The specific sequence of bases in DNA gives the material an element of programmability by determining its binding behavior. Carefully designing the sequences of the staple strands enables scientists to precisely direct the scaffold strand's folding into a predetermined shape with high precision. [ 9 ]


On a chemical level, the hydrogen bonds that exist between the complementary base pairs provide strength and stability to the folded DNA origami structures. Additionally, DNA is a relatively stable molecule, offering resilience in physiological conditions. [ 9 ]


One of the advantages of using a DNA Origami nanostructure over an otherwise classified DNA nanostructure is the ease of defining finite structures. [ 10 ]  In the design of some other DNA nanostructures, it can be impractical to design the extremely large number of individualized strands if the entire structure is composed of smaller strands. One method of bypassing the need for a huge number of different strands is to use repeating units, which comes with the disadvantage of a distribution of sizes and sometimes shapes. DNA Origami, however, forms discrete structures. [ 10 ]


Applications for DNA Origami are primarily focused around the ability to exert fine control on systems, especially by constraining positions of molecules, typically by attachment to the DNA Origami nanostructures. Current applications are primarily focused around sensing and drug delivery, but many additional applications have been investigated.


Fabrication of DNA origami objects requires a preliminary intuition of 3-dimensional DNA structural design. This can be difficult to grasp due to the complexity of exclusively using  adenine - thymine  pairings and  guanine - cytosine  pairings to both fold and unravel double helical DNA molecules such that the output strands produce uniquely desired shapes.


The design software and the choice of base-pair sequences become crucial for creating intricate 2D or even 3D shapes as the key to DNA origami lies in the precise  base-pairing  between the technique's two building blocks: staple strands and the scaffold. This ensures specific binding and accurate folding. A scaffold strand is a long, single-stranded DNA molecule, often sourced from a virus. Staple strands are shorter DNA strands designed to bind to specific sequences on the scaffold strand, dictating its folding. [ 9 ]


To produce a desired shape, images are drawn with a  raster fill  of a single long DNA  molecule . This design is then fed into a computer program that calculates the placement of individual staple strands. Each staple binds to a specific region of the DNA template, and thus due to  Watson–Crick base pairing , the necessary sequences of all staple strands are known and displayed. The DNA is mixed, then heated and cooled. As the DNA cools, the various staples pull the long strand into the desired shape. Designs are directly observable via several methods, including  electron microscopy ,  atomic force microscopy , or  fluorescence microscopy  when DNA is coupled to fluorescent materials. [ 6 ]


Bottom-up  self-assembly  methods are considered promising alternatives that offer cheap, parallel synthesis of nanostructures under relatively mild conditions.


Since the creation of this method, software was developed to assist the process using  CAD  software. This allows researchers to use a computer to determine the way to create the correct staples needed to form a certain shape. One such software called caDNAno is an open source software for creating such structures from DNA. The use of software has not only increased the ease of the process but has also drastically reduced the errors made by manual calculations. [ 11 ] [ 5 ]


After meticulously planning the sequence of the staple strands with software to ensure they bind the scaffold strand at the intended points, the designed staple strand sequences are synthesized in a lab using techniques like automated DNA synthesis. Finally, the scaffold strand and staple strands are mixed in a buffer solution and subjected to a specific temperature cycle. This cycle allows the staple strands to find their complementary sequences on the scaffold strand and bind through  hydrogen bonding , causing the scaffold to fold into the desired shape. [ 9 ]


As in the broader field of DNA nanotechnology, DNA Origami may be made dynamic in nature through the use of a variety of methods. The three primary methods of creating a dynamic DNA Origami machine are toehold mediated strand displacement, enzymatic reactions, and base stacking. [ 12 ]  While these methods are most commonly used, additional methods for creating dynamic DNA Origami machines exist, such as designing a directional component and using brownian motion to drive rotational movement of structures  [ 13 ]  or leveraging less commonly used DNA self-assembly phenomena like G-quadruplexes or i-motifs which can be pH sensitive. [ 14 ]


Modifications can be otherwise used to affect structural properties, to impart unique chemistry to the nanostructures, or to add stimuli responses to the nanostructures. Modifications to structures can be made through conjugation of molecules such as proteins, or through chemical modification of the DNA bases themselves. pH dependent responses, light dependent responses, and more have been shown through modified systems.


One example application of creating dynamic structures is the ability to have a stimuli response resulting in drug release, which is presented by several groups. [ 15 ] [ 16 ]  Other, less common applications comes in sensing moving mechanisms in vivo such as the unwinding of helicase. [ 17 ]


DNA Origami, being made of a natural biological polymer, is well suited to the biological environment when salt concentrations allow, [ 1 ]  and offers fine control over the positioning of molecules and structures in the system. This allows DNA Origami to be applicable to a number of scenarios in biomedical engineering. Current biomedical applications include drug release with 0 order mechanisms, [ 2 ]  vaccines, [ 3 ]  cell signaling, [ 4 ]  and sensing applications. [ 5 ]


DNA is folded into an  octahedron  and coated with a single bilayer of  phospholipid , mimicking the envelope of a  virus  particle.  The DNA nanoparticles, each at about the size of a virion, are able to remain in circulation for hours after injected into mice.  It also elicits much lower immune response than the uncoated particles.  It presents a potential use in drug delivery, reported by researchers in Wyss Institute at Harvard University. [ 18 ] [ 19 ]


Researchers at the  Harvard University   Wyss Institute  reported the self-assembling and self-destructing drug delivery vessels using the DNA origami in the lab tests.  The DNA nanorobot they created is an open DNA tube with a hinge on one side which can be clasped shut.  The drug filled DNA tube is held shut by a DNA  aptamer , configured to identify and seek certain diseased related protein.  Once the origami nanobots get to the infected cells, the aptamers break apart and release the drug.  The first disease model the researchers used was  leukemia  and  lymphoma . [ 20 ]


Researchers in the  National Center for Nanoscience and Technology  in  Beijing  and  Arizona State University  reported a DNA origami delivery vehicle for  Doxorubicin , a well-known anti-cancer drug.   The drug was non-covalently attached to DNA origami nanostructures through intercalation and a high drug load was achieved.  The DNA-Doxorubicin complex was taken up by human breast adenocarcinoma cancer cells ( MCF-7 ) via cellular internalization with much higher efficiency than doxorubicin in free form.  The enhancement of cell killing activity was observed not only in regular  MCF-7 , more importantly, also in doxorubicin-resistant cells.  The scientists theorized that the doxorubicin-loaded DNA origami inhibits  lysosomal  acidification, resulting in cellular redistribution of the drug to action sites, thus increasing the  cytotoxicity  against the tumor cells. [ 21 ] [ 22 ]  Further testing on in vivo on mice suggests that over a 12 day period, Doxorubicin was more effective at reducing tumor sizes in mice when it was contained in DNA Origami Nanostructures or DONs. [ 23 ]


