In 2012, a group of American scientists led by Floyd Romesberg, a chemical biologist at the  Scripps Research Institute  in San Diego, California, published that his team designed an unnatural base pair (UBP). The two new artificial nucleotides or  Unnatural Base Pair  (UBP) were named  d5SICS  and  dNaM . More technically, these artificial  nucleotides  bearing hydrophobic  nucleobases , feature two fused  aromatic rings  that form a (d5SICS–dNaM) complex or base pair in DNA. In 2014 the same team from the Scripps Research Institute reported that they synthesized a stretch of circular DNA known as a  plasmid  containing natural T-A and C-G base pairs along with the best-performing UBP Romesberg's laboratory had designed, and inserted it into cells of the common bacterium  E. coli  that successfully replicated the unnatural base pairs through multiple generations. This is the first known example of a living organism passing along an expanded genetic code to subsequent generations. This was in part achieved by the addition of a supportive algal gene that expresses a  nucleotide triphosphate  transporter which efficiently imports the triphosphates of both d5SICSTP and dNaMTP into  E. coli  bacteria. Then, the natural bacterial replication pathways use them to accurately replicate the  plasmid  containing d5SICS–dNaM.


The successful incorporation of a third base pair is a significant breakthrough toward the goal of greatly expanding the number of  amino acids  which can be encoded by DNA, from the existing 20 amino acids to a theoretically possible 172, thereby expanding the potential for living organisms to produce novel  proteins . [ 20 ]  In the future, these unnatural base pairs could be synthesised and incorporated into oligonucleotides via DNA printing methods.


DNA printing can thus be used to produce DNA parts, which are defined as sequences of DNA that encode a specific biological function (for example,  promoters , transcription regulatory sequences or  open reading frames ). [ 21 ]  However, because oligonucleotide synthesis typically cannot accurately produce oligonucleotides sequences longer than a few hundred base pairs, DNA assembly methods have to be employed to assemble these parts together to create functional genes, multi-gene circuits or even entire synthetic chromosomes or genomes. Some DNA assembly techniques only define protocols for joining DNA parts, while other techniques also define the rules for the format of DNA parts that are compatible with them. These processes can be scaled up to enable the assembly of entire chromosomes or genomes. In recent years, there has been proliferation in the number of different DNA assembly standards with 14 different assembly standards developed as of 2015, each with their pros and cons. [ 22 ]   Overall, the development of DNA assembly standards has greatly facilitated the workflow of synthetic biology, aided the exchange of material between research groups and also allowed for the creation of modular and reusable DNA parts. [ 22 ]


The various DNA assembly methods can be classified into three main categories – endonuclease-mediated assembly, site-specific recombination, and long-overlap-based assembly. [ 22 ]  Each group of methods has its distinct characteristics and their own advantages and limitations.


Endonucleases  are enzymes that recognise and cleave nucleic acid segments and they can be used to direct DNA assembly. Of the different types of restriction enzymes, the type II restriction enzymes are the most commonly available and used because their cleavage sites are located near or in their recognition sites. Hence, endonuclease-mediated assembly methods make use of this property to define DNA parts and assembly protocols.


The BioBricks assembly standard was described and introduced by Tom Knight in 2003 and it has been constantly updated since then. [ 23 ]  Currently, the most commonly used BioBricks standard is the assembly standard 10, or BBF RFC 10. BioBricks defines the prefix and suffix sequences required for a DNA part to be compatible with the BioBricks assembly method, allowing the joining of all DNA parts which are in the BioBricks format.


The prefix contains the restriction sites for EcoRI, NotI and XBaI, while the suffix contains the SpeI, NotI and PstI restriction sites. Outside of the prefix and suffix regions, the DNA part must not contain these restriction sites. To join two BioBrick parts together, one of the plasmids is digested with EcoRI and SpeI while the second plasmid is digested with EcoRI and XbaI. The two EcoRI overhangs are complementary and will thus anneal together, while SpeI and XbaI also produce complementary overhangs which can also be ligated together. As the resulting plasmid contains the original prefix and suffix sequences, it can be used to join with more BioBricks parts. [ 24 ]  Because of this property, the BioBricks assembly standard is said to be  idempotent  in nature. However, there will also be a "scar" sequence (either TACTAG or TACTAGAG) formed between the two fused BioBricks. This prevents BioBricks from being used to create fusion proteins, as the 6bp scar sequence codes for a tyrosine and a stop codon, causing translation to be terminated after the first domain is expressed, while the 8bp scar sequence causes a  frameshift , preventing continuous readthrough of the codons. To offer alternative scar sequences that for example give a 6bp scar, or scar sequences that do not contain stop codons, other assembly standards such as the BB-2 Assembly, BglBricks Assembly, Silver Assembly and the Freiburg Assembly were designed. [ 25 ] [ 26 ] [ 27 ] [ 28 ]


While the easiest method to assemble BioBrick parts is described above, there also exist several other commonly used assembly methods that offer several advantages over the standard assembly. The 3 antibiotic (3A) assembly allows for the correct assembly to be selected via antibiotic selection, while the amplified insert assembly seeks to overcome the low transformation efficiency seen in 3A assembly. [ 29 ] [ 30 ]


The BioBrick assembly standard has also served as inspiration for using other types of endonucleases for DNA assembly. For example, both the iBrick standard and the HomeRun vector assembly standards employ homing endonucleases instead of type II restriction enzymes. [ 31 ] [ 32 ]


Some assembly methods also make use of type IIs restriction endonucleases. These differ from other type II endonucleases as they cut several base pairs away from the recognition site. As a result, the overhang sequence can be modified to contain the desired sequence. This provides Type IIs assembly methods with two advantages – it enables "scar-less" assembly, and allows for one-pot, multi-part assembly. Assembly methods that use type IIs endonucleases include  Golden Gate  and its associated variants.


The Golden Gate assembly protocol was defined by Engler et al. 2008 to define a DNA assembly method that would give a final construct without a scar sequence, while also lacking the original restriction sites. This allows the protein to be expressed without containing unwanted protein sequences which could negatively affect protein folding or expression. By using the BsaI restriction enzyme that produces a 4 base pair overhang, up to 240 unique, non-palindromic sequences can be used for assembly. [ 33 ]


Plasmid design and assembly


In Golden Gate cloning, each DNA fragment to be assembled is placed in a plasmid, flanked by inward facing BsaI restriction sites containing the programmed overhang sequences. For each DNA fragment, the 3' overhang sequence is complementary to the 5' overhang of the next downstream DNA fragment. For the first fragment, the 5' overhang is complementary to the 5' overhang of the destination plasmid, while the 3' overhang of the final fragment is complementary to the 3' overhang of the destination plasmid. Such a design allows for all DNA fragments to be assembled in a one-pot reaction (where all reactants are mixed together), with all fragments arranged in the correct sequence. Successfully assembled constructs are selected by detecting the loss of function of a screening cassette that was originally in the destination plasmid. [ 33 ]


MoClo and Golden Braid


The original Golden Gate Assembly only allows for a single construct to be made in the destination vector . To enable this construct to be used in a subsequent reaction as an entry vector, the MoClo and Golden Braid standards were designed. [ 34 ]


The MoClo standard involves defining multiple tiers of DNA assembly:


Each assembly tier alternates the use of BsaI and BpiI restriction sites to minimise the number of forbidden sites, and sequential assembly for each tier is achieved by following the Golden Gate plasmid design. Overall, the MoClo standard allows for the assembly of a construct that contains multiple transcription units, all assembled from different DNA parts, by a series of one-pot Golden Gate reactions. However, one drawback of the MoClo standard is that it requires the use of 'dummy parts' with no biological function, if the final construct requires less than four component parts. [ 35 ]  The Golden Braid standard on the other hand introduced a pairwise Golden Gate assembly standard.


The Golden Braid standard uses the same tiered assembly as MoClo, but each tier only involves the assembly of two DNA fragments, i.e. a pairwise approach. Hence in each tier, pairs of genes are cloned into a destination fragment in the desired sequence, and these are subsequently assembled two at a time in successive tiers. Like MoClo, the Golden Braid standard alternates the BsaI and BpiI restriction enzymes between each tier.


The development of the Golden Gate assembly methods and its variants has allowed researchers to design tool-kits to speed up the synthetic biology workflow. For example, EcoFlex was developed as a toolkit for  E. Coli  that uses the MoClo standard for its DNA parts, while a similar toolkit has also been developed for engineering the  Chlamydomonas reinhardtii  microalgae. [ 36 ] [ 37 ]


Site-specific recombination makes use of phage  integrases  instead of restriction enzymes, eliminating the need for having restriction sites in the DNA fragments. Instead, integrases make use of unique attachment (att) sites, and catalyse DNA rearrangement between the target fragment and the destination vector. The Invitrogen Gateway cloning system was invented in the late 1990s and uses two proprietary enzyme mixtures, BP clonase and LR clonase. The BP clonase mix catalyses the recombination between attB and attP sites, generating hybrid attL and attR sites, while the LR clonase mix catalyse the recombination of attL and attR sites to give attB and attP sites. As each enzyme mix recognises only specific att sites, recombination is highly specific and the fragments can be assembled in the desired sequence. [ 38 ]


Vector design and assembly


Because Gateway cloning is a proprietary technology, all Gateway reactions must be carried out with the Gateway kit that is provided by the manufacturer. The reaction can be summarised into two steps. The first step involves assembling the entry clones containing the DNA fragment of interest, while the second step involves inserting this fragment of interest into the destination clone.


The earliest iterations of the Gateway cloning method only allowed for only one entry clone to be used for each destination clone produced. However, further research revealed that four more orthogonal att sequences could be generated, allowing for the assembly of up to four different DNA fragments, and this process is now known as the Multisite Gateway technology. [ 39 ]


Besides Gateway cloning, non-commercial methods using other integrases have also been developed. For example, the Serine Integrase Recombinational Assembly (SIRA) method uses the ϕC31 integrase, while the Site-Specific Recombination-based Tandem Assembly (SSRTA) method uses the  Streptomyces  phage φBT1 integrase. [ 40 ] [ 41 ]  Other methods, like the HomeRun Vector Assembly System (HVAS), build on the Gateway cloning system and further incorporate homing endonucleases to design a protocol that could potentially support the industrial synthesis of synthetic DNA constructs. [ 31 ]


There have been a variety of long-overlap-based assembly methods developed in recent years. One of the most commonly used methods, the Gibson assembly method, was developed in 2009, and provides a one-pot DNA assembly method that does not require the use of restriction enzymes or integrases. [ 42 ]  Other similar overlap-based assembly methods include Circular Polymerase Extension Cloning (CPEC), Sequence and Ligase Independent Cloning (SLIC) and Seamless Ligation Cloning Extract (SLiCE). [ 43 ] [ 44 ] [ 45 ]  Despite the presence of many overlap assembly methods, the Gibson assembly method is still the most popular. [ 46 ]  Besides the methods listed above, other researchers have built on the concepts used in Gibson assembly and other assembly methods to develop new assembly strategies like the Modular Overlap-Directed Assembly with Linkers (MODAL) strategy, or the Biopart Assembly Standard for Idempotent Cloning (BASIC) method. [ 47 ] [ 48 ]


The Gibson assembly method is a relatively straightforward DNA assembly method, requiring only a few additional reagents: the 5' T5  exonuclease , Phusion  DNA polymerase , and  Taq   DNA ligase . The DNA fragments to be assembled are synthesised to have overlapping 5' and 3' ends in the order that they are to be assembled in. These reagents are mixed together with the DNA fragments to be assembled at 50 °C and the following reactions occur:


Because the T5 exonuclease is heat labile, it is inactivated at 50 °C after the initial chew back step. The product is thus stable, and the fragments assembled in the desired order. This one-pot protocol can assemble up to 5 different fragments accurately, while several commercial providers have kits to accurately assemble up to 15 different fragments in a two-step reaction. [ 49 ]  However, while the Gibson assembly protocol is fast and uses relatively few reagents, it requires bespoke DNA synthesis as each fragment has to be designed to contain overlapping sequences with the adjacent fragments and amplified via PCR. This reliance on PCR may also affect the fidelity of the reaction when long fragments, fragments with high GC content or repeat sequences are used. [ 48 ]


The MODAL strategy defines overlap sequences known as "linkers" to reduce the amount of customisation that needs to be done with each DNA fragment. The linkers were designed using the  R2oDNA Designer  software and the overlap regions were designed to be 45 bp long to be compatible with Gibson assembly and other overlap assembly methods. To attach these linkers to the parts to be assembled, PCR is carried using part-specific primers containing 15 bp prefix and suffix adaptor sequences. The linkers are then attached to the adaptor sequences via a second PCR reaction. To position the DNA fragments, the same linker will be attached to the suffix of the desired upstream fragment and the prefix of the desired downstream fragments. Once the linkers are attached, Gibson assembly, CPEC, or the other overlap assembly methods can all be used to assemble the DNA fragments in the desired order.


The BASIC assembly strategy was developed in 2015 and sought to address the limitations of previous assembly techniques, incorporating six key concepts from them: standard reusable parts; single-tier format (all parts are in the same format and are assembled using the same process); idempotent cloning; parallel (multipart) DNA assembly; size independence; automatability. [ 48 ]


DNA parts and linker design


The DNA parts are designed and cloned into storage plasmids, with the part flanked by an integrated prefix ( i P) and an integrated suffix ( i S) sequence. The  i P and  i S sequences contain inward facing BsaI restriction sites, which contain overhangs complementary to the BASIC linkers. [ 48 ]  Like in MODAL, the 7 standard linkers used in BASIC were designed with the R2oDNA Designer software, and screened to ensure that they do not contain sequences with homology to chassis genomes, and that they do not contain unwanted sequences like secondary structure sequences, restriction sites or ribosomal binding sites. Each linker sequence is split into two halves, each with a 4 bp overhang complementary to the BsaI restriction site, a 12 bp double stranded sequence and sharing a 21 bp overlap sequence with the other half. The half that is will bind to the upstream DNA part is known as the suffix linker part (e.g. L1S) and the half that binds to the downstream part is known as the prefix linker part (e.g. L1P). These linkers form the basis of assembling the DNA parts together.


Besides directing the order of assembly, the standard BASIC linkers can also be modified to carry out other functions. To allow for idempotent assembly, linkers were also designed with additional methylated  i P and  i S sequences inserted to protect them from being recognised by BsaI. This methylation is lost following transformation and in vivo plasmid replication, and the plasmids can be extracted, purified, and used for further reactions.


Because the linker sequence are relatively long (45bp for a standard linker), there is an opportunity to incorporate functional DNA sequences to reduce the number of DNA parts needed during assembly. The BASIC assembly standard provides several linkers embedded with RBS of different strengths. Similarly to facilitate the construction of fusion proteins containing multiple protein domains, several fusion linkers were also designed to allow for full read-through of the DNA construct. These fusion linkers code for a 15 amino acid glycine and serine polypeptide, which is an ideal linker peptide for fusion proteins with multiple domains.


Assembly


There are three main steps in the assembly of the final construct.


As DNA printing and DNA assembly methods have allowed commercial gene synthesis to become progressively and exponentially cheaper over the past years, [ 50 ]  artificial gene synthesis represents a powerful and flexible engineering tool for creating and designing new DNA sequences and protein functions. Besides synthetic biology, various research areas like those involving  heterologous gene expression ,  vaccine  development,  gene therapy  and molecular engineering, would benefit greatly from having fast and cheap methods to synthesise DNA to code for proteins and peptides. [ 51 ]  The methods used for DNA printing and assembly have even enabled the use of DNA as an  information storage medium .


On June 28, 2007, a team at the  J. Craig Venter Institute  published an article in  Science Express , saying that they had successfully transplanted the natural DNA from a  Mycoplasma mycoides  bacterium into a  Mycoplasma capricolum  cell, creating a bacterium which behaved like a  M. mycoides . [ 52 ]


On Oct 6, 2007,  Craig Venter  announced in an interview with UK's  The Guardian  newspaper that the same team had synthesized a modified version of the single  chromosome  of  Mycoplasma genitalium  artificially. The chromosome was modified to eliminate all genes which tests in live bacteria had shown to be unnecessary. The next planned step in this  minimal genome project  is to transplant the synthesized minimal genome into a bacterial cell with its old DNA removed; the resulting bacterium will be called  Mycoplasma laboratorium . The next day the Canadian  bioethics  group,  ETC Group  issued a statement through their representative,  Pat Mooney , saying Venter's "creation" was "a chassis on which you could build almost anything". The synthesized genome had not yet been transplanted into a working cell. [ 53 ]


On May 21, 2010,  Science  reported that the Venter group had successfully synthesized the genome of the bacterium  Mycoplasma mycoides  from a computer record, and transplanted the synthesized genome into the existing cell of a  Mycoplasma capricolum  bacterium that had its DNA removed. The "synthetic" bacterium was viable, i.e. capable of replicating billions of times. The team had originally planned to use the  M. genitalium  bacterium they had previously been working with, but switched to  M. mycoides  because the latter bacterium grows much faster, which translated into quicker experiments. [ 54 ]  Venter describes it as "the first species.... to have its parents be a computer". [ 55 ]  The transformed bacterium is dubbed " Synthia " by ETC. A Venter spokesperson has declined to confirm any breakthrough at the time of this writing.


As part of the Synthetic Yeast 2.0 project, various research groups around the world have participated in a project to synthesise synthetic yeast genomes, and through this process, optimise the genome of the model organism  Saccharomyces cerevisiae . [ 56 ]  The Yeast 2.0 project applied various DNA assembly methods that have been discussed above, and in March 2014,  Jef Boeke  of the Langone Medical Centre at New York University, revealed that his team had synthesized chromosome III of  S. cerevisiae . [ 57 ] [ 58 ]  The procedure involved replacing the genes in the original chromosome with synthetic versions and the finished synthetic chromosome was then integrated into a yeast cell. It required designing and creating 273,871 base pairs of DNA – fewer than the 316,667 pairs in the original chromosome. In March 2017, the synthesis of 6 of the 16 chromosomes had been completed, with synthesis of the others still ongoing. [ 59 ]


The  AT-hook  is a  DNA-binding   motif  present in many proteins, including the  high mobility group  (HMG)  proteins , [ 1 ]  DNA-binding  proteins  from  plants [ 2 ]  and hBRG1 protein, a central ATPase of the  human  switching/sucrose non-fermenting (SWI/SNF) remodeling  complex . [ 3 ]


This motif consists of a conserved, palindromic, core sequence of  proline - arginine - glycine - arginine - proline , although some AT-hooks contain only a single proline in the core sequence. AT-hooks also include a variable number of positively charged  lysine  and  arginine  residues on either side of the core sequence. [ 4 ]  The AT-hook binds to the  minor groove  of  adenine - thymine  (AT) rich DNA, hence the AT in the name. The rest of the name derives from a predicted  asparagine / aspartate  "hook" in the earliest AT-hooks reported in 1990. [ 5 ]  In 1997 structural studies using  NMR  determined that a DNA-bound AT-hook adopted a crescent or hook shape around the minor groove of a target DNA strand (pictured at right). [ 6 ]  HMGA proteins contain three AT-hooks, although some proteins contain as many as 30. [ 5 ]  The optimal binding sequences for AT-hook proteins are repeats of the form (ATAA) n  or (TATT) n , although the optimal binding sequences for the core sequence of the AT-hook are AAAT and AATT. [ 7 ]


The DNA dodecamer has eight consecutive AT base pairs, allowing the AT-hook to be positioned in several positions, with the preferred position being at one of the AATT regions to fully occupy the minor groove. Van der Waals interactions of the AT-hook with the adenines play an important role for the specificity of the position. [ 8 ]  Van der Waals interactions of the AT-hook with the adenines play an important role for the specificity of the position. [ 8 ]


The figure shows the position of the main chain to allow hydrogen bonds with the minor groove thymine oxygen atoms. The interactions shown, caused the DNA to bend, extending the minor groove. The distorted DNA causes the complementary major groove to form interactions between the side chains.


AT-hook proteins can form hydrogen bonds between NH groups of Gly37 and Arg38 on the main-chain and thymine oxygen atoms in the minor groove, which bends the DNA and widens the minor groove. [ 8 ]  The binding to the minor groove facilitates binding of other proteins in the major groove. [ 9 ]   That enables HMG proteins to regular expression of genes and influence biological processes.


The AT-hooks have also been proposed to anchor chromatin-modifying proteins to AT-rich DNA sequences through their association with chromatin remodeling, histone modifications, and chromatin insulator function. [ 9 ]


Alterations or abnormal expression of the HMG proteins have led to metabolic disorders, such as obesity, type 2 diabetes, and cancer. [ 8 ]


BioBrick  parts are  DNA  sequences which conform to a  restriction-enzyme  assembly standard. [ 1 ] [ 2 ]  These building blocks are used to design and assemble larger  synthetic biological circuits  from individual parts and combinations of parts with defined functions, which would then be incorporated into living cells such as  Escherichia coli  cells to construct new biological systems. [ 3 ]  Examples of BioBrick parts include  promoters ,  ribosomal binding sites (RBS) ,  coding sequences  and  terminators .


The BioBrick parts are used by applying engineering principles of abstraction and modularization. BioBrick parts form the base of the hierarchical system on which  synthetic biology  is based. There are three levels to the hierarchy:


The development of standardized biological parts allows for the rapid assembly of sequences. The ability to test individual parts and devices to be independently tested and characterized also improves the reliability of higher-order systems. [ 2 ]


The first attempt to create a list of standard biological parts was in 1996, by  Rebatchouk et al . This team introduced a cloning strategy for the assembly of short DNA fragments. However, this early attempt was not widely recognised by the scientific research community at the time. [ 2 ] [ 4 ]  In 1999, Arkin and Endy realized that the heterogeneous elements that made up a genetic circuit were lacking standards, so they proposed a list of standard biological parts. [ 5 ]  BioBricks were described and introduced by  Tom Knight  at  MIT  in 2003. [ 1 ]  Since then, various research groups have utilized the BioBrick standard parts to engineer novel biological devices and systems.


The BioBricks Foundation was formed in 2006 by engineers and scientists alike as a not-for-profit organization to standardize biological parts across the field. [ 6 ]  The Foundation focuses on improving in areas of Technology, Law, Education and the Global Community as they apply to  synthetic biology . BioBricks Foundation's activities include hosting SBx.0 Conferences, technical and educational programs. The SBx.0 conferences are international conferences on synthetic biology hosted across the world. Technical programs are aimed at the production of a series of standard biological parts, and their education expansion is creating acts which help create open, standardized sources of biological parts. [ 7 ]


As an alternative to traditional biotechnology patent systems and in an effort to allow BioBricks to be utilized as an open-source community standard, the BioBricks Foundation created the BioBrick Public Agreement, which consists of a Contributor Agreement and a User Agreement. Those who want to give a part to the community sign the Contributor Agreement, agreeing not to assert against Users Contributor-held intellectual property rights that might limit the use of the contributed materials. Signers of the User Agreement may freely use the whole collection of parts given by contributors. There is no requirement for users to contribute to the community in order to use the parts, and users may assert intellectual property rights to inventions developed by using the parts. [ 8 ]  The User Agreement allows users to establish invention of uses of parts, to disclose patents on parts combinations, and to freely build on the contributions of other users. [ 9 ] [ 10 ]


The BioBrick assembly standard was introduced to overcome the lack of standardization posed by traditional  molecular cloning  methods. The BioBrick assembly standard is a more reliable approach for combining parts to form larger composites. The assembly standard enables two groups of synthetic biologists in different parts of the world to re-use a BioBrick part without going through the whole cycle of design and manipulation. [ 2 ]  This means the newly designed part can be used by other teams of researchers more easily. Besides that, when compared to the old-fashioned  ad hoc  cloning method, the assembly standard process is faster and promotes automation. [ 11 ]  The BioBrick assembly standard 10 was the first assembly standard to be introduced. Over the years, several other assembly standards, such as the Biofusion standard and Freiburg standard have been developed.


Assembly standard 10 was developed by Tom Knight, and is the most widely used assembly standard. It involves the use of  restriction enzymes . Every BioBrick part is a DNA sequence which is carried by a  circular plasmid , which acts as a  vector . [ 12 ]  The vector acts as a transport system to carry the BioBrick parts. The first approach towards a BioBrick standard was the introduction of standard sequences, the prefix and suffix sequences, which flank the 5 ′  and 3 ′  ends of the DNA part respectively. [ 13 ]  These standard sequences encode specific restriction enzyme sites. The prefix sequence encodes  EcoRI  (E) and  Xbal  (X) sites, while the suffix sequence encodes  SpeI  (S) and  PstI  (P) sites. The prefix and the suffix  are not considered part of the BioBrick part. [ 3 ]  To facilitate the assembly process, the BioBrick part itself must not contain any of these restriction sites. During the assembly of two different parts, one of the plasmids is digested with  EcoRI  and  SpeI . The plasmid carrying the other BioBrick part is digested with  EcoRI  and  Xbal . This leaves both plasmids with 4 base pair (bp) overhangs at the 5 ′  and 3 ′  ends. The  EcoRI  sites will ligate since they are complementary to each other. The  Xbal  and  SpeI  sites will also ligate as the digestion produces compatible ends. Now, both the DNA parts are in one plasmid. The ligation produces an 8 base pair "scar" site between the two BioBrick parts. Since the scar site is a hybrid of the  Xbal  and  SpeI  sites, it is not recognized by either restriction enzyme. [ 13 ]  The prefix and suffix sequences remain unchanged by this digestion and ligation process, which allows for subsequent assembly steps with more BioBrick parts.


This assembly is an  idempotent  process: multiple applications do not change the end product, and maintain the prefix and suffix. Although the BioBrick standard assembly allows for the formation of functional modules, there is a limitation to this standard 10 approach. The 8 bp scar site does not allow the creation of a  fusion protein . [ 12 ]  The scar site causes a  frame shift  which prevents the continuous reading of codons, which is required for the formation of fusion protein.


Tom Knight later developed the BB-2 assembly standard in 2008 to address problems with joining the scars of protein domains and that the scars consist of eight bases, which will yield an altered reading frame when joining protein domains. The enzymes used for digestion of the initial parts are almost the same, but with modified prefixes and suffixes. [ 14 ]


The BglBrick assembly standard was proposed by J. Christopher Anderson, John E. Dueber, Mariana Leguia, Gabriel C. Wu, Jonathan C. Goler, Adam P. Arkin, and Jay D. Keasling in September 2009 as a standard very similar in concept to BioBrick, but enabling the generation of fusion proteins without altering the reading frame or introducing stop codons and while creating a relatively neutral amino acid linker scar (GlySer). A BglBrick part is as a DNA sequence flanked by 5 ′  EcoRI and BglII sites (GAATTCaaaA GATCT ) and 3 ′  BamHI and XhoI sites ( G GATCCaaaCTCGAG), and lacking in these same restriction sites internally. The upstream part in the pairwise assembly is purified from an EcoRI/BamHI digest, and the downstream part+vector is purified from an EcoRI/BglII digest. Ligation of these two fragments creates a composite part reforming the original flanking sites required in the part definition and leaving a  GGATCT  scar sequence at the junction of the parts, a scar that encodes the amino acids glycine and serine when fusing CDS parts together in-frame, convenient due to the GlySer dipeptide being a popular linker of protein domains. [ 2 ]


Pam Silver's lab created the Silver assembly standard to overcome the issue surrounding the formation of fusion protein. This assembly standard is also known as Biofusion standard, and is an improvement of the BioBrick assembly standard 10. Silver's standard involves deletion of one nucleotide from the  Xbal  and  SpeI  site, which shortens the scar site by 2 nucleotides, which now forms a 6 bp scar sequence. The 6 bp sequence allows the reading frame to be maintained. The scar sequence codes for the amino acid  threonine  (ACT) and  arginine  (AGA). [ 15 ]   This minor improvement allows for the formation of in-frame fusion protein. However, arginine's being a large, charged  amino acid  is a disadvantage to the Biofusion assembly technique: these properties of arginine result in the destabilisation of the protein by the  N-end rule .


The 2007 Freiburg iGEM team [ 16 ]  introduced a new assembly standard to overcome the disadvantages of the existing Biofusion standard technique. The Freiburg team created a new set of prefix and suffix sequences by introducing additional restriction enzyme sites,  AgeI  and  NgoMIV  to the existing prefix and suffix respectively. These newly introduced restriction enzyme sites are BioBrick standard compatible. The Freiburg standard still forms a 6 bp scar site, but the scar sequence (ACCGGC) now codes for threonine and  glycine  respectively. This scar sequence results in a much more stable protein [ 17 ]  as the glycine forms a stable N-terminal, unlike the arginine, which signals for N-terminal degradation. The assembly technique proposed by the Freiburg team  diminishes the limitations of the Biofusion standard.


Different methods are used when it comes to assembling BioBricks. This is because some standards require different materials and methods (use of different restriction enzymes), while others are due to preferences in protocol because some methods of assembly have higher efficiency and is user-friendly.


The 3A assembly method is the most commonly used, as its compatible with assembly Standard 10, Silver standard as well as the Freiburg standard. This assembly method involves two BioBrick parts and a destination plasmid. The destination plasmid contains a toxic (lethal) gene, to ease the selection of a correctly assembled plasmid. The destination plasmids also have different antibiotic resistance genes than the plasmids carrying the BioBrick parts. All three plasmids are digested with an appropriate restriction enzyme and then allowed to ligate. Only the correctly assembled part will produce a viable composite part contained in the destination plasmid. This allows a good selection as only the correctly assembled BioBrick parts survive.


The amplified insert assembly method does not depend on prefix and suffix sequences, allowing to be used in combination with a majority of assembly standards. It also has a higher transformation rate than 3A assembly, and it does not require the involved plasmids to have different antibiotic resistance genes. This method reduces noise from uncut plasmids by amplifying a desired insert using PCR prior to digestion and treating the mixture with the restriction enzyme  DpnI , which digests methylated DNA like plasmids. Eliminating the template plasmids with DpnI leaves only the insert to be amplified by PCR. To decrease the possibility of
creating plasmids with unwanted combinations of insert and backbone, the backbone can be treated with phosphatase to prevent its religation. [ 14 ]


The Gibson scarless assembly method allows the joining of multiple BioBricks simultaneously. This method requires the desired sequences to have an overlap of 20 to 150  bps . Because BioBricks do not have this overlap, this method requires PCR primers to create overhangs between adjacent BioBricks. T5 exonuclease attacks the 5 ′  ends of sequences, creating single-stranded DNA in the ends of all sequences where the different components are designed to anneal. DNA polymerase then adds DNA parts to gaps in the anneal components, and a Taq ligase can seal the final strands. [ 14 ]


The 4R/2M assembly method was designed to combine parts (BioBrick Assembly Standard 10 or Silver Standard) within existing plasmids (i.e. without PCR or subcloning). The plasmids are reacted in vivo with sequence-specific DNA methyltransferases, so that each is modified and protected from one of two restriction endonucleases that are later used to linearize undesired circular ligation products.  [ 18 ]


The MIT group led by Tom Knight that developed BioBricks and International Genetically Engineered Machines (iGEM) competition are also the pioneers of The Registry of Standard Biological Parts (Registry). [ 19 ]  Registry being one of the foundations of synthetic biology, provides web-based information and data on over 20,000 BioBrick parts. The Registry contains:


Every BioBrick part has its unique identification code which makes the search for the desired BioBrick part easier (for example, BBa_J23100, a constitutive promoter). [ 2 ]  The registry is open access, whereby anyone can submit a BioBrick part. Most of the BioBrick submission is from students participating in the annual iGEM competition hosted every summer. [ 20 ]  The Registry allows exchange of data and materials online which allows rapid re-use and modifications of parts by the participating community.


Professional parts registries have also been developed. Since most of the BioBrick parts are submitted by undergraduates as part of the iGEM competition, the parts may lack important characterisation data and metadata which would be essential when it comes to designing and modelling the functional components. [ 19 ]  One example of a professional parts registry is the USA-based publicly funded facility, The International Open Facility Advancing Biotechnology (BIOFAB), which contains detailed descriptions of each biological part. It is also an open-source registry, and is available commercially.  BIOFAB aims to catalogue high-quality BioBrick parts to accommodate the needs of professional synthetic biology community.


The BioBrick Foundation (BBF) is a public-benefit organization established to promote the use of standardized BioBrick parts on a scale beyond the iGEM competition. The BBF is currently working on the derivation of standard framework to promote the production high quality BioBrick parts which would be freely available to everyone. [ 21 ]


The  blue–white screen  is a  screening technique  that allows for the rapid and convenient detection of recombinant bacteria in  vector -based  molecular cloning  experiments.  This method of screening is usually performed using a suitable  bacterial strain , but other organisms such as yeast may also be used.  DNA  of transformation
is ligated into a  vector .  The vector is then  inserted  into a  competent host cell  viable for transformation, which are then grown in the presence of  X-gal . Cells transformed with vectors containing  recombinant DNA  will produce white colonies; cells transformed with non-recombinant plasmids (i.e. only the vector) grow into blue colonies.


Molecular cloning  is one of the most commonly used procedures in  molecular biology .  A gene of interest may be inserted into a plasmid vector via  ligation , and the plasmid is then transformed into  Escherichia coli  cells.  However, not all the plasmids transformed into cells may contain the desired gene insert, and checking each individual colony for the presence of the insert is time-consuming. Therefore, a method for the detection of the insert would be useful for making this procedure less time- and labor-intensive.  One of the early methods developed for the detection of insert is blue–white screening which allows for identification of successful products of cloning reactions through the colour of the  bacterial  colony.


The method is based on the principle of α-complementation of the  β-galactosidase  gene.  This phenomenon of α-complementation was first demonstrated in work done by  Agnes Ullmann  in the laboratory of  François Jacob  and  Jacques Monod , where the function of an inactive mutant β-galactosidase with deleted sequence was shown to be rescued by a fragment of β-galactosidase in which that same sequence, the α-donor peptide, is still intact. [ 1 ]   Langley  et al.  showed that the mutant non-functional  β-galactosidase  was lacking in part of its N-terminus with its residues 11—41 deleted, but it may be complemented by a peptide formed of residues 3—90 of β-galactosidase. [ 2 ]   M13 filamentous phage  containing sequence coding for the first 145 amino acid was later constructed by  Messing et al. , and α-complementation via the use of a vector was demonstrated by the formation of blue plaques when cells containing the inactive protein were infected by the phage and then grown in plates containing X-gal. [ 3 ]


The pUC series of plasmid  cloning vectors  by Vieira and Messing was developed from the M13 system and were the first plasmids constructed to take advantage of this screening method. [ 4 ]   In this method, DNA ligated into the plasmid disrupts the α peptide and therefore the complementation process, and no functional β-galactosidase can form.  Cells transformed with plasmid containing an insert therefore form white colonies, while cells transformed with plasmid without an insert form blue colonies; result of a successful ligation can thus be easily identified by the white coloration of cells formed from the unsuccessful blue ones. [ 5 ]


β-galactosidase  is a protein encoded by the  lacZ  gene of the  lac  operon , and it exists as a  homotetramer  in its active state.  However, a mutant β-galactosidase derived from the M15 strain of  E. coli  has its N-terminal residues 11—41 deleted and this mutant, the ω-peptide, is unable to form a tetramer and is inactive.  This mutant form of protein however may return fully to its active tetrameric state in the presence of an N-terminal fragment of the protein, the α-peptide.  The rescue of function of the mutant β-galactosidase by the α-peptide is called α-complementation.


In this method of screening, the host  E. coli  strain carries the  lacZ  deletion mutant ( lacZΔM15 ) which contains the ω-peptide, while the plasmids used carry the  lacZα  sequence which encodes the first 59 residues of β-galactosidase, the α-peptide.  Neither is functional by itself.  However, when the two peptides are expressed together, as when a plasmid containing the  lacZα  sequence is transformed into a  lacZΔM15  cells, they form a functional  β-galactosidase  enzyme.


The blue–white screening method works by disrupting this α-complementation process.  The plasmid carries within the  lacZα  sequence an internal  multiple cloning site  (MCS).  This MCS within the  lacZα  sequence can be cut by restriction enzymes so that the foreign DNA may be inserted within the  lacZ α gene, thereby disrupting the gene that produces α-peptide.  Consequently, in cells containing the plasmid with an insert, no functional  β-galactosidase  may be formed.


The presence of an active β-galactosidase can be detected by  X-gal , a colourless  analog  of lactose that may be cleaved by β-galactosidase to form 5-bromo-4-chloro-indoxyl, which then spontaneously dimerizes and oxidizes to form a bright blue insoluble pigment 5,5'-dibromo-4,4'-dichloro-indigo.  This results in a characteristic blue colour in cells containing a functional β-galactosidase.  Blue colonies therefore show that they may contain a vector with an uninterrupted  lacZα  (therefore no insert), while white colonies, where X-gal is not hydrolyzed, indicate the presence of an insert in  lacZα  which disrupts the formation of an active β-galactosidase.


The recombinant clones can be further analyzed by isolating and purifying small amounts of plasmid DNA from the transformed colonies and restriction enzymes can be used to cut the clone and determine if it has the fragment of interest. [ 6 ]  If the DNA is necessary to be sequenced, the plasmids from the colonies will need to be isolated at a point, whether to cut using restriction enzymes or performing other assays.


The correct type of  vector  and  competent cells  are important considerations when planning a blue–white screen.  The plasmid must contain the  lacZ α, and examples of such plasmids are  pUC19  and  pBluescript .  The  E. coli  cell should contain the mutant  lacZ  gene with deleted sequence (i.e.  lacZΔM15 ), and some of the commonly used cells with such genotype are JM109, DH5α, and XL1-Blue. It should also be understood that the lac operon is affected by the presence of glucose.  The protein  EIIA Glc , which is involved in glucose import, shuts down lactose permease when glucose is being transported into the cell.  The media used in agar plate therefore should not include glucose.


X-gal is light-sensitive and therefore its solution and plates containing X-gal should be stored in the dark.   Isopropyl β-D-1-thiogalactopyranoside  (IPTG), which functions as the inducer of the  lac  operon, may be used in the media to enhance the expression of LacZ.


X-gal  is an expensive material, thus other methods have been developed in order to screen bacteria.  GFP  has been developed as an alternative to help screen bacteria. The concept is similar to α-complementation in which a DNA insert can disrupt the coding sequence within a vector and thus disrupt the GFP production resulting in non-fluorescing bacteria. [ 7 ]  Bacteria that have recombinant vectors (vector + insert), will be white and not express the GFP protein, while non-recombinant (vector), will and fluoresce under UV light. GFP in general has been used as a reporter gene where individuals can definitively determine if a clone carries a gene that researchers are analyzing. On occasion, the medium in which the colonies grow can influence the screen and introduce false-positive results. [ 8 ]  X-gal on the medium can occasionally degrade to produce a blue color or GFP can lose its fluorescence because of the medium and can impact researchers capabilities to determine colonies with the desire recombinant and those that do not possess it. [ 9 ]


Some white colonies may not contain the desired recombinant plasmid for a number of reasons.  The ligated DNA may not be the correct one or not properly ligated, and it is possible for some linearized vector to be transformed, its ends "repaired" and ligated together such that no LacZα is produced and no blue colonies may be formed.  Mutation can also lead to the α-fragment not being expressed.  A colony with no vector at all will also appear white, and may sometimes appear as satellite colonies after the  antibiotic  used has been depleted. It is also possible that blue colonies may contain the insert. This occurs when the insert is "in frame" with the LacZα gene and a STOP codon is absent in the insert. This can lead to the expression of a fusion protein that has a functional LacZα if its structure is not disrupted. The correct recombinant construct can sometimes give lighter blue colonies which may complicate its identification.


Bromodeoxyuridine  ( 5-bromo-2'-deoxyuridine ,  BrdU ,  BUdR ,  BrdUrd ,  broxuridine ) is a synthetic  nucleoside analogue  with a chemical structure similar to  thymidine . BrdU is commonly used to study  cell proliferation  in living tissues [ 1 ]  and has been studied as a  radiosensitizer [ 2 ]  and  diagnostic tool  in people with  cancer . [ 3 ]


During the  S phase  of the  cell cycle  (when  DNA replication  occurs), BrdU can be incorporated in place of thymidine in newly synthesized  DNA  molecules of dividing cells. [ 4 ]  Cells that have recently performed DNA replication or  DNA repair  can be detected with  antibodies  specific for BrdU using techniques such as  immunohistochemistry  or  immunofluorescence . [ 5 ]  BrdU-labelled cells in humans can be detected up to two years after BrdU infusion. [ 6 ]


Because BrdU can replace  thymidine  during DNA replication, it can cause  mutations , and its use is therefore potentially a health hazard. [ citation needed ]  However, because it is neither  radioactive  nor  myelotoxic  at labeling concentrations, it is widely preferred for  in vivo  studies of  cancer  cell proliferation. [ 7 ] [ 8 ]   However, at  radiosensitizing  concentrations, BrdU becomes myelosuppressive, thus limiting its use for radiosensitizing. [ 2 ]


BrdU differs from  thymidine  in that BrdU substitutes a  bromine  atom for  thymidine's  CH 3  group.  The Br substitution can be used in X-ray diffraction experiments in crystals containing either DNA or RNA.  The Br atom acts as an anomalous scatterer and its larger size will affect the crystal's X-ray diffraction enough to detect  isomorphous  differences as well. [ 9 ] [ 10 ]


Bromodeoxyuridine releases gene silencing caused by DNA methylation. [ 11 ]


BrdU can also be used to identify microorganisms that respond to specific carbon substrates in aquatic [ 12 ]  and soil [ 13 ]  environments. A carbon substrate added to the incubations of environmental samples will cause the growth of microorganisms that can utilize that substrate. These microorganisms will then incorporate BrdU into their DNA as they grow. Community DNA can then be isolated and BrdU-labeled DNA purified using an immunocapture technique. [ 14 ]  Subsequent sequencing of the labeled DNA can then be used to identify the microbial taxa that participated in the degradation of the added carbon source.


However, it is not certain whether all microbes present in an environmental sample can incorporate BrdU into their biomass during  de novo  DNA synthesis. Therefore, a group of microorganisms may respond to a carbon source but go undetected using this technique. Additionally, this technique is biased towards identifying microorganisms with A- and T-rich genomes.


DNA with BrdU transcribes as usual DNA, with  guanine  included in RNA as a complement to BrdU. [ 15 ]


Calf-intestinal alkaline phosphatase  ( CIAP / CIP ) is a type of  alkaline phosphatase  that catalyzes the removal of   phosphate groups  from the 5' end of DNA strands and  phosphomonoesters  from RNA. [ 1 ] [ 2 ]  This enzyme is frequently used in DNA  sub-cloning , as DNA fragments that lack the 5' phosphate groups cannot  ligate . [ 3 ]  This prevents recircularization of the linearized DNA  vector  and improves the yield of the vector containing the appropriate insert.


Calf-intestinal alkaline phosphatase can serve as an effective tool for removing uranium from groundwater and soil that can pose major health risks. [ 4 ]  Furthermore, the toxicity of lipopolysaccharide (LPS) was mitigated by calf-intestinal alkaline phosphatase in mice and piglets, which indicates that it could be a promising new therapeutic agent for treating diseases associated with LPS. [ 5 ]


Cassette mutagenesis  is a type of  site-directed mutagenesis  that uses a short, double-stranded  oligonucleotide sequence  ( gene cassette ) to replace a fragment of target  DNA . It uses complementary  restriction enzyme digest ends  on the target DNA and  gene cassette  to achieve specificity. It is different from methods that use single  oligonucleotide  in that a single gene cassette can contain multiple mutations. Unlike many site directed mutagenesis methods, cassette mutagenesis also does not involve primer extension by  DNA polymerase . [ 1 ] [ 2 ] [ 3 ] [ 4 ] [ 5 ]


First,  restriction enzymes  are used to cleave near the target sequence on DNA contained in a suitable  vector . This step removes the target sequence and everything between the restriction sites. Then, the  synthetic double stranded DNA  containing the desired mutation and  ends  that are complementary to the  restriction digest ends  are  ligated  in place of the sequence removed. Finally, the resultant  construct  is  sequenced  to check that the target sequence contains the intended mutation. [ 1 ]


The use of synthetic gene cassette allows total control over the type of mutation that can be generated. When studying protein functions, cassette mutagenesis can allow a scientist to change individual  amino acids  by introducing different  codons  or omitting codons. [ 1 ] [ 2 ]


By including the  SD sequence  and the first few codons of a gene, a scientist can easily and dramatically affect the expression level of a protein by altering these  regulatory sequences . [ 2 ]


To use this method, the sequence of the target sequence and nearby restriction sites must be known. Since  restriction enzymes  are used, for this method to be useful, the restriction sites flanking the target DNA has to be unique in the  gene / vector  system so that the  gene cassette  can be inserted with specificity. The length of the sequence flanked by the restriction sites is also a limiting factor due to the use of synthetic gene cassettes. [ 2 ] [ 3 ]


Since one gene cassette can contain multiple mutations, less total oligonucleotide synthesis and purification is needed. Compared to mutagenesis methods that requires the synthesis of double stranded DNA using a single stranded template (1-30% in vitro in  M13 ), the efficiency of the ligation of oligodeoxynucleotide cassette is close to 100%. The high efficiency of the mutagenesis means mutants can be screened directly by sequencing. [ 2 ]  Once the vector is set up with flanking restriction sites, all manipulations (i.e.,  mutagenesis ,  sequencing ,  expression ) can be performed in the same  plasmid . [ 2 ]


Chargaff's rules  (given by  Erwin Chargaff ) state that in the  DNA  of any species and any organism, the amount of  guanine  should be equal to the amount of  cytosine  and the amount of  adenine  should be equal to the amount of  thymine . Further, a 1:1  stoichiometric  ratio of  purine  and  pyrimidine  bases (i.e.,  A+G=T+C ) should exist. This pattern is found in both strands of the DNA. They were discovered by Austrian-born chemist  Erwin Chargaff [ 1 ] [ 2 ]  in the late 1940s.


The first rule holds that a double-stranded  DNA  molecule,  globally  has percentage base pair equality: A% = T% and G% = C%. The rigorous validation of the rule constitutes the basis of  Watson–Crick base pairs  in the DNA double helix model.


The second rule holds that both Α% ≈ Τ% and G% ≈ C% are valid for each of the two DNA strands. [ 3 ]  This describes only a global feature of the base composition in a single DNA strand. [ 4 ]


The second parity rule was discovered in 1968. [ 3 ]  It states that, in single-stranded DNA, the number of adenine units is  approximately  equal to that of thymine (%A  ≈  %T), and the number of cytosine units is  approximately  equal to that of guanine (%C  ≈  %G).


The first empirical generalization of Chargaff's second parity rule, called the Symmetry Principle, was proposed by Vinayakumar V. Prabhu [ 5 ]  in 1993. This principle states that for any given oligonucleotide, its frequency is approximately equal to the frequency of its complementary reverse oligonucleotide. A theoretical generalization [ 6 ]  was mathematically derived by Michel E. B. Yamagishi and Roberto H. Herai in 2011. [ 7 ]


In 2006, it was shown that this rule applies to four [ 2 ]  of the five types of double stranded genomes; specifically it applies to the  eukaryotic   chromosomes , the  bacterial  chromosomes, the double stranded  DNA  viral genomes, and the  archaeal  chromosomes. [ 8 ]  It does not apply to  organellar genomes  ( mitochondria  and  plastids ) smaller than ~20-30  kbp , nor does it apply to single stranded DNA (viral) genomes or any type of  RNA  genome. The basis for this rule is still under investigation, although genome size may play a role.


The rule itself has consequences. In most bacterial genomes (which are generally 80-90% coding) genes are arranged in such a fashion that approximately 50% of the coding sequence lies on either strand.  Wacław Szybalski , in the 1960s, showed that in  bacteriophage  coding sequences  purines  (A and G) exceed  pyrimidines  (C and T). [ 9 ]  This rule has since been confirmed in other organisms and should probably be now termed " Szybalski's rule ". While Szybalski's rule generally holds, exceptions are known to exist. [ 10 ] [ 11 ] [ 12 ]  The biological basis for Szybalski's rule is not yet known.


The combined effect of Chargaff's second rule and Szybalski's rule can be seen in bacterial genomes where the coding sequences are not equally distributed. The  genetic code  has 64  codons  of which 3 function as termination codons: there are only 20  amino acids  normally present in proteins. (There are two uncommon amino acids— selenocysteine  and  pyrrolysine —found in a limited number of proteins and encoded by the  stop codons —TGA and TAG respectively.) The mismatch between the number of codons and amino acids allows several codons to code for a single amino acid—such codons normally differ only at the third codon base position.


Multivariate statistical analysis of codon use within genomes with unequal quantities of coding sequences on the two strands has shown that codon use in the third position depends on the strand on which the gene is located. This seems likely to be the result of Szybalski's and Chargaff's rules. Because of the asymmetry in pyrimidine and purine use in coding sequences, the strand with the greater coding content will tend to have the greater number of purine bases (Szybalski's rule). Because the number of purine bases will, to a very good approximation, equal the number of their complementary pyrimidines within the same strand and, because the coding sequences occupy 80–90% of the strand, there appears to be (1) a selective pressure on the third base to minimize the number of purine bases in the strand with the greater coding content; and (2) that this pressure is proportional to the mismatch in the length of the coding sequences between the two strands.


The origin of the deviation from Chargaff's rule in the organelles has been suggested to be a consequence of the mechanism of replication. [ 13 ]  During replication the DNA strands separate. In single stranded DNA,  cytosine  spontaneously slowly deaminates to  adenosine  (a C to A  transversion ). The longer the strands are separated the greater the quantity of deamination. For reasons that are not yet clear the strands tend to exist longer in single form in mitochondria than in chromosomal DNA. This process tends to yield one strand that is enriched in  guanine  (G) and  thymine  (T) with its complement enriched in cytosine (C) and adenosine (A), and this process may have given rise to the deviations found in the mitochondria.  [ citation needed ] [ dubious  –  discuss ]


Chargaff's second rule appears to be the consequence of a more complex parity rule: within a single strand of DNA any oligonucleotide ( k-mer  or  n-gram ; length ≤ 10) is present in equal numbers to its reverse complementary nucleotide. Because of the computational requirements this has not been verified in all genomes for all oligonucleotides. It has been verified for triplet oligonucleotides for a large data set. [ 14 ]  Albrecht-Buehler has suggested that this rule is the consequence of genomes evolving by a process of  inversion  and  transposition . [ 14 ]  This process does not appear to have acted on the mitochondrial genomes. Chargaff's second parity rule appears to be extended from the nucleotide-level to populations of codon triplets, in the case of whole single-stranded Human genome DNA. [ 15 ]
A kind of "codon-level second Chargaff's parity rule" is proposed as follows:


Examples — computing whole human genome using the first codons reading frame provides:


In 2020, it is suggested that the physical properties of the dsDNA (double stranded DNA) and the tendency to maximum entropy of all the physical systems are the cause of Chargaff's second parity rule. [ 16 ]  The symmetries and patterns present in the dsDNA sequences can emerge from the physical peculiarities of the dsDNA molecule and the maximum entropy principle alone, rather than from biological or environmental evolutionary pressure.


The following table is a representative sample of Erwin Chargaff's 1952 data, listing the base composition of DNA from various organisms and support both of Chargaff's rules. [ 17 ]  An organism such as φX174 with significant variation from A/T and G/C equal to one, is indicative of single stranded DNA.


Chromosome combing  (also known as molecular combing or DNA combing) [ 1 ]  is a technique used to produce an array of uniformly stretched  DNA  that is then highly suitable for  nucleic acid hybridization  studies such as  fluorescent in situ hybridisation  (FISH) which benefit from the uniformity of stretching, the easy access to the hybridisation target sequences, [ 2 ]  and the  resolution  offered by the large distance between two probes, which is due to the stretching of the DNA by a factor of 1.5 times the crystallographic length of DNA.


DNA in solution (i.e. with a randomly-coiled structure) is stretched by retracting the  meniscus  of the solution at a constant rate (typically 300 μm/s). The ends of DNA strands, which are thought to be frayed (i.e. open and exposing polar groups) bind to ionisable groups coating a  silanized  glass plate at a  pH  below the  pKa  of the ionizable groups (ensuring they are charged enough to interact with the ends of DNA). The rest of the DNA, which is mostly dsDNA, cannot form these interactions (aside from a few "touch down" segments along the length of the DNA strand) so is available for hybridisation to probes. As the meniscus retracts, surface retention creates a force that acts on DNA to retain it in the liquid phase; however this force is inferior to the strength of the DNA's attachment; the result is that the DNA is stretched as it enters the air phase; as the force acts in the locality of the air/liquid phase, it is invariant to different lengths or conformations of the DNA in solution, so DNA of any length will be stretched the same as the meniscus retracts. As this stretching is constant along the length of a DNA, distance along the strand can be related to base content; 1 μm is approximately equivalent to 2 kb.


DNA regions of interest are observed by hybridising them with probes labelled by  haptens  like  biotin ; this can then be bound by one or more layers of fluorochrome-associated ligands (such as immunofluorescence antibodies). Multicolour tagging is also possible. This has several potential uses, typically as a high-resolution physical mapping technique (e.g. for positional cloning), an example of which was the correct mapping of 200 kb of the CAPN3 gene region, or the mapping of non-overlapping sequences (since the distance between two probes can be accurately measured). It is therefore useful for finding exons, microdeletions, amplifications, or rearrangements. Before the combing improvement, FISH was too low-res to be of use in this case. With this technique, the resolution of FISH is theoretically limited only by the resolution of the  epifluorescence microscope ; in practice, resolutions of around 2 μm are obtained, for DNA molecules usually 200–600 kb long (though combing-FISH has been used with some success on molecules in excess of 1 Mb long), and there may be room for improvement through optimisation. Since DNA analyses using this technique are single-molecule, genomes from different cells can be compared to find anomalies, with implications for diagnosis of cancer and other genetic alterations.


Chromosome combing is also used to study  DNA replication , a highly regulated process that is reliant on a specific program of temporal and spatial distribution of activation of  origins of replication . Each origin occupies a distinct genetic locus and must fire only once per  cell cycle . Chromosome combing allows a genome-wide view of the firing of origins and propagation of  replication forks . As no assumptions are made about the sequence of the origins, this technique is particularly useful for mapping origins in  eukaryotes , which are not thought to have precisely defined initiation sequences.


Strategies involving combing recently replicated DNA typically involve incorporating modified nucleotides (such as BrdU,  bromodeoxyuridine ) into the nascent DNA, then fluorescently detecting it. As replication forks spread bidirectionally from origins of replication at (approximately) equal speeds, [ 3 ]  then origin position can be inferred. Replacing the modified nucleotide pool with a different type of modified nucleotide after a certain amount of time allows development of a time-resolved picture of the firing of sites, and the kinetics of replication forks. Pause sites can be identified, merged replication forks resolved, and the frequency of origin firings in different time periods to be studied.


Firing frequencies have shown in  in vitro  studies of Xenopus laevis egg extract to increase as S phase progresses. In another study [ 4 ]  on Epstein-Barr virus  episomes , hybridised probes were used to visualise the regional distribution of firing events; a particular zone showed preference for firing, whilst a few pause sites were also inferred.


Chromosome combing is performed by the company Genomic Vision, based in Paris.


Cross-linking and immunoprecipitation  ( CLIP, or CLIP-seq ) is a method used in  molecular biology  that combines  UV   crosslinking  with  immunoprecipitation  in order to identify  RNA  binding sites of  proteins  on a transcriptome-wide scale, thereby increasing our understanding of post-transcriptional regulatory networks. [ 1 ] [ 2 ] [ 3 ]  CLIP can be used either with antibodies against endogenous proteins, or with common peptide tags (including FLAG, V5, HA, and others) or affinity purification, which enables the possibility of profiling model organisms or RBPs otherwise lacking suitable antibodies. [ 4 ]


CLIP begins with the  in vivo  cross-linking of RNA-protein complexes using ultraviolet light (UV). Upon UV exposure, covalent bonds are formed between proteins and nucleic acids that are in close proximity (on the order of Angstroms apart). [ 5 ]  The cross-linked cells are then lysed, RNA is fragmented, and the protein of interest is isolated via immunoprecipitation. In order to allow for priming of reverse transcription, RNA adapters are ligated to the 3' ends, and RNA fragments are labelled to enable the analysis of the RNA-protein complexes after they have been separated from free RNA using gel electrophoresis and membrane transfer.  Proteinase K  digestion is then performed in order to remove protein from the crosslinked RNA, which leaves a few amino acids at the crosslink site. This often leads to truncation of cDNAs at the crosslinked nucleotide, which is exploited in variants such as  iCLIP  to increase the resolution of the method. [ 6 ]  cDNA is then synthesized via RT-PCR followed by high-throughput sequencing followed by mapping the reads back to the transcriptome and other computational analyses to study the interaction sites. [ 2 ]


CLIP was originally undertaken to study interactions between the neuron-specific  RNA-binding protein  and  splicing factors   NOVA1  and NOVA2 in the  mouse brain , identifying RNA binding sites that contained the expected Nova-binding motifs. Sequencing of the cDNA library identified many positions  close to alternative exons, several of which were found to require Nova1/2 for their brains-specific splicing patterns. [ 1 ]  In 2008, CLIP was combined with high-throughput sequencing (termed "HITS-CLIP") to generate genome-wide protein-RNA interaction maps for Nova; [ 7 ]  since then a number of other RNA-binding proteins have been studied with CLIP, including PTBP1, [ 8 ]  RbFox2 (where it was referred to as "CLIP-seq"), [ 9 ]  SFRS1, [ 10 ]  Argonaute, [ 11 ] [ 12 ] [ 13 ]  hnRNP C, [ 6 ]  the Fragile-X mental retardation protein FMRP, [ 14 ] [ 15 ]  Ptbp2 (in the mouse brain), [ 16 ]  Mbnl2, [ 17 ]  the nElavl proteins (the neuron-specific Hu proteins), [ 18 ]  and has been applied to RNA binding proteins from all kingdoms of life, including prokaryotes. [ 19 ]  CLIP analysis of the RNA-binding protein  Argonaute  led to identification of microRNA targets [ 20 ]  by decoding  microRNA -mRNA and protein-RNA interaction maps in the mouse brain [ 11 ] [ 21 ]  and subsequently in budding yeast ( Saccharomyces cerevisiae ) , [ 22 ]   Caenorhabditis elegans , [ 23 ]   embryonic stem cells [ 24 ]  and tissue culture cells. [ 25 ]


HITS-CLIP  combines UV  cross-linking  and  immunoprecipitation  with  high-throughput sequencing  to identify binding sites of RNA-binding proteins. [ 5 ]  HITS-CLIP also introduced the addition of dinucleotide barcodes to primers, providing the ability to sequence and then deconvolute multiple experiments simultaneously. [ 7 ]  With analysis of cross-linking induced mutation sites (CIMS) at  high sequencing depths , crosslink sites can be differentiated from other sources of sequence variation. [ 28 ]


PAR-CLIP  (photoactivatable ribonucleoside–enhanced cross-linking and immunoprecipitation) is also used for identifying the binding sites of cellular RNA-binding proteins (RBPs) and microRNA-containing ribonucleoprotein complexes (miRNPs). [ 25 ]  The method relies on the incorporation of photoreactive ribonucleoside analogs, such as 4-thiouridine (4-SU) and 6-thioguanosine (6-SG) into nascent RNA transcripts by living cells. Irradiation of the cells by UV light of 365 nm induces efficient cross-linking of photoreactive nucleoside-labeled cellular RNAs to interacting RBPs. Immunoprecipitation of the RBP of interest is followed by the isolation of the cross-linked and co-immunoprecipitated RNA. The isolated RNA is converted into a cDNA library and deep sequenced using  high-throughput sequencing  technology. Cross-linking the 4-SU and 6-SG analogs results in thymidine to cytidine, and guanosine to adenosine transitions respectively. As a result, PAR-CLIP can identify binding site locations with high accuracy.


However, PAR-CLIP is limited mainly to cultured cells, and nucleoside cytotoxicity is a concern; [ 2 ]  it has been reported that 4-SU inhibits ribosomal RNA synthesis, induces a nucleolar stress response, and reduces cell proliferation. [ 29 ]  PAR-CLIP has been employed to determine the transcriptome-wide binding sites of several known RBPs and microRNA-containing ribonucleoprotein complexes at high resolution. This includes the miRNA targeting AGO and TNRC6 proteins. [ 21 ]


iCLIP  (individual nucleotide–resolution crosslinking and immunoprecipitation) is a variant of CLIP that enabled amplification of truncated cDNAs, which are produced when reverse transcription stops prematurely at the cross-link site. [ 6 ]  Other approaches to identify protein-RNA crosslink sites include mutational analysis of read-through cDNAs, such as nucleotide transitions in  PAR-CLIP , [ 25 ]  or rare errors introduced by reverse transcriptase when it reads through the crosslink sites in standard  HITS-CLIP  methods, termed Crosslink induced mutation site (CIMS) analysis. [ 30 ]


iCLIP also added a random sequence (unique molecular identifier,  UMI ) along with experimental barcodes to the primer used for reverse transcription, thereby barcoding unique cDNAs to minimise any errors or quantitative biases of PCR, and thus improving the quantification of binding events. Enabling amplification of truncated cDNAs led to identification of the sites of RNA-protein interactions at high resolution by analysing the starting position of truncated cDNAs, as well as their precise quantification using UMIs with software called " iCount ". These innovations of iCLIP were adopted by later variants of CLIP such as eCLIP and irCLIP. [ 4 ]  Another modification of iCLIP, miCLIP, identifies methylated RNA sites with use of mutant enzyme or modification-specific antibody. [ 31 ] [ 32 ] [ 2 ]  The quantitative nature of iCLIP enabled  comparison across samples at the level of full RNAs, [ 33 ]  or to study competitive binding of multiple RNA-binding proteins [ 34 ]  or subtle changes in binding of a mutant protein at the level of binding peaks. [ 35 ]


eCLIP (enhanced CrossLinking and ImmunoPrecipitation followed by high-throughput sequencing) is also used to map RBP binding sites on RNAs transcriptome-wide. [ 36 ]  eCLIP was designed to improve upon iCLIP by increasing the efficiency in converting purified RNA fragments into cDNA library. At its publication, eCLIP was reported to increase such efficiency by >1000-fold, which not only decreases wasted sequencing of PCR duplicate molecules, but also dramatically decreases experimental failures during the CLIP procedure. Additionally, the amplification in eCLIP is now comparable to RNA-seq, enabling rigorous quantitative normalization against paired input controls (to remove background at ribosomal and other highly abundant RNAs) as well as quantitative comparison across peaks and samples, enabling the ability to detect allele-specific binding or differential RNA binding between conditions. As in other CLIP methods, eCLIP relies on RBP-RNA interactions covalently linked using UV crosslinking of live cells. Cells are then lysed, and RNA is fragmented using limited RNase treatment. A specific RBP (and its bound RNA) is then immunoprecipitated using an antibody that specifically recognizes the targeted RBP. After ligation of a 3’ RNA adapter, immunoprecipitated material (as well as a paired input sample) are run on denaturing protein gels and transferred to nitrocellulose membranes. A region from the protein size to 75 kDa above is cut from the membrane and treated with Proteinase K to release RNA. After cleanup, RNA is then reverse transcribed to ssDNA, when a second adapter is ligated. By ligating the second adapter to cDNAs, eCLIP can identify truncated cDNAs, similar to iCLIP, and thereby study RNA-protein interaction sites with high resolution. PCR amplification is then used to obtain sufficient material for high-throughput sequencing. eCLIP can also be used to identify miRNA targets and profile RNA modifications such as m6A. eCLIP datasets have been produced for over 150 RBPs with validated commercially available antibodies. [ 37 ]


sCLIP (simple CLIP) is a technique that requires lower amounts of input RNA and omits radio-labeling of the immunoprecipitated RNA. The method is based on linear amplification of the immunoprecipitated RNA and thereby improves the complexity of the sequencing-library despite significantly reducing the amount of input material and omitting several purification steps. Additionally, it permits a radiolabel-free visualization of immunoprecipitated RNA by using a highly sensitive biotin-based labeling technique. Along with a bioinformatical platform this method is designed to provide deep insights into RNA–protein interactomes in biomedical science, where the amount of starting material is often limited (i.e. in case of precious clinical samples). [ 38 ]  Additional iCLIP variants have also been developed that retain the individual nucleotide resolution but differ in one or more steps from the original iCLIP method. These include iCLIP2, irCLIP, iiCLIP, and iCLIP1.5, a few to name.


As a modification of CLIP, methylated RNA sites were identified with the use of mutant enzyme or modification-specific antibody with the methods termed miCLIP or m6A-CLIP. [ 31 ] [ 32 ] [ 39 ] [ 2 ]


RNA-binding proteins are frequently components of multi-protein complexes, and RNAs from various genes are present in cells at a range of abundance, therefore it is common that RNAs bound to co-purified proteins or non-specifically sticking to beads may be isolated when immunoprecipitating a specific protein. The data specificity obtained using early immunoprecipitation methods such as  RIP  have been demonstrated to be dependent on the reaction conditions of the experiment, such as protein concentrations and ionic conditions, and reassociation of RNA-binding proteins following cell lysis could lead to detection of artificial interactions. [ 40 ]  Formaldehyde crosslinking methods have been used to preserve RNA-protein interactions, but these also generate protein-protein cross-links. By employing UV crosslinking that is specific to direct protein-RNA contacts, CLIP avoid protein-protein cross-links and ensures high specificity, while also obtaining positional information on the sites of protein-RNA interactions.


Since UV crosslinking creates a covalent bond, the crosslinked RNA fragments retain a short peptide after Proteinase K digestion, which can be exploited to identify the crosslink site. Reverse transcription most often truncates at the crosslink sites, creating truncated cDNAs that are exploited by iCLIP, while read-through cDNAs often contain mutations at the crosslink site (see HITS-CLIP and PAR-CLIP). [ 2 ]


All CLIP library generation protocols require moderate quantities of cells or tissue (50–100 mg), require numerous enzymatic steps, and customised computational analyses. [ 12 ] [ 41 ]  Certain steps are difficult to optimize and frequently have low efficiencies. For example, overdigestion with RNase can decrease the number of identified binding sites and thus needs to be optimised. [ 27 ]  Crosslinking efficiency also varies between proteins, [ 42 ]  and nucleotide bias of crosslinking has been reported, [ 43 ]  for example by comparing cross-linking sites and motifs enriched when protein-RNA complexes are studied  in vivo  in living cells and  in vitro , [ 44 ]  though methods are being developed to minimise such bias for enriched motif discovery. [ 45 ]  Computationally predicted miRNA targets derived from  TargetScan  are comparable to CLIP in identifying miRNA targets, raising questions as to its utility relative to existing predictions. [ 46 ]  Because CLIP methods rely on immunoprecipitation, crosslinked RNA could in some cases affect antibody-epitope interactions. Finally, significant differences have been observed. Therefore, raw CLIP results require further computational analyses to thoroughly investigate RNA-protein binding site interactions within the cell.


PAR-CLIP   [ 1 ]  ( photoactivatable ribonucleoside-enhanced crosslinking and immunoprecipitation ) is a  biochemical  method for identifying the binding sites of cellular  RNA-binding proteins  (RBPs) and  microRNA -containing ribonucleoprotein complexes (miRNPs). The method relies on the incorporation of  ribonucleoside  analogs that are  photoreactive , such as 4-thiouridine (4-SU) and 6-thioguanosine (6-SG), into nascent RNA transcripts by living cells. Irradiation of the cells by  ultraviolet  light of 365 nm  wavelength  induces efficient  crosslinking  of photoreactive nucleoside– labeled  cellular RNAs to interacting RBPs.  Immunoprecipitation  of the RBP of interest is followed by isolation of the crosslinked and coimmunoprecipitated RNA. The isolated RNA is converted into a  cDNA  library and is  deep sequenced  using  next-generation sequencing  technology. [ 1 ] [ 2 ]


Recently,  PAR-CLIP  have been applied to determine the transcriptome-wide binding sites of several known RBPs and microRNA-containing ribonucleoprotein complexes at high resolution. [ 1 ] [ 3 ] [ 4 ] [ 5 ]


Community fingerprinting  is a set of  molecular biology  techniques that can be used to quickly profile the  diversity  of a  microbial  community. Rather than directly identifying or counting individual cells in an environmental sample, these techniques show how many variants of a  gene  are present. In general, it is assumed that each different gene variant represents a different type of microbe. Community fingerprinting is used by  microbiologists  studying a variety of microbial systems (e.g. marine, freshwater, soil, and human microbial communities) to measure biodiversity or track changes in community structure over time. The method analyzes environmental samples by assaying genomic  DNA . This approach offers an alternative to  microbial culturing , which is important because most microbes cannot be cultured in the laboratory. [ 1 ]  Community fingerprinting does not result in identification of individual microbe species; instead, it presents an overall picture of a microbial community.  These methods are now largely being replaced by high throughput sequencing, such as targeted  microbiome  analysis (e.g., 16s rRNA sequencing) and  metagenomics .


A fingerprinting analysis begins with an environmental sample (e.g. seawater or soil), from which total DNA is extracted. (Total DNA contains a mix of genetic material from all the microbes present in the sample.) A particular gene or DNA region is then selected as a target for analysis, under the assumption that each microbe species will have a different gene variant (also called a " phylotype "). Different methods (see below) can be used to visualize the phylotypes present in a sample. Because the aim of community fingerprinting is to gain an overall understanding of community structure, it is a particularly useful technique for analyzing  time-series  data collected from the field. For example, one could study the pattern of microbial  succession  in a habitat, or one could examine the response of a microbial community to an environmental perturbation, such as the release of a pollutant. Depending on what information is desired, different genes may be targeted. The most common are small subunit  ribosomal RNA  (rRNA) genes, such as  16S rRNA . These genes are frequently used in microbial  phylogenetic  analyses, so well-established techniques exist for their study. Other genes of interest might be those that are key in various metabolic processes. [ 1 ]


The advantages of community fingerprinting are that it can be performed quickly and relatively cheaply, and the analyses can accommodate a large number of samples simultaneously. [ 2 ] [ 3 ]  These properties make community fingerprinting especially useful for monitoring changes in microbial communities over time. Also, fingerprinting techniques do not require one to have  a priori  sequence data for organisms in a sample.
A disadvantage of community fingerprinting is that it results in largely qualitative, not quantitative data. [ 4 ]  When using qualitative data, it can be difficult to compare patterns observed in different studies or between different investigators.  Also, community fingerprinting does not directly identify taxa in an environmental sample, though the data output from certain techniques (e.g.  DGGE ) can be analyzed further if one desires identification. Some authors point to poor reproducibility of results for certain fingerprinting methods, [ 3 ]  while other authors have criticized the inaccuracy of abundance estimates and the inability of some techniques to capture the presence of rare taxa. [ 5 ]  For example, it is difficult for the DGGE method to detect microbes that comprise less than 0.5%-1% of a bacterial community. [ 6 ]


This section presents three methods of community fingerprinting.


Terminal restriction fragment length polymorphism  (T-RFLP) is a method that uses  fluorescently -labeled DNA fragments to produce a community fingerprint. [ 2 ] [ 7 ]  This section presents a brief explanation of T-RFLP  in the specific context of community fingerprinting . For a more detailed explanation, refer to the  T-RFLP  article.


To perform T-RFLP (Figure 1), one must select a target gene (e.g. the 16S rRNA gene) to amplify by  PCR . At least one  primer  used in the PCR reaction is fluorescently labeled at the  5´  end. After PCR amplification, each copied DNA segment carries the fluorescent label. Next,  restriction enzymes  are used to cut the amplified DNA at specific  recognition sites . The underlying assumption is that each microbe in the sample will have a different sequence on the target gene, so a restriction enzyme will cut each microbe's DNA in a different place. (Each different restriction site is considered to represent a single  operational taxonomic unit  [OTU]). Thus, the enzyme will produce one fragment length for each type of microbe present in the sample. The result of digestion is a set of restriction fragments of different lengths, each of which is fluorescently labeled at one end. These are known as "terminal fragments" because they are labeled at the end where the PCR primer attached.  (The unlabeled ends are not recorded in the final analysis.) Next, the fragments are separated by size through either  gel  or  capillary electrophoresis . Laser detection captures the size and fluorescence-intensity patterns of the terminal fragments. DNA standards of known size and fluorescence are included in the analysis as references. By setting a minimum threshold for fluorescence, background noise will be excluded. [ 4 ]


The output of the laser detection step is an  electropherogram  that shows a series of peaks, with fragment length on the horizontal axis and fluorescence intensity on the vertical axis (Figure 1). A second output [ 4 ]  is a data table that lists the migration time of the fragments, the size in  base pairs  of each peak, and the height of and area under each peak.


The theoretical basis of T-RFLP assumes that peaks at different positions along the horizontal axis represent different types of organisms (or OTUs). The area under each fluorescence intensity peak is a proxy for relative abundance of each phylotype in the community. However, a number of caveats must be taken into account. Different types of organisms may share a restriction site in the gene of interest; if that is the case, these organisms would not be distinguished as different peaks on the electropherogram. Furthermore, area under a peak represents relative abundance rather than absolute abundance, and there are biases in abundance measurement and PCR amplification. For example, organisms that are scarce in the original total DNA sample will not be amplified enough to be detected in the final analysis. [ 7 ]  This leads to underestimation of community diversity. Liu  et al. [ 7 ]  cite other possible factors that may distort results, including "differences in gene copy number between species and biases introduced during cell lysis, DNA extraction, and PCR amplification" (p. 4521). For those who seek detailed technical information, Marsh [ 2 ]  provides a catalog of potential biases that could be introduced in each step of the T-RFLP process.


The major advantages of T-RFLP are that it is fast and can easily accommodate many samples. [ 2 ]  Also, the visual output simplifies comparison of community structure patterns across different samples. A disadvantage is the broad, qualitative nature of the data output, which must be interpreted with the above caveats in mind. Also, direct identification of microbes in a sample is not possible through T-RFLP.


Disayathanoowat  et al. [ 8 ]  used T-RFLP to assess the microbial gut community, or microbiome, in two species of honeybees in Thailand. They found that the two species harbor different microbial communities and that the microbiome changes over the lifetime of the bees.


Joo  et al. [ 9 ]   tested T-RFLP as a method for phytoplankton monitoring. The authors collected environmental water samples from reservoirs in a time series. After comparison of samples with known terminal restriction fragments (from a database built from cultures), they concluded that T-RFLP can be used effectively as a technique for monitoring  changes  in the phytoplankton community. However, diversity and abundance estimates were found to be less accurate than those found through other methods.


Denaturing gradient gel electrophoresis (DGGE)  is a microbial fingerprinting technique that separates  amplicons  of roughly the same size based on sequence properties [ 1 ]  (Figure 2). These properties dictate the threshold at which DNA denatures. The DGGE gel uses a gradient DNA denaturant (a mixture of urea and formamide), or a linear temperature gradient. [ 10 ]  When the fragment reaches its melting point (threshold of enough denaturant), it stops moving. This is due to the fact that a partially melted double-stranded DNA can no longer migrate through the gel. [ 11 ]  A GC clamp (about 40 bases with high GC content) is used as a special  primer  to anchor the  PCR  fragments together once they have  denatured . [ 12 ]


Each lane on a gel represents one microbial community. The shared bands among the samples are the same size and roughly the same position on the gel. The gene variants that are not shared amongst microbial community samples do not match up horizontally with others. For example, if the gene of interest is 16S rRNA, as it was when the technique was first described, the PCR-amplified fragments will be in the same vertical location because they are all roughly the same size. [ 13 ]  Another target gene may have greater variation in length, but the denaturant gradient uses a second element (of melting point) to further distinguish between the samples. The DGGE gel will separate genes of the same size based on base sequence.


This technique shows to what extent microbial communities are the same or different in taxonomic composition. Each band in a different location on the gel represents a different phylotype (one unique sequence of a phylogenetic marker gene). [ 1 ]  For microbial communities this method profiles many individual 16S rRNA sequences. [ 14 ]  The number of bands at differing horizontal positions can be used to estimate the level of biodiversity in that sample and infer phylogenetic affiliation. [ 10 ]  In order to know more about phylogenetic affiliation, one could excise those bands from the gel and then sequence them.


The use of denaturing profile serves as a way to separate DNA fragments of similar sizes. This is beneficial in assessing microbial diversity due to the fact that the 16S rRNA gene does not vary much in size across bacterial phyla. The DGGE gel provides a quick way of looking at biodiversity in a microbial sample and does not preclude the option of sequencing the bands of interest. This method does not require that the microbes be cultured in the lab and does not require any sequence data needed to design probes for hybridization methods. [ 10 ]  The main disadvantage is that this is a qualitative assessment of biodiversity and one must sequence the genes in order to make inferences about the phylogenetic relatedness. Another disadvantage is that the GC clamp can be variable each time it is synthesized. This leads to the potential for different DGGE profiles for the same 16S rRNA sequence. [ 12 ]


Stephen  et al. [ 15 ]  utilized DGGE for a rapid analysis of Proteobacteria in soil. They obtained an initial assessment of microbial diversity in their environmental samples from soil maintained for 36 years at the various pH values. They combined DGGE and hybridization techniques by probing the DNA fragments to obtain more detail about the natural populations. In this study, they were looking at a group of closely related bacterial types, all autotrophic β–proteobacterial ammonia oxidizers. The 16S rDNA samples yielded ambiguous overlapping bands when run out on a gel. The ambiguous overlapping bands were separated with cluster-specific radiolabelled probes, which yielded information of the relative abundance of the different genotypes in samples.


Ward  et al. [ 14 ]  examined cyanobacterial mat communities in a Yellowstone hot spring by using DGGE analysis of 16S rRNA gene segments of aerobic chemoorganotrophic populations. DGGE allowed them to profile the community gaining new insight to diversity. They characterized the bands of interest by purifying and sequencing and detected many lineages previously unknown to be present.


(Automated)  ribosomal intergenic spacer analysis , or (A)RISA (Figure 3), takes advantage of the fact that prokaryotic DNA encodes for two highly conserved genes, the 16SrRNA gene and the 23SrRNA gene. These encode the small and large subunit genes in the rRNA operon. Between these two genes, there is an internal transcribed spacer (ITS) region. Due to the fact it is non-coding for proteins, it is a highly variable nucleotide sequence and length. Once DNA is isolated from a community, PCR amplifies this spacer region. The fragments can be run out on a gel (RISA), or the fluorescent primers can be translated into peaks in abundance of the different fragments lengths on an  electropherogram  (ARISA). [ 16 ] [ 17 ]


Due to variable non-coding regions, the output for RISA is a gel with different banding patterns, and output for ARISA is an electropherogram  with different peaks (similar to T-RFLP).


The brightness of the fluorescently labeled primers correlates to how prevalent that bacterial type is in the community. The banding pattern on the gel can be interpreted as a community-specific profile. Each DNA band or peak indicates at least one representative of that organism. [ 16 ]  In RISA, the bands on gel that do not match up in length represent different organisms in the community because they have different spacer regions between the two highly conserved genes. The electropherogram shows peaks correlating to the relative abundance of that spacer region in the sample.


ARISA can have a higher resolution in detecting microbial diversity as compared to  T-RFLP . [ 18 ]  This fingerprinting method is a quick and sensitive method to estimate microbial diversity. The observed length heterogeneities can be compared to databases for overlap with culturable organisms. [ 16 ]  One can design phylum-level oligonucleotide primers to get at questions regarding phylogenetic groups. [ 16 ]  One disadvantage to ARISA is the fact that a single organism may contribute more than one peak to the community profile. Unrelated organisms can also have similar spacer lengths, which leads to underestimates of community diversity. Due to these biases, researchers often use this method on multiple samples from each community in order to get an average assessment.


Ranjard  et al. [ 17 ]  discuss several examples of types of studies in which RISA can be used. They cite several studies that have used this technique to fingerprint bacterial communities following such disturbances as antibiotic treatment, mercury stress and deforestation. They also demonstrated the successful use of ARISA for characterizing fungal communities, which is an aspect of microbial ecology that remains to be fully explored.


Schloss  et al. [ 19 ]  conducted a study examining environmental variables and linking them to changes in the microbial ecology of a compost pile. They used ARISA to profile the community structure and look at microbial succession over stages in the composting process. They took DNA samples and sequenced the 16S rRNA gene to identify community members at the different phases of the process. Then they used ARISA to look at community-wide changes and then matched up the 16S rRNA gene sequence with the most common ARISA fragments to identify these microbial community members.


Cre recombinase  is a tyrosine recombinase  enzyme  derived from the  P1 bacteriophage . The enzyme uses a  topoisomerase I -like mechanism to carry out  site specific recombination  events. The enzyme (38 kDa) is a member of the  integrase  family of site specific recombinase and it is known to catalyse the  site specific recombination  event between two DNA recognition sites ( LoxP sites ). This 34 base pair (bp) loxP recognition site consists of two 13 bp  palindromic sequences  which flank an 8bp spacer region. The products of  Cre-mediated recombination  at loxP sites are dependent upon the location and relative orientation of the loxP sites. Two separate DNA species both containing loxP sites can undergo fusion as the result of Cre mediated recombination. DNA sequences found between two loxP sites are said to be " floxed ". In this case the products of Cre mediated recombination depends upon the orientation of the loxP sites. DNA found between two loxP sites oriented in the same direction will be excised as a circular loop of DNA whilst intervening DNA between two loxP sites that are opposingly orientated will be inverted. [ 1 ]  The enzyme requires no additional  cofactors  (such as  ATP ) or accessory proteins for its function. [ 2 ]


The enzyme plays important roles in the life cycle of the P1 bacteriophage, such as cyclization of the linear genome and resolution of dimeric  chromosomes  that form after  DNA replication . [ 3 ]


Cre recombinase is a widely used tool in the field of  molecular biology . The enzyme's unique and specific recombination system is exploited to manipulate genes and chromosomes in a huge range of research, such as  gene knock out  or  knock in  studies. The enzyme's ability to operate efficiently in a wide range of cellular environments (including mammals, plants, bacteria, and yeast) enables the Cre-Lox recombination system to be used in a vast number of organisms, making it a particularly useful tool in scientific research. [ 4 ]


Studies carried out in 1981 by Sternberg and Hamilton demonstrated that the bacteriophage ' P1 ' had a unique site specific recombination system.  EcoRI  fragments of the P1 bacteriophage  genome  were generated and  cloned  into  lambda vectors . A 6.5kb EcoRI fragment (Fragment 7) was found to permit efficient recombination events. [ 5 ]  The mechanism of these recombination events was known to be unique as they occurred in the absence of bacterial  RecA  and  RecBCD  proteins. The components of this recombination system were elucidated using deletion  mutagenesis  studies. These studies showed that a P1 gene product and a recombination site were both required for efficient recombination events to occur. The P1 gene product was named  Cre  ( c yclization  re combination) and the recombination site was named loxP ( lo cus of crossing ( x ) over,  P 1). [ 5 ]  The Cre protein was purified in 1983 and was found to be a 35,000 Da protein. [ 2 ]  No high energy cofactors such as  ATP  or accessory proteins are required for the recombinase activity of the purified protein. [ 2 ]  Early studies also demonstrated that Cre binds to non specific DNA sequences whilst having a 20 fold higher affinity for loxP sequences and results of early  DNA footprinting  studies also suggested that Cre molecules bind loxP sites as  dimers . [ 2 ]