self-assembly is possible without any outside help using different substrates such as provision of catatonic surface of Aluminium foil, DNA ligase can provide the enzymatic assistance that is required to make DNA lattice structure from DNA over hangs. [ 15 ]


The first DNA ligase was purified and characterized in 1967 by the Gellert, Lehman, Richardson, and Hurwitz laboratories. [ 16 ]  It was first purified and characterized by Weiss and Richardson using a six-step chromatographic-fractionation process beginning with elimination of cell debris and addition of streptomycin, followed by several Diethylaminoethyl (DEAE)-cellulose column washes and a final phosphocellulose fractionation. The final extract contained 10% of the activity initially recorded in the  E. coli  media; along the process it was discovered that ATP and Mg++ were necessary to optimize the reaction. The common commercially available DNA ligases were originally discovered in  bacteriophage T4 ,  E. coli  and other  bacteria . [ 17 ]


Genetic deficiencies in human DNA ligases have been associated with clinical syndromes marked by immunodeficiency, radiation sensitivity, and developmental abnormalities,  [ 16 ]   LIG4 syndrome  (Ligase IV syndrome) is a rare disease associated with mutations in DNA ligase 4 and interferes with dsDNA break-repair mechanisms. Ligase IV syndrome causes immunodeficiency in individuals and is commonly associated with microcephaly and marrow hypoplasia. [ 18 ]  A list of prevalent diseases caused by lack of or malfunctioning of DNA ligase is as follows.


Xeroderma pigmentosum , which is commonly known as XP, is an inherited condition characterized by an extreme sensitivity to ultraviolet (UV) rays from sunlight. This condition mostly affects the eyes and areas of skin exposed to the sun. Some affected individuals also have problems involving the nervous system. [ 19 ]


Mutations in the ATM gene cause  ataxia–telangiectasia . The ATM gene provides instructions for making a protein that helps control cell division and is involved in DNA repair. This protein plays an important role in the normal development and activity of several body systems, including the nervous system and immune system. The ATM protein assists cells in recognizing damaged or broken DNA strands and coordinates DNA repair by activating enzymes that fix the broken strands. Efficient repair of damaged DNA strands helps maintain the stability of the cell's genetic information. Affected children typically develop difficulty walking, problems with balance and hand coordination, involuntary jerking movements (chorea), muscle twitches (myoclonus), and disturbances in nerve function (neuropathy). The movement problems typically cause people to require wheelchair assistance by adolescence. People with this disorder also have slurred speech and trouble moving their eyes to look side-to-side (oculomotor apraxia). [ 20 ]


Fanconi anemia  (FA) is a rare, inherited blood disorder that leads to bone marrow failure. FA prevents bone marrow from making enough new blood cells for the body to work normally. FA also can cause the bone marrow to make many faulty blood cells. This can lead to serious health problems, such as  leukemia . [ 21 ]


Bloom syndrome  results in skin that is sensitive to sun exposure, and usually the development of a butterfly-shaped patch of reddened skin across the nose and cheeks. A skin rash can also appear on other areas that are typically exposed to the sun, such as the back of the hands and the forearms. Small clusters of enlarged blood vessels (telangiectases) often appear in the rash; telangiectases can also occur in the eyes. Other skin features include patches of skin that are lighter or darker than the surrounding areas (hypopigmentation or hyperpigmentation respectively). These patches appear on areas of the skin that are not exposed to the sun, and their development is not related to the rashes.


In recent studies, human DNA ligase I was used in  Computer-aided drug design  to identify DNA ligase inhibitors as possible therapeutic agents to treat cancer. [ 22 ]  Since excessive cell growth is a hallmark of cancer development, targeted chemotherapy that disrupts the functioning of DNA ligase can impede adjuvant cancer forms. Furthermore, it has been shown that DNA ligases can be broadly divided into two categories, namely, ATP- and NAD + -dependent. Previous research has shown that although NAD + -dependent DNA ligases have been discovered in sporadic cellular or viral niches outside the bacterial domain of life, there is no instance in which a NAD + -dependent ligase is present in a  eukaryotic  organism. The presence solely in non-eukaryotic organisms, unique substrate specificity, and distinctive domain structure of NAD+ dependent compared with ATP-dependent human DNA ligases together make NAD + -dependent ligases ideal targets for the development of new antibacterial drugs. [ 16 ]


A  DNA machine  is a  molecular machine  constructed from  DNA .  Research into DNA machines was pioneered in the late 1980s by  Nadrian Seeman  and co-workers from  New York University . DNA is used because of the numerous biological tools already found in nature that can affect DNA, and the immense knowledge of how DNA works previously researched by  biochemists .


DNA machines can be logically designed since DNA assembly of the double helix is based on strict rules of base pairing that allow portions of the strand to be predictably connected based on their sequence.  This "selective stickiness" is a key advantage in the construction of DNA machines.


An example of a DNA machine was reported by  Bernard Yurke  and co-workers at  Lucent Technologies  in the year 2000, who constructed  molecular tweezers  out of DNA. [ 1 ]
The DNA tweezers contain three strands: A, B and C. Strand A latches onto half of strand B and half of strand C, and so it joins them all together. Strand A acts as a hinge so that the two "arms" — AB and AC — can move. The structure floats with its arms open wide. They can be pulled shut by adding a fourth strand of DNA (D) "programmed" to stick to both of the dangling, unpaired sections of strands B and C.   The closing of the tweezers was proven by tagging strand A at either end with light-emitting molecules that do not emit light when they are close together.  To re-open the tweezers add a further strand (E) with the right sequence to pair up with strand D. Once paired up, they have no connection to the machine BAC, so float away.  The DNA machine can be opened and closed repeatedly by cycling between strands D and E. These tweezers can be used for removing drugs from inside  fullerenes  as well as from a self assembled DNA  tetrahedron . The state of the device can be determined by measuring the separation between donor and acceptor fluorophores using  FRET .


DNA walkers  are another type of DNA machine.


A  DNA microarray  (also commonly known as  DNA  chip or  biochip ) is a collection of microscopic DNA spots attached to a solid surface. Scientists use DNA  microarrays  to measure the  expression  levels of large numbers of genes simultaneously or to  genotype  multiple regions of a genome. Each DNA spot contains  picomoles  (10 −12   moles ) of a specific DNA sequence, known as  probes  (or  reporters  or  oligos ). These can be a short section of a  gene  or other DNA element that are used to  hybridize  a  cDNA  or cRNA (also called anti-sense RNA)  sample (called  target ) under high-stringency conditions. Probe-target hybridization is usually detected and quantified by detection of  fluorophore -, silver-, or  chemiluminescence -labeled targets to determine relative abundance of nucleic acid sequences in the target. The original nucleic acid arrays were macro arrays approximately 9 cm × 12 cm and the first computerized image based analysis was published in 1981. [ 1 ]  It was invented by  Patrick O. Brown . An example of its application is in SNPs arrays for polymorphisms in cardiovascular diseases, cancer, pathogens and GWAS analysis. It is also used for the identification of structural variations and the measurement of gene expression.


The core principle behind microarrays is hybridization between two DNA strands, the property of  complementary  nucleic acid sequences to specifically pair with each other by forming  hydrogen bonds  between complementary  nucleotide base pairs . A high number of complementary base pairs in a nucleotide sequence means tighter  non-covalent  bonding between the two strands. After washing off non-specific bonding sequences, only strongly paired strands will remain hybridized. Fluorescently labeled target sequences that bind to a probe sequence generate a signal that depends on the hybridization conditions (such as temperature), and washing after hybridization. Total strength of the signal, from a spot (feature), depends upon the amount of target sample binding to the probes present on that spot. Microarrays use relative quantitation in which the intensity of a feature is compared to the intensity of the same feature under a different condition,  and the identity of the feature is known by its position.


Many types of arrays exist and the broadest distinction is whether they are spatially arranged on a surface or on coded beads:


DNA microarrays can be used to detect DNA (as in  comparative genomic hybridization ), or detect RNA (most commonly as  cDNA  after  reverse transcription ) that may or may not be translated into proteins. The process of measuring gene expression via cDNA is called  expression analysis  or  expression profiling .


Applications include:


Specialised arrays tailored to particular  crops  are becoming increasingly popular in  molecular breeding  applications. In the future they could be used to screen  seedlings  at early stages to lower the number of unneeded seedlings tried out in breeding operations. [ 10 ]


Microarrays can be manufactured in different ways, depending on the number of probes under examination, costs, customization requirements, and the type of scientific question being asked. Arrays from commercial vendors may have as few as 10 probes or as many as 5 million or more micrometre-scale probes.


Microarrays can be fabricated using a variety of technologies, including printing with fine-pointed pins onto glass slides,  photolithography  using pre-made masks, photolithography using dynamic micromirror devices, ink-jet printing, [ 11 ] [ 12 ]  or  electrochemistry  on microelectrode arrays.


In  spotted microarrays , the probes are  oligonucleotides ,  cDNA  or small fragments of  PCR  products that correspond to  mRNAs . The probes are  synthesized  prior to deposition on the array surface and are then "spotted" onto glass. A common approach utilizes an array of fine pins or needles controlled by a robotic arm that is dipped into wells containing DNA probes and then depositing each probe at designated locations on the array surface. The resulting "grid" of probes represents the nucleic acid profiles of the prepared probes and is ready to receive complementary cDNA or cRNA "targets" derived from experimental or clinical samples.
This technique is used by research scientists around the world to produce "in-house" printed microarrays in their own labs. These arrays may be easily customized for each experiment, because researchers can choose the probes and printing locations on the arrays, synthesize the probes in their own lab (or collaborating facility), and spot the arrays. They can then generate their own labeled samples for hybridization, hybridize the samples to the array, and finally scan the arrays with their own equipment. This provides a relatively low-cost microarray that may be customized for each study, and avoids the costs of purchasing often more expensive commercial arrays that may represent vast numbers of genes that are not of interest to the investigator.
Publications exist which indicate in-house spotted microarrays may not provide the same level of sensitivity compared to commercial oligonucleotide arrays, [ 13 ]  possibly owing to the small batch sizes and reduced printing efficiencies when compared to industrial manufactures of oligo arrays.


In  oligonucleotide microarrays , the probes are short sequences designed to match parts of the sequence of known or predicted  open reading frames . Although oligonucleotide probes are often used in "spotted" microarrays, the term "oligonucleotide array" most often refers to a specific technique of manufacturing. Oligonucleotide arrays are produced by printing short oligonucleotide sequences designed to represent a single gene or family of gene splice-variants by  synthesizing  this sequence directly onto the array surface instead of depositing intact sequences. Sequences may be longer (60-mer probes such as the  Agilent  design) or shorter (25-mer probes produced by  Affymetrix ) depending on the desired purpose; longer probes are more specific to individual target genes, shorter probes may be spotted in higher density across the array and are cheaper to manufacture.
One technique used to produce oligonucleotide arrays include  photolithographic  synthesis (Affymetrix) on a silica substrate where light and light-sensitive masking agents are used to "build" a sequence one nucleotide at a time across the entire array. [ 14 ]  Each applicable probe is selectively "unmasked" prior to bathing the array in a solution of a single nucleotide, then a masking reaction takes place and the next set of probes are unmasked in preparation for a different nucleotide exposure. After many repetitions, the sequences of every probe become fully constructed. More recently, Maskless Array Synthesis from NimbleGen Systems has combined flexibility with large numbers of probes. [ 15 ]


Two-color microarrays  or  two-channel microarrays  are typically  hybridized  with cDNA prepared from two samples to be compared (e.g. diseased tissue versus healthy tissue) and that are labeled with two different  fluorophores . [ 16 ]   Fluorescent  dyes commonly used for cDNA labeling include  Cy 3, which has a fluorescence emission wavelength of 570 nm (corresponding to the green part of the light spectrum), and  Cy 5 with a fluorescence emission wavelength of 670 nm (corresponding to the red part of the light spectrum). The two Cy-labeled cDNA samples are mixed and hybridized to a single microarray that is then scanned in a microarray scanner to visualize fluorescence of the two fluorophores after  excitation  with a  laser  beam of a defined wavelength. Relative intensities of each fluorophore may then be used in ratio-based analysis to identify up-regulated and down-regulated genes. [ 17 ]


Oligonucleotide microarrays often carry control probes designed to hybridize with  RNA spike-ins . The degree of hybridization between the spike-ins and the control probes is used to  normalize  the hybridization measurements for the target probes. Although absolute levels of gene expression may be determined in the two-color array in rare instances, the relative differences in expression among different spots within a sample and between samples is the preferred method of  data analysis  for the two-color system. Examples of providers for such microarrays includes  Agilent  with their Dual-Mode platform,  Eppendorf  with their DualChip platform for colorimetric  Silverquant  labeling, and TeleChem International with  Arrayit .


In  single-channel microarrays  or  one-color microarrays , the arrays provide intensity data for each probe or probe set indicating a relative level of hybridization with the labeled target. However, they do not truly indicate abundance levels of a gene but rather relative abundance when compared to other samples or conditions when processed in the same experiment. Each RNA molecule encounters protocol and batch-specific bias during amplification, labeling, and hybridization phases of the experiment making comparisons between genes for the same microarray uninformative. The comparison of two conditions for the same gene requires two separate single-dye hybridizations. Several popular single-channel systems are the Affymetrix "Gene Chip", Illumina "Bead Chip", Agilent single-channel arrays, the Applied Microarrays "CodeLink" arrays, and the Eppendorf "DualChip & Silverquant". One strength of the single-dye system lies in the fact that an aberrant sample cannot affect the raw data derived from other samples, because each array chip is exposed to only one sample (as opposed to a two-color system in which a single low-quality sample may drastically impinge on overall data precision even if the other sample was of high quality). Another benefit is that data are more easily compared to arrays from different experiments as long as batch effects have been accounted for.


One channel microarray may be the only choice in some situations. Suppose

 i

 {\displaystyle i}

  samples need to be compared: then the number of experiments required using the two channel arrays quickly becomes unfeasible, unless a sample is used as a reference.


two channel microarray (with reference)


This is an example of a  DNA microarray experiment  which includes details for a particular case to better explain DNA microarray experiments, while listing modifications for RNA or other alternative experiments.


The advent of inexpensive microarray experiments created several specific bioinformatics challenges: [ 19 ]  the multiple levels of replication in experimental design ( Experimental design ); the number of platforms and independent groups and data format ( Standardization ); the statistical treatment of the data ( Data analysis ); mapping each probe to the  mRNA  transcript that it measures ( Annotation ); the sheer volume of data and the ability to share it ( Data warehousing ).


Due to the biological complexity of gene expression, the considerations of experimental design that are discussed in the  expression profiling  article are of critical importance if statistically and biologically valid conclusions are to be drawn from the data.


There are three main elements to consider when designing a microarray experiment. First, replication of the biological samples is essential for drawing conclusions from the experiment. Second, technical replicates (e.g. two RNA samples obtained from each experimental unit) may help to quantitate precision. The biological replicates include independent RNA extractions. Technical replicates may be two  aliquots  of the same extraction. Third, spots of each cDNA clone or oligonucleotide are present as replicates (at least duplicates) on the microarray slide, to provide a measure of technical precision in each hybridization. It is critical that information about the sample preparation and handling is discussed, in order to help identify the independent units in the experiment and to avoid inflated estimates of  statistical significance . [ 20 ]


Microarray data is difficult to exchange due to the lack of standardization in platform fabrication, assay protocols, and analysis methods. This presents an  interoperability  problem in  bioinformatics . Various  grass-roots   open-source  projects are trying to ease the exchange and analysis of data produced with non-proprietary chips:


For example, the "Minimum Information About a Microarray Experiment" ( MIAME ) checklist helps define the level of detail that should exist and is being adopted by many  journals  as a requirement for the submission of papers incorporating microarray results. But MIAME does not describe the format for the information, so while many formats can support the MIAME requirements, as of 2007 [update]  no format permits verification of complete semantic compliance. The "MicroArray Quality Control (MAQC) Project" is being conducted by the US  Food and Drug Administration  (FDA) to develop standards and quality control metrics which will eventually allow the use of MicroArray data in drug discovery, clinical practice and regulatory decision-making. [ 21 ]  The  MGED Society  has developed standards for the representation of gene expression experiment results and relevant annotations.


Microarray data sets are commonly very large, and analytical precision is influenced by a number of variables.  Statistical  challenges include taking into account effects of background noise and appropriate  normalization  of the data. Normalization methods may be suited to specific platforms and, in the case of commercial platforms, the analysis may be proprietary. [ 22 ]  Algorithms that affect statistical analysis include:


Microarray data may require further processing aimed at reducing the dimensionality of the data to aid comprehension and more focused analysis. [ 33 ]  Other methods permit analysis of data consisting of a low number of biological or technical  replicates ; for example, the Local Pooled Error (LPE) test pools  standard deviations  of genes with similar expression levels in an effort to compensate for insufficient replication. [ 34 ]


The relation between a probe and the  mRNA  that it is expected to detect is not trivial. [ 35 ]  Some mRNAs may cross-hybridize probes in the array that are supposed to detect another mRNA. In addition, mRNAs may experience amplification bias that is sequence or molecule-specific. Thirdly, probes that are designed to detect the mRNA of a particular gene may be relying on genomic  EST  information that is incorrectly associated with that gene.


Microarray data was found to be more useful when compared to other similar datasets. The sheer volume of data, specialized formats (such as  MIAME ), and curation efforts associated with the datasets require specialized databases to store the data. A number of open-source data warehousing solutions, such as  InterMine  and  BioMart , have been created for the specific purpose of integrating diverse biological datasets, and also support analysis.


Advances in massively parallel sequencing has led to the development of  RNA-Seq  technology, that enables a whole transcriptome shotgun approach to characterize and quantify gene expression. [ 36 ] [ 37 ]  Unlike microarrays, which need a reference genome and transcriptome to be available before the microarray itself can be designed, RNA-Seq can also be used for new model organisms whose genome has not been sequenced yet. [ 37 ]


DNA origami  is the nanoscale folding of  DNA  to create arbitrary two- and three-dimensional shapes at the  nanoscale . The specificity of the interactions between  complementary base pairs  make DNA a useful construction material, through design of its base sequences. [ 2 ]  DNA is a well-understood material that is suitable for creating scaffolds that hold other molecules in place or to create structures all on its own.


DNA origami was the cover story of  Nature  on March 16, 2006. [ 3 ]  Since then, DNA origami has progressed past an art form and has found a number of applications from drug delivery systems to uses as circuitry in plasmonic devices; however, most commercial applications remain in a concept or testing phase. [ 4 ]


The idea of using DNA as a construction material was first introduced in the early 1980s by  Nadrian Seeman . [ 5 ]  The method of DNA origami was developed by  Paul Rothemund  at the  California Institute of Technology . [ 6 ]  In contrast to common top-down fabrication methods such as 3D printing or lithography which involve depositing or removing material through a tool, DNA Nanotechnology, as well as DNA Origami as a subset, is a bottom-up fabrication method. By rationally designing the constituent subunits of the DNA polymer, DNA can self-assemble into a variety of shapes. The process of constructing DNA Origami involves the folding of a long single strand of  viral   DNA  (typically the 7,249 bp genomic DNA of  M13 bacteriophage ) aided by multiple smaller "staple" strands. These shorter strands bind the longer in various places, resulting in the formation of a pre-defined two- or three-dimensional shape. [ 7 ]  Examples include a  smiley face  and a coarse map of China and the Americas, along with many three-dimensional structures such as cubes. [ 8 ]


There are several DNA properties that make the molecule an ideal building material for DNA origami. DNA strands have a natural tendency to bind to their complementary sequences through  Watson–Crick base pairing . This allows staple strands to locate the position on the scaffold strand without any external manipulation, leading to self-assembly of the desired structure.