Researchers from the  Massachusetts Institute of Technology  are developing a method to attach various viral antigens to Virus-shaped DNA particles to mimic the virus to be used to develop new vaccines. [ 24 ]  This was started in 2016 when Bathe's lab created an algorithm known as DAEDALUS (DNA Origami Sequence Design Algorithm for User-defined Structures) to generate precision-controlled three-dimensional shapes of DNA. [ 25 ]  Using the tool they designed virus-shaped scaffolding that can modularly attach different antigens to the surface of the DNA scaffold. Currently, MIT is working to develop optimal geometries for B cells to recognize HIV antigens. Further research has attempted to replace HIV antigens with SARS-CoV-2 and are testing whether vaccines show proper immune response from isolated  B cells  and in mice. [ 26 ]


Similarly, researchers from the  Technical University of Munich  have developed a method to have  T-cells  target tumor cells by using antigen coated DNA origami. [ 27 ]  The researchers developed a method to create chassis known as programmable T-cell Engagers or (PTEs) which are DNA Origami structures that can be configured to bind to user-defined target cells and T-cells based on which antigens are coated on the surfaces of the nanostructure. The in vitro results show that after 24 hours of exposure 90% of the tumor cells were destroyed. Meanwhile in vivo testing showed that their PTEs were capable of binding to the target proteins for several hours which validates the mechanism they designed. [ 28 ]


Many potential applications have been suggested in literature, including enzyme immobilization, drug delivery systems, and nanotechnological self-assembly of materials. Though DNA is not the natural choice for building active structures for nanorobotic applications, due to its lack of structural and catalytic versatility, several papers have examined the possibility of molecular walkers on origami and switches for algorithmic computing. [ 8 ] [ 29 ]   The following paragraphs list some of the reported applications conducted in the laboratories with clinical potential.


In a study conducted by a group of scientists from  iNANO center  and  CDNA Center  at  Aarhus university , researchers were able to construct a small multi-switchable 3D DNA Box Origami. The proposed nanoparticle was characterized by  AFM ,  TEM  and  FRET . The constructed box was shown to have a unique reclosing mechanism, which enabled it to repeatedly open and close in response to a unique set of DNA or RNA keys. The authors proposed that this "DNA device can potentially be used for a broad range of applications such as controlling the function of single molecules, controlled drug delivery, and molecular computing." [ 30 ]


Nanorobots made of DNA origami demonstrated computing capacities and completed pre-programmed task inside the living organism was reported by a team of bioengineers at Wyss Institute at Harvard University and Institute of Nanotechnology and Advanced Materials at  Bar-Ilan University .  As a proof of concept, the team injected various kinds of nanobots (the curled DNA encasing molecules with  fluorescent  markers) into live cockroaches.  By tracking the markers inside the cockroaches, the team found the accuracy of delivery of the molecules (released by the uncurled DNA) in target cells, the interactions among the nanobots and the control are equivalent to a computer system.  The complexity of the logic operations, the decisions and actions, increases with the increased number of nanobots.  The team estimated that the computing power in the cockroach can be scaled up to that of an 8-bit computer. [ 31 ] [ 32 ]


A research group at the  Indian Institute of Science  used nanostructures to develop a platform to elucidate the coaxial stacking between DNA bases. This approach utilized  DNA-PAINT  based super-resolution microscopy for visualizing these DNA nanostructures and performed DNA binding kinetics analysis to elucidate the fundamental force of base-stacking that helps stabilize the DNA double helical structure. They went on to assemble multimeric DNA origami nanostructures termed as a 'three-point star' into a  tetrahedral  3D origami structure. The assembly relied chiefly on base-stacking interactions between each subunit. The group further showed that the knowledge of such interactions can be used to predict and thus tune the relative stabilities of these multimeric DNA nanostructures. [ 33 ]


The idea of using  protein design  to accomplish the same goals as DNA origami has surfaced as well. Researchers at the National Institute of Chemistry in Slovenia are working on using  rational design  of  protein folding  to create structures much like those seen with DNA origami. The main focus of current research in protein folding design is in the drug delivery field, using antibodies attached to proteins as a way to create a targeted vehicle. [ 34 ] [ 35 ]


A  DNA sequencer  is a  scientific instrument  used to automate the  DNA sequencing  process.  Given a sample of  DNA , a DNA sequencer is used to determine the order of the four bases: G ( guanine ), C ( cytosine ), A ( adenine ) and T ( thymine ). This is then reported as a text  string , called a read. Some DNA sequencers can be also considered  optical instruments  as they analyze light signals originating from  fluorochromes  attached to  nucleotides .


The first automated DNA sequencer, invented by  Lloyd M. Smith , was introduced by  Applied Biosystems  in 1987. [ 1 ]  It used the  Sanger sequencing  method, a technology which formed the basis of the "first generation" of DNA sequencers [ 2 ] [ 3 ]  and enabled the completion of the  human genome project  in 2001. [ 4 ]  This first generation of DNA sequencers are essentially automated  electrophoresis  systems that detect the migration of labelled DNA fragments. Therefore, these sequencers can also be used in the  genotyping  of genetic markers where only the length of a DNA fragment(s) needs to be determined (e.g.  microsatellites ,  AFLPs ).


The  Human Genome Project  spurred the development of cheaper, high throughput and more accurate platforms known as  Next Generation Sequencers  (NGS) to sequence the  human genome . These include the 454,  SOLiD  and  Illumina  DNA sequencing platforms. Next generation sequencing machines have increased the rate of DNA sequencing substantially, as compared with the previous Sanger methods. DNA samples can be prepared automatically in as little as 90 mins, [ 5 ]  while a human genome can be sequenced at 15 times coverage in a matter of days. [ 6 ]


More recent, third-generation DNA sequencers such as PacBio  SMRT  and  Oxford Nanopore  offer the possibility of sequencing long molecules, compared to short-read technologies such as Illumina SBS or MGI Tech's DNBSEQ.


Because of limitations in DNA sequencer technology, the reads of many of these technologies are short, compared to the length of a  genome  therefore the reads must be  assembled  into longer  contigs . [ 7 ]  The data may also contain errors, caused by limitations in the DNA sequencing technique or by errors during  PCR amplification . DNA sequencer manufacturers use a number of different methods to detect which DNA bases are present. The specific protocols applied in different sequencing platforms have an impact in the final data that is generated. Therefore, comparing data quality and cost across different technologies can be a daunting task. Each manufacturer provides their own ways to inform sequencing errors and scores. However, errors and scores between different platforms cannot always be compared directly. Since these systems rely on different DNA sequencing approaches, choosing the best DNA sequencer and method will typically depend on the experiment objectives and available budget. [ 2 ]


The first  DNA sequencing  methods were developed by Gilbert (1973) [ 8 ]  and Sanger (1975). [ 9 ]  Gilbert introduced a sequencing method based on chemical modification of DNA followed by cleavage at specific bases whereas Sanger's technique is based on  dideoxynucleotide  chain termination. The Sanger method became popular due to its increased efficiency and low radioactivity. The first automated DNA sequencer was the AB370A, introduced in 1986 by  Applied Biosystems . The AB370A was able to sequence 96 samples simultaneously, 500 kilobases per day, and reaching read lengths up to 600 bases. This was the beginning of the "first generation" of DNA sequencers, [ 2 ] [ 3 ]  which implemented Sanger sequencing, fluorescent dideoxy nucleotides and polyacrylamide gel sandwiched between glass plates - slab gels. The next major advance was the release in 1995 of the AB310 which utilized a linear polymer in a capillary in place of the slab gel for DNA strand separation by electrophoresis. These techniques formed the base for the completion of the human genome project in 2001. [ 4 ]  The human genome project spurred the development of cheaper, high throughput and more accurate platforms known as Next Generation Sequencers (NGS). In 2005,  454 Life Sciences  released the 454 sequencer, followed by Solexa Genome Analyzer and SOLiD (Supported Oligo Ligation Detection) by Agencourt in 2006. Applied Biosystems acquired Agencourt in 2006, and in 2007,  Roche  bought 454 Life Sciences, while Illumina purchased Solexa. Ion Torrent entered the market in 2010 and was acquired by Life Technologies (now  Thermo Fisher Scientific ). And  BGI  started manufacturing sequencers in China after acquiring  Complete Genomics  under their  MGI  arm. These are still the most common NGS systems due to their competitive cost, accuracy, and performance.


More recently, a third generation of DNA sequencers was introduced. The sequencing methods applied by these sequencers do not require DNA amplification (polymerase chain reaction – PCR), which speeds up the sample preparation before sequencing and reduces errors. In addition, sequencing data is collected from the reactions caused by the addition of nucleotides in the complementary strand in real time. Two companies introduced different approaches in their third-generation sequencers.  Pacific Biosciences  sequencers utilize a method called Single-molecule real-time (SMRT), where sequencing data is produced by light (captured by a camera) emitted when a nucleotide is added to the complementary strand by enzymes containing fluorescent dyes.  Oxford Nanopore Technologies  is another company developing third-generation sequencers using electronic systems based on nanopore sensing technologies.


DNA sequencers have been developed, manufactured, and sold by the following companies, among others.


The 454 DNA sequencer was the first next-generation sequencer to become commercially successful. [ 10 ]  It was developed by  454 Life Sciences  and purchased by Roche in 2007. 454 utilizes the detection of pyrophosphate released by the DNA polymerase reaction when adding a nucleotide to the template strain.


Roche currently manufactures two systems based on their pyrosequencing technology: the GS FLX+ and the GS Junior System. [ 11 ]  The GS FLX+ System promises read lengths of approximately 1000 base pairs while the GS Junior System promises 400 base pair reads. [ 12 ] [ 13 ]  A predecessor to GS FLX+, the 454 GS FLX Titanium system was released in 2008, achieving an output of 0.7G of data per run, with 99.9% accuracy after quality filter, and a read length of up to 700bp. In 2009, Roche launched the GS Junior, a bench top version of the 454 sequencer with read length up to 400bp, and simplified library preparation and data processing.


One of the advantages of 454 systems is their running speed. Manpower can be reduced with automation of library preparation and semi-automation of emulsion PCR. A disadvantage of the 454 system is that it is prone to errors when estimating the number of bases in a long string of identical nucleotides. This is referred to as a homopolymer error and occurs when there are 6 or more identical bases in row. [ 14 ]  Another disadvantage is that the price of reagents is relatively more expensive compared with other next-generation sequencers.


In 2013 Roche announced that they would be shutting down development of 454 technology and phasing out 454 machines completely in 2016 when its technology became noncompetitive. [ 15 ] [ 16 ]


Roche produces a number of software tools which are optimised for the analysis of 454 sequencing data. [ 17 ]  Such as,


Illumina  produces a number of next-generation sequencing machines using technology acquired from  Manteia Predictive Medicine  and developed by Solexa. [ 19 ]  Illumina makes a number of next generation sequencing machines using this technology including the HiSeq, Genome Analyzer IIx, MiSeq and the HiScanSQ, which can also process  microarrays . [ 20 ]


The technology leading to these DNA sequencers was first released by Solexa in 2006 as the Genome Analyzer. [ 10 ]  Illumina purchased Solexa in 2007. The Genome Analyzer uses a sequencing by synthesis method. The first model produced 1G per run. During the year 2009 the output was increased from 20G per run in August to 50G per run in December. In 2010 Illumina released the HiSeq 2000 with an output of 200 and then 600G per run which would take 8 days. At its release the HiSeq 2000 provided one of the cheapest sequencing platforms at $0.02 per million bases as costed by the  Beijing Genomics Institute .


In 2011 Illumina released a benchtop sequencer called the MiSeq. At its release the MiSeq could generate 1.5G per run with paired end 150bp reads. A sequencing run can be performed in 10 hours when using automated DNA sample preparation. [ 10 ]


The Illumina HiSeq uses two software tools to calculate the number and position of DNA clusters to assess the sequencing quality: the HiSeq control system and the real-time analyzer. These methods help to assess if nearby clusters are interfering with each other. [ 10 ]


Life Technologies  (now  Thermo Fisher Scientific ) produces DNA sequencers under the  Applied Biosystems  and  Ion Torrent  brands. Applied Biosystems makes the SOLiD next-generation sequencing platform, [ 21 ]  and Sanger-based DNA sequencers such as the 3500 Genetic Analyzer. [ 22 ]  Under the Ion Torrent brand, Applied Biosystems produces four next-generation sequencers: the Ion PGM System, Ion Proton System, Ion S5 and Ion S5xl systems. [ 23 ]  The company is also believed to be developing their new capillary DNA sequencer called SeqStudio that will be released early 2018. [ 24 ]


SOLiD systems was acquired by  Applied Biosystems  in 2006. SOLiD applies sequencing by ligation and  dual base encoding . The first SOLiD system was launched in 2007, generating reading lengths of 35bp and 3G data per run. After five upgrades, the 5500xl sequencing system was released in 2010, considerably increasing read length to 85bp, improving accuracy up to 99.99% and producing 30G per 7-day run. [ 10 ]


The limited read length of the SOLiD has remained a significant shortcoming [ 25 ]  and has to some extent limited its use to experiments where read length is less vital such as resequencing and transcriptome analysis and more recently ChIP-Seq and methylation experiments. [ 10 ]  The DNA sample preparation time for SOLiD systems has become much quicker with the automation of sequencing library preparations such as the Tecan system. [ 10 ]


The colour space data produced by the SOLiD platform can be decoded into DNA bases for further analysis, however software that considers the original colour space information can give more accurate results. Life Technologies has released BioScope, [ 26 ]  a data analysis package for resequencing, ChiP-Seq and transcriptome analysis. It uses the MaxMapper algorithm to map the colour space reads.


Beckman Coulter  (now  Danaher ) has previously manufactured chain termination and capillary electrophoresis-based DNA sequencers under the model name CEQ, including the CEQ 8000. [ 27 ]  The company now produces the GeXP Genetic Analysis System, which uses  dye terminator sequencing . [ 28 ]  This method uses a  thermocycler  in much the same way as  PCR  to denature, anneal, and extend DNA fragments, amplifying the sequenced fragments. [ 29 ] [ 30 ]


Pacific Biosciences  produces the PacBio RS and Sequel sequencing systems using a  single molecule real time sequencing , or SMRT, method. [ 31 ]  This system can produce read lengths of multiple thousands of base pairs.  Higher raw read errors are corrected using either circular consensus - where the same strand is read over and over again - or using optimized  assembly  strategies. [ 32 ]  Scientists have reported 99.9999% accuracy with these strategies. [ 33 ]  The Sequel system was launched in 2015 with an increased capacity and a lower price. [ 34 ] [ 35 ]


Oxford Nanopore Technologies ' MinION sequencer is based on evolving  nanopore sequencing  technology to nucleic acid analyses. [ 37 ]  The device is four inches long and gets power from a  USB port . MinION decodes DNA directly as the molecule is drawn at the rate of 450 bases/second through a  nanopore  suspended in a membrane. [ 38 ]  Changes in electric current indicate which base is present. Initially, the device was 60 to 85 percent accurate, compared with 99.9 percent in conventional machines. [ 39 ]  Even inaccurate results may prove useful because it produces long read lengths. [ 40 ]  In early 2021, researchers from  University of British Columbia  has used special molecular tags and able to reduce the five-to-15 per cent error rate of the device to less than 0.005 per cent even when sequencing many long stretches of DNA at a time. [ 41 ]  There are two more product iterations based on MinION; the first one is the GridION which is a slightly larger sequencer that processes up to five MinION flow cells at once. And, the second one is the PromethION which uses as many as 100,000 pores in parallel, more suitable for high volume sequencing. [ 42 ]


MGI  produces high-throughput sequencers for scientific research and clinical applications such as DNBSEQ-G50, DNBSEQ-G400, and DNBSEQ-T7, under a proprietary DNBSEQ technology. [ 43 ]  It is based upon  DNA nanoball sequencing  and combinatorial probe anchor synthesis technologies, in which DNA nanoballs (DNBs) are loaded onto a patterned array chip via the fluidic system, and later a sequencing primer is added to the adaptor region of DNBs for  hybridization . DNBSEQ-T7 can generate short reads at a very large scale—up to 60 human genomes per day. [ 44 ]  DNBSEQ-T7 was used to generate 150 bp paired-end reads, sequencing 30X, to sequence the genome of SARS-CoV-2 or COVID-19 to identify the genetic variants predisposition in severe COVID-19 illness. [ 45 ]  Using a novel technique the researchers from  China National GeneBank  sequenced  PCR -free libraries on MGI's PCR-free DNBSEQ arrays to obtain for the first time a true PCR-free  whole genome sequencing . [ 46 ]  MGISEQ-2000 was used in single-cell RNA sequencing to study the underlying pathogenesis and recovery in COVID-19 patients, as published in  Nature Medicine . [ 47 ]


Current offerings in DNA sequencing technology  show a dominant player:  Illumina  (December 2019), followed by  PacBio ,  MGI  and  Oxford Nanopore .


A  dot blot  (or  slot blot ) is a technique in  molecular biology  used to detect proteins. It represents a simplification of the  western blot  method, with the exception that the proteins to be detected are not first separated by  electrophoresis . Instead, the sample is applied directly on a membrane in a single spot, and the blotting procedure is performed.


The technique offers significant savings in time, as  chromatography  or  gel electrophoresis , and the complex blotting procedures for the gel are not required. However, it offers no information on the size of the target protein. [ 1 ]


Performing a dot blot is similar in idea to performing a western blot, with the advantage of faster speed and lower cost. [ 2 ]  Dot blots are also performed to screen the binding capabilities of an antibody. [ 3 ]


A general dot blot protocol involves spotting 1–2 microliters of a samples onto a  nitrocellulose  or  PVDF  membrane and letting it air dry. Samples can be in the form of tissue culture supernatants, blood serum, cell extracts, or other preparations. [ 4 ]  After the protein samples are spotted onto the membrane, the membrane is placed in a plastic container and sequentially incubated in blocking buffer, antibody solutions, or rinsing buffer on shaker. After antibody binding, the membrane is incubated with a  chemiluminescent  substrate and imaged.


Vacuum-assisted dot blot apparatus has been used to facilitate the rinsing and incubating process by using vacuum to extract the solution from underneath the membrane, which is assembled in between several layers of plates to ensure good seal between sample wells, hold waste solution, and deliver suction force. For chemiluminescence signal detection, apparatus need to be disassembled and the membrane need to be taken out and wrapped in a transparent plastic film.


An  electropherogram  (also called  electrophoretogram ,  sequencing chromatogram ,  EPG , and  e-gram ) is a record or chart produced when  electrophoresis  is used in an analytical technique, primarily in the fields of  forensic biology ,  molecular biology , and  biochemistry . [ 1 ]  The method plots data points that represent a specific time and  fluorescence  intensity at various wavelengths of light to represent a DNA profile. [ 2 ] [ page needed ]


In the field of genetics, an electropherogram is a plot of DNA fragment sizes, typically used for  genotyping  such as  DNA sequencing . [ 3 ]  The data is plotted with time, shown via base pairs (bps), on the x-axis and fluorescence intensity on the y-axis. Such plots are often achieved using an instrument such as an automated  DNA sequencer  paired with capillary electrophoresis (CE). Such electropherograms may be used to determine DNA sequence genotypes, or genotypes that are based on the length of specific DNA fragments or number of short tandem repeats (STR) at a specific locus by comparing the sample to internal size standards and allelic ladder data using the same size standard. [ 2 ] [ page needed ]  These genotypes can be used for:


An  electrophoretic color marker  is a chemical used to monitor the progress of  agarose gel electrophoresis  and  polyacrylamide gel electrophoresis  (PAGE) since DNA, RNA, and most proteins are colourless. [ 1 ]  The color markers are made up of a mixture of  dyes  that migrate through the gel matrix alongside the sample of interest. They are typically designed to have different mobilities from the sample components and to generate colored bands that can be used to assess the migration and separation of sample components. [ 2 ]


Color markers are often used as molecular weight standards, loading dyes, tracking dyes, or staining solutions. Molecular weight ladders are used to estimate the size of DNA and protein fragments by comparing their migration distance to that of the colored bands. [ 2 ]  DNA and protein standards are available commercially in a wide range of sizes, and are often provided with pre-stained or color-coded bands for easy identification. Loading dyes are usually added to the sample buffer before loading the sample onto the gel, and they migrate through the gel along with the sample to help track its progress during electrophoresis. [ 3 ]  Tracking dyes are added to the electrophoresis buffer rather to provide a visual marker of the buffer front. Staining solutions are applied after electrophoresis to visualize the sample bands, and are available in a range of colors.


Different types of electrophoretic color markers are available commercially, with varying numbers and types of dyes or pigments used in the mixture. Some markers generate a series of colored bands with known mobilities, while others produce a single band of a specific color that can be used as a reference point. They are widely used in research,  clinical diagnostics , and  forensic science . [ 4 ]


Loading buffers often contain anionic dyes that are visible under the visible light spectrum, and are added to the gel before the nucleic acid. Tracking dyes should not be reactive so as not to alter the sample, and move down the gel with the DNA or RNA sample. [ 5 ]  Commonly used color markers include  Bromophenol blue ,  Cresol Red ,  Orange G  and  Xylene cyanol . Xylene and bromophenol blue are the most commonly used dyes. [ citation needed ]  Generally speaking, Orange G migrates faster than bromophenol blue, which migrates faster than xylene cyanol, but the apparent "sizes" of these dyes (compared to DNA molecules) varies with the concentration of agarose and the  buffer  system used. For instance, in a 1%  agarose  gel made in TAE buffer ( Tris - acetate - EDTA ), xylene cyanol migrates at the speed of a 3000 base pair (bp) molecule of DNA and bromophenol blue migrates at 400 bp. However, in a 1% gel made in TBE buffer (Tris- borate -EDTA), they migrate at 2000 bp and 250 bp respectively. [ 3 ]


Agarose gel electrophoresis is a technique widely used to estimate the size of nucleic acid fragments and identify them based on their differential mobility in the gel. Nucleic acids are commonly stained and detected using either ethidium bromide or SYBR Green dyes. [ citation needed ]  The most common electrophoretic stain in agarose gel is ethidium bromide, however, SYBR green presents greater resolution and yield for single-stranded nucleic acid detection. [ citation needed ]  The dyes grant fluorescence to DNA and RNA under 300 nm UV light. This occurs due to their intercalating nature. In double helical nucleic acids, the dyes bind between two strands, and in single-stranded nucleic acids, the dyes bind short, duplex segments formed within a strand. [ citation needed ]


The most commonly used dye in agarose gel gel electrophoresis of DNA and RNA, dating as far back as the 1970s, is  ethidium bromide  (2,7-diamino-10-ethyl-9-phenylphenanthridiniumbromide). [ citation needed ]  Ethidium Bromide (EtBr) is an orange-colored fluorescent intercalating dye. The dye inserts itself between the double helical structure of nucleic acids, allowing for visualization of the molecules under UV light. [ 6 ]  EtBr has absorbance maxima at 300-360 nm and fluorescent emission maxima at 500-590 nm, with the detection limit of 0.5-5.0 ng/band. [ 6 ]  The dye, however, has reduced sensitivity in the detection of single-stranded nucleic acid samples EtBr should be handled with care, as it is a potent mutagen. [ 6 ]


A more sensitive alternative for nucleic acid staining in gel electrophoresis is SYBR™ Green I. [ 7 ]   The dye is 25 times more sensitive than EtBr in the staining of dsDNA, and is especially useful in staining assays containing single-stranded nucleic acids. [ citation needed ]  SYBR Green is, however, more expensive when compared to EtBr.


Coomassie Blue  is the most commonly used non-covalent stain in SDS polyacrylamide gel electrophoresis for protein quantification. The staining dye binds to the protein bands and creates a blue color that can be detected visually.  Coomassie Brilliant Blue R-250 (red), is typically used for electrophoresis, while Coomassie Brilliant Blue G-250 (green), for Bradford Assay. [ 8 ]   The limitation of this dye is that it is non-specific, and will bind to almost any protein in solution, and is less sensitive. [ citation needed ]   Another common method of visualization of proteins in the gel is silver staining, where soluble silver ions permanently mark proteins and are reduced by formaldehyde to form a brown precipitate. [ citation needed ]   Silver staining is a more sensitive staining method when compared to Coomassie Blue, however, results are more vulnerable to contamination.


Color markers are sometimes added to loading dyes for gel electrophoresis in the separation of DNA fragments. Loading dyes keep DNA samples below the surface of the agarose gel, and the color markers within help keep track of the migration front of the DNA as it moves along the gel. [ 9 ]


For PAGE, some commercially available molecular weight markers (also called "ladders" because they look like the rungs of a ladder after separation) contain pre-stained proteins of different colours, so it is possible to determine more accurately where the proteins of interest in the samples might be.


An  electrophoretic mobility shift assay  (EMSA) or  mobility shift electrophoresis , also referred as a  gel shift assay ,  gel mobility shift assay ,  band shift assay , or  gel retardation assay , is a common  affinity electrophoresis  technique used to study  protein–DNA  or  protein – RNA  interactions.  This procedure can determine if a protein or mixture of proteins is capable of binding to a given DNA or RNA sequence, and can sometimes indicate if more than one protein molecule is involved in the binding complex.  Gel shift assays are often performed  in vitro  concurrently with  DNase footprinting ,  primer extension , and promoter-probe experiments when studying  transcription  initiation, DNA gang replication, DNA repair or RNA processing and maturation, as well as pre-mRNA splicing. [ 1 ]  Although precursors can be found in earlier literature, most current assays are based on methods described by Garner and Revzin  [ 2 ]  and Fried and  Crothers . [ 3 ]


A mobility shift assay is  electrophoretic separation  of a protein–DNA or protein–RNA mixture on a  polyacrylamide  or  agarose gel  for a short period (about 1.5-2 hr for a 15- to 20-cm gel). [ 4 ]  The speed at which different molecules (and combinations thereof) move through the gel is determined by their size and charge, and to a lesser extent, their shape (see  gel electrophoresis ). The control lane (DNA probe without protein present) will contain a single band corresponding to the unbound DNA or RNA fragment. However, assuming that the protein is capable of binding to the fragment, the lane with a protein that binds present will contain another band that represents the larger, less mobile complex of nucleic acid probe bound to protein which is 'shifted' up on the gel (since it has moved more slowly).


Under the correct experimental conditions, the interaction between the DNA (or RNA) and protein is stabilized and the ratio of bound to unbound nucleic acid on the gel reflects the fraction of free and bound probe molecules as the binding reaction enters the gel. This stability is in part due to a "caging effect", in that the protein, surrounded by the gel matrix, is unable to diffuse away from the probe before they recombine. [ 5 ]  If the starting concentrations of protein and probe are known, and if the stoichiometry of the complex is known, the apparent affinity of the protein for the nucleic acid sequence may be determined. [ 6 ]  Unless the complex is very long lived under gel conditions, or dissociation during electrophoresis is taken into account, the number derived is an apparent Kd.  If the protein concentration is not known but the complex stoichiometry is, the protein concentration can be determined by increasing the concentration of DNA probe until further increments do not increase the fraction of protein bound. By comparison with a set of standard dilutions of free probe run on the same gel, the number of moles of protein can be calculated. [ 4 ]


An antibody that recognizes the protein can be added to this mixture to create an even larger complex with a greater shift. This method is referred to as a  supershift assay , and is used to unambiguously identify a protein present in the protein – nucleic acid complex.


Often, an extra lane is run with a competitor  oligonucleotide  to determine the most favorable binding sequence for the binding protein. The use of different oligonucleotides of defined sequence allows the identification of the precise binding site by competition (not shown in diagram). Variants of the competition assay are useful for measuring the specificity of binding and for measurement of association and dissociation kinetics. Thus, EMSA might also be used as part of a SELEX experiment to select for oligonucleotides that do actually bind a given protein. [ citation needed ]


Once DNA-protein binding is determined  in vitro , a number of algorithms can narrow the search for identification of the transcription factor. Consensus sequence oligonucleotides for the transcription factor of interest will be able to compete for the binding, eliminating the shifted band, and must be confirmed by supershift. If the predicted consensus sequence fails to compete for binding, identification of the transcription factor may be aided by Multiplexed Competitor EMSA (MC-EMSA), whereby large sets of consensus sequences are multiplexed in each reaction, and where one set competes for binding, the individual consensus sequences from this set are run in a further reaction. [ 7 ]


For visualization purposes, the nucleic acid fragment is usually labelled with a  radioactive ,  fluorescent  or  biotin  label. Standard  ethidium bromide  staining is less sensitive than these methods and can lack the sensitivity to detect the nucleic acid if small amounts of nucleic acid or single-stranded nucleic acid(s) are used in these experiments. When using a biotin label,  streptavidin  conjugated to an enzyme such as horseradish peroxidase is used to detect the DNA fragment. [ 8 ] [ 9 ]  While isotopic DNA labeling has little or no effect on protein binding affinity, use of non-isotopic labels including flurophores or biotin can alter the affinity and/or stoichiometry of the protein interaction of interest. Competition between fluorophore- or biotin-labeled probe and unlabeled DNA of the same sequence can be used to determine whether the label alters binding affinity or stoichiometry.


Exonucleases  are  enzymes  that work by cleaving  nucleotides  one at a time from the end (exo) of a polynucleotide chain. A  hydrolyzing  reaction that breaks phosphodiester bonds at either the 3′ or the  5′ end  occurs.  Its close relative is the  endonuclease , which cleaves  phosphodiester bonds  in the middle (endo) of a polynucleotide chain.   Eukaryotes  and  prokaryotes  have three types of exonucleases involved in the normal turnover of  mRNA :  5′ to 3′ exonuclease (Xrn1) , which is a dependent  decapping protein ; 3′ to 5′ exonuclease, an independent protein; and poly(A)-specific 3′ to 5′ exonuclease. [ 1 ] [ 2 ]


In both  archaea  and  eukaryotes , one of the main routes of RNA degradation is performed by the multi-protein  exosome complex , which consists largely of 3′ to 5′  exoribonucleases .


RNA  polymerase  II is known to be in effect during transcriptional termination; it works with a 5' exonuclease (human gene Xrn2) to degrade the newly formed transcript downstream, leaving the polyadenylation site and simultaneously shooting the polymerase. This process involves the exonuclease's catching up to the pol II and terminating the transcription. [ 3 ]


Pol I  then synthesizes DNA nucleotides in place of the RNA primer it had just removed. DNA polymerase I also has 3' to 5' and 5' to 3' exonuclease activity, which is used in editing and proofreading DNA for errors. The 3' to 5' can only remove one mononucleotide at a time, and the 5' to 3' activity can remove mononucleotides or up to 10 nucleotides at a time.


In 1971, Lehman IR discovered exonuclease I in  E. coli . Since that time, there have been numerous discoveries including: exonuclease, II,  III , IV,  V , VI,  VII , and VIII. Each type of exonuclease has a specific type of function or requirement. [ 4 ]


Exonuclease I  breaks apart single-stranded DNA in a 3' → 5' direction, releasing deoxyribonucleoside 5'-monophosphates one after another. It does not cleave DNA strands without  terminal 3'-OH groups   because they are blocked by phosphoryl or acetyl groups.
 [ 5 ]


Exonuclease II  is associated with DNA polymerase I, which contains a 5' exonuclease that clips off the RNA primer contained immediately upstream from the site of DNA synthesis in a 5' → 3' manner.


Exonuclease III  has four catalytic activities:


Exonuclease IV  adds a water molecule, so it can break the bond of an oligonucleotide to nucleoside 5' monophosphate. This exonuclease requires  Mg 2+  in order to function and works at higher temperatures than exonuclease I. [ 7 ]


Exonuclease V  is a 3' to 5' hydrolyzing enzyme that catalyzes linear double-stranded DNA and single-stranded DNA, which requires  Ca2+ . [ 8 ]  This enzyme is extremely important in the process of  homologous recombination .


Exonuclease VIII  is 5' to 3' dimeric protein that does not require ATP or any gaps or nicks in the strand, but requires a  free 5' OH group  to carry out its function  [ citation needed ] .


In  Escherichia coli  the  dnaQ  gene encodes the ε subunit of DNA polymerase III . [ 9 ]  The ε subunit is one of three core proteins of the DNA polymerase complex. It acts as a 3’→5’ DNA directed proofreading exonuclease that removes incorrectly incorporated bases during replication. [ 10 ]   Similarly, in  Salmonella  typhimurium  bacteria, the 3’ to 5’ editing function employed during  DNA replication  is also encoded by a gene,  dnaQ , which specifies a 3’ to 5’ exonuclease subunit, one of the three separately encoded core proteins of the DNA polymerase III holoenzyme. [ 11 ]


In contrast to  E. coli  and  S. typhimurium , where the polymerase and editing functions are encoded by separate genes, in the  bacterial species  Buchnera aphidicola  the DNA polymerase encoded by the DNA III (polC) gene contains both DNA polymerase and 3’ to 5’ exonuclease domains. [ 11 ]   An evolutionary divergence (about 0.25 to 1.2 billion years ago), appears to have been associated with the separation of the DNA polymerase gene function from the 3’ to 5’ exonuclease editing gene function in the lineage that led to  E. coli  and  S. typhimurium . [ 11 ]


The 3' to 5' human type endonuclease is known to be essential for the proper processing of histone pre-mRNA, in which U7 snRNP directs the single cleavage process. Following the removal of the downstream cleavage product (DCP)  Xrn1  continues to further breakdown the product until it is completely degraded. [ 12 ]  This allows the nucleotides to be recycled.  Xrn1  is linked to a co-transcriptional cleavage (CoTC) activity that acts as a precursor to develop a free 5' unprotected end, so the exonuclease can remove and degrade the downstream cleavage product (DCP). This initiates transcriptional termination because one does not want DNA or RNA strands building up in their bodies. [ 13 ]


CCR4-Not  is a general transcription regulatory complex in  budding yeast  that is found to be associated with  mRNA  metabolism, transcription initiation, and mRNA degradation. CCR4 has been found to contain  RNA  and single-stranded  DNA  3' to 5' exonuclease activities. [ 14 ]   Another component associated with the  CCR4-Not  is CAF1 protein, which has been found to contain 3' to 5' or 5' to 3' exonuclease domains in the  mouse  and  Caenorhabditis elegans . [ 15 ]  This protein has not been found in yeast, which suggests that it is likely to have an abnormal exonuclease domain like the one seen in a metazoan. [ 16 ]  Yeast contains  Rat1  and Xrn1 exonuclease. The Rat1 works just like the human type (Xrn2) and  Xrn1  function in the cytoplasm is in the 5' to 3' direction to degrade RNAs (pre-5.8s and 25s rRNAs) in the absence of Rat1. [ 17 ] [ 18 ]


In beta  Coronaviruses , including  SARS-CoV-2 , a proof reading exonuclease, nsp14-ExoN, that is part of the viral genome is responsible for recombination that is implicated in novel strain emergence. [ 19 ]


Expression cloning  is a technique in  DNA cloning  that uses  expression vectors  to generate a library of clones, with each clone expressing one  protein . This  expression library  is then screened for the property of interest and clones of interest are recovered for further analysis. An example would be using an expression library to isolate genes that could confer  antibiotic resistance . [ 1 ]


Expression vectors are a specialized type of  cloning vector  in which the transcriptional and translational signals needed for the regulation of the gene of interest are included in the cloning vector.  The transcriptional and translational signals may be synthetically created to make the expression of the gene of interest easier to regulate.


Usually the ultimate aim of expression cloning is to produce large quantities of specific  proteins .  To this end, a  bacterial  expression  clone  may include a  ribosome   binding site  ( Shine-Dalgarno sequence ) to enhance translation of the gene of interest's mRNA, a  transcription  termination sequence, or, in  eukaryotes , specific sequences to promote the  post-translational modification  of the protein product.


An  expression vector , otherwise known as an  expression construct , is usually a  plasmid  or virus designed for  gene expression  in cells. The  vector  is used to introduce a specific  gene  into a target cell, and can commandeer the cell's mechanism for  protein synthesis  to produce the  protein   encoded  by the gene.  Expression vectors are the basic tools in  biotechnology  for the  production of proteins .


The  vector  is engineered to contain regulatory sequences that act as  enhancer  and  promoter  regions and lead to efficient transcription of the gene carried on the expression vector. [ 1 ]  The goal of a well-designed expression vector is the efficient production of protein, and this may be achieved by the production of significant amount of stable  messenger RNA , which can then be  translated  into protein. The expression of a protein may be tightly controlled, and the protein is only produced in significant quantity when necessary through the use of an  inducer .  In some systems, however, the protein may be expressed constitutively.  Escherichia coli  is commonly used as the host for  protein production , but other cell types may also be used. An example of the use of expression vector is the production of  insulin , which is used for medical treatments of  diabetes .


An expression vector has features that any  vector  may have, such as an  origin of replication , a  selectable marker , and a suitable site for the insertion of a gene like the  multiple cloning site . The cloned gene may be transferred from a specialized  cloning vector  to an expression vector, although it is possible to clone directly into an expression vector. The cloning process is normally performed in  Escherichia coli . Vectors used for protein production in organisms other than  E.coli  may have, in addition to a suitable origin of replication for its propagation in  E. coli , elements that allow them to be maintained in another organism, and these vectors are called  shuttle vectors .


An expression vector must have elements necessary for gene expression. These may include a  promoter , the correct translation initiation sequence such as a  ribosomal binding site  and  start codon , a  termination codon , and a  transcription termination sequence . [ 2 ]  There are differences in the machinery for protein synthesis between prokaryotes and eukaryotes, therefore the expression vectors must have the elements for expression that are appropriate for the chosen host.  For example, prokaryotes expression vectors would have a  Shine-Dalgarno sequence  at its translation initiation site for the binding of ribosomes, while eukaryotes expression vectors would contain the  Kozak consensus sequence .


The  promoter  initiates the  transcription  and is therefore the point of control for the expression of the cloned gene. The promoters used in expression vector are normally  inducible , meaning that protein synthesis is only initiated when required by the introduction of an  inducer  such as  IPTG . Gene expression however may also be constitutive (i.e. protein is constantly expressed) in some expression vectors. Low level of constitutive protein synthesis may occur even in expression vectors with tightly controlled promoters.


After the expression of the gene product, it may be necessary to purify the expressed protein; however, separating the protein of interest from the great majority of proteins of the host cell can be a protracted process. To make this purification process easier, a  purification tag  may be added to the cloned gene. This tag could be  histidine (His) tag , other marker peptides, or a  fusion partners  such as  glutathione S-transferase  or  maltose-binding protein . [ 3 ]  Some of these fusion partners may also help to increase the solubility of some expressed proteins. Other fusion proteins such as  green fluorescent protein  may act as a  reporter gene  for the identification of successful cloned genes, or they may be used to study protein expression in  cellular imaging . [ 4 ] [ 5 ]


The expression vector is  transformed  or  transfected  into the host cell for protein synthesis. Some expression vectors may have elements for transformation or the insertion of DNA into the host chromosome, for example the  vir  genes  for  plant transformation , and  integrase  sites for chromosomal integration .


Some vectors may include targeting sequence that may target the expressed protein to a specific location such as the  periplasmic space  of bacteria.


Different organisms may be used to express a gene's target protein, and the expression vector used will therefore have elements specific for use in the particular organism. The most commonly used organism for  protein production  is the bacterium  Escherichia coli . However, not all proteins can be successfully expressed in  E. coli , or be expressed with the correct form of post-translational modifications such as glycosylations, and other systems may therefore be used.


The expression host of choice for the expression of many proteins is  Escherichia coli  as the production of heterologous protein in  E. coli  is relatively simple and convenient, as well as being rapid and cheap. A large number of  E. coli  expression plasmids are also available for a wide variety of needs. Other bacteria used for protein production include  Bacillus subtilis .


Most heterologous proteins are expressed in the cytoplasm of  E. coli . However, not all proteins formed may be soluble in the cytoplasm, and incorrectly folded proteins formed in cytoplasm can form insoluble aggregates called  inclusion bodies . Such insoluble proteins will require refolding, which can be an involved process and may not necessarily produce high yield. [ 6 ]  Proteins which have  disulphide bonds  are often not able to fold correctly due to the reducing environment in the cytoplasm which prevents such bond formation, and a possible solution is to target the protein to the  periplasmic space  by the use of an N-terminal  signal sequence . Another possibility is to manipulate the redox environment of the cytoplasm. [ 7 ]  Other more sophisticated systems are also being developed; such systems may allow for the expression of proteins previously thought impossible in  E. coli , such as  glycosylated  proteins. [ 8 ] [ 9 ] [ 10 ]


The promoters used for these vector are usually based on the promoter of the  lac  operon  or the  T7  promoter, [ 11 ]  and they are normally regulated by the  lac   operator . These promoters may also be hybrids of different promoters, for example, the  Tac-Promoter  is a hybrid of  trp  and  lac  promoters. [ 12 ]  Note that most commonly used  lac  or  lac -derived promoters are based on the  lac UV5  mutant which is insensitive to  catabolite repression . This mutant allows for expression of protein under the control of the  lac  promoter when the  growth medium  contains glucose since glucose would inhibit gene expression if wild-type  lac  promoter is used. [ 13 ]  Presence of glucose nevertheless may still be used to reduce background expression through residual inhibition in some systems. [ 14 ]


Examples of  E. coli  expression vectors are the pGEX series of vectors where  glutathione S-transferase  is used as a fusion partner and gene expression is under the control of the tac promoter, [ 15 ] [ 16 ] [ 17 ]  and the pET series of vectors which uses a  T7  promoter. [ 18 ]


It is possible to simultaneously express two or more different proteins in  E. coli  using different plasmids. However, when 2 or more plasmids are used, each plasmid needs to use a different antibiotic selection as well as a different origin of replication, otherwise one of the plasmids may not be stably maintained. Many commonly used plasmids are based on the  ColE1  replicon and are therefore incompatible with each other; in order for a ColE1-based plasmid to coexist with another in the same cell, the other would need to be of a different replicon, e.g. a p15A replicon-based plasmid such as the pACYC series of plasmids. [ 19 ]  Another approach would be to use a single two-cistron vector or design the coding sequences in tandem as a bi- or poly-cistronic construct. [ 20 ] [ 21 ]


A yeast commonly used for protein production is  Pichia pastoris . [ 22 ]  Examples of yeast expression vector in  Pichia  are the pPIC series of vectors, and these vectors use the  AOX1  promoter which is inducible with  methanol . [ 23 ]  The plasmids may contain elements for insertion of foreign DNA into the yeast genome and signal sequence for the secretion of expressed protein. Proteins with disulphide bonds and glycosylation can be efficiently produced in yeast. Another yeast used for protein production is  Kluyveromyces lactis  and the gene is expressed, driven by a variant of the strong  lactase  LAC4 promoter. [ 24 ]


Saccharomyces cerevisiae  is particularly widely used for gene expression studies in yeast, for example in  yeast two-hybrid system  for the study of protein-protein interaction. [ 25 ]  The vectors used in yeast two-hybrid system contain fusion partners for two cloned genes that allow the transcription of a reporter gene when there is interaction between the two proteins expressed from the cloned genes.


Baculovirus , a rod-shaped virus which infects insect cells, is used as the expression vector in this system. [ 26 ]  Insect cell lines derived from  Lepidopterans  (moths and butterflies), such as  Spodoptera frugiperda , are used as host. A cell line derived from the  cabbage looper  is of particular interest, as it has been developed to grow fast and without the expensive serum normally needed to boost cell growth. [ 27 ] [ 28 ]  The  shuttle vector  is called bacmid, and gene expression is under the control of a strong promoter pPolh. [ 29 ]  Baculovirus has also been used with mammalian cell lines in the  BacMam  system. [ 30 ]


Baculovirus is normally used for production of  glycoproteins , although the glycosylations may be different from those found in vertebrates. In general, it is safer to use than mammalian virus as it has a limited host range and does not infect vertebrates without modifications.


Many plant expression vectors are based on the  Ti plasmid  of  Agrobacterium tumefaciens . [ 31 ]  In these expression vectors, DNA to be inserted into plant is cloned into the  T-DNA , a stretch of DNA flanked by a 25-bp direct repeat sequence at either end, and which can integrate into the plant genome. The T-DNA also contains the selectable marker. The  Agrobacterium  provides a mechanism for  transformation , integration of into the plant genome, and the promoters for its  vir  genes may also be used for the cloned genes.  Concerns over the transfer of bacterial or viral genetic material into the plant however have led to the development of vectors called intragenic vectors whereby functional equivalents of plant genome are used so that there is no transfer of genetic material from an alien species into the plant. [ 32 ]


Plant viruses may be used as vectors since the  Agrobacterium  method does not work for all plants. Examples of plant virus used are the  tobacco mosaic virus  (TMV),  potato virus X , and  cowpea mosaic virus . [ 33 ]  The protein may be expressed as a fusion to the coat protein of the virus and is displayed on the surface of assembled viral particles, or as an unfused protein that accumulates within the plant. Expression in plant using plant vectors is often constitutive, [ 34 ]  and a commonly used constitutive promoter in plant expression vectors is the  cauliflower mosaic virus  (CaMV) 35S promoter. [ 35 ] [ 36 ]


Mammalian expression vectors offer considerable advantages for the expression of mammalian proteins over bacterial expression systems - proper folding, post-translational modifications, and relevant enzymatic activity. It may also be more desirable than other eukaryotic non-mammalian systems whereby the proteins expressed may not contain the correct glycosylations. It is of particular use in producing membrane-associating proteins that require chaperones for proper folding and stability as well as containing numerous post-translational modifications. The downside, however, is the low yield of product in comparison to prokaryotic vectors as well as the costly nature of the techniques involved. Its complicated technology, and potential contamination with animal viruses of mammalian cell expression have also placed a constraint on its use in large-scale industrial production. [ 37 ]


Cultured mammalian cell lines such as the  Chinese hamster ovary (CHO) ,  COS , including human cell lines such as  HEK  and  HeLa  may be used to produce protein. Vectors are  transfected  into the cells and the DNA may be integrated into the genome by  homologous recombination  in the case of stable transfection, or the cells may be transiently transfected. Examples of mammalian expression vectors include the  adenoviral  vectors, [ 38 ]  the pSV and the pCMV series of plasmid vectors,  vaccinia  and  retroviral  vectors, [ 39 ]  as well as baculovirus. [ 30 ]  The promoters for  cytomegalovirus  (CMV) and  SV40  are commonly used in mammalian expression vectors to drive gene expression. Non-viral promoter, such as the elongation factor (EF)-1 promoter, is also known. [ 40 ]


E. coli   cell lysate  containing the cellular components required for transcription and translation are used in this  in vitro  method of protein production. The advantage of such system is that protein may be produced much faster than those produced  in vivo  since it does not require time to culture the cells, but it is also more expensive. Vectors used for  E. coli  expression can be used in this system although specifically designed vectors for this system are also available. Eukaryotic cell extracts may also be used in other cell-free systems, for example, the  wheat germ  cell-free expression systems. [ 41 ]  Mammalian cell-free systems have also been produced. [ 42 ]


Expression vector in an expression host is now the usual method used in laboratories to produce proteins for research. Most proteins are produced in  E. coli , but for glycosylated proteins and those with disulphide bonds, yeast, baculovirus and mammalian systems may be used.


Most protein  pharmaceuticals  are now produced through recombinant DNA technology using expression vectors. These peptide and protein pharmaceuticals may be hormones, vaccines, antibiotics, antibodies, and enzymes. [ 43 ]   The first human recombinant protein used for disease management, insulin, was introduced in 1982. [ 43 ]  Biotechnology allows these peptide and protein pharmaceuticals, some of which were previously rare or difficult to obtain, to be produced in large quantity. It also reduces the risks of contaminants such as host viruses, toxins and  prions . Examples from the past include  prion  contamination in  growth hormone  extracted from  pituitary glands  harvested from human cadavers, which caused  Creutzfeldt–Jakob disease  in patients receiving treatment for  dwarfism , [ 44 ]  and viral contaminants in clotting  factor VIII  isolated from human blood that resulted in the transmission of viral diseases such as  hepatitis  and  AIDS . [ 45 ] [ 46 ]  Such risk is reduced or removed completely when the proteins are produced in non-human host cells.


In recent years, expression vectors have been used to introduce specific genes into plants and animals to produce  transgenic  organisms, for example in  agriculture  it is used to produce  transgenic plants . Expression vectors have been used to introduce a  vitamin A  precursor,  beta-carotene , into rice plants. This product is called  golden rice . This process has also been used to introduce a gene into plants that produces an  insecticide , called  Bacillus thuringiensis toxin  or  Bt toxin  which reduces the need for farmers to apply insecticides since it is produced by the modified organism. In addition expression vectors are used to extend the ripeness of tomatoes by altering the plant so that it produces less of the chemical that causes the tomatoes to rot. [ 47 ]  There have been  controversies  over using expression vectors to modify crops due to the fact that there might be unknown health risks, possibilities of companies patenting certain  genetically modified food  crops, and ethical concerns. Nevertheless, this technique is still being used and heavily researched.


Transgenic animals  have also been produced to study animal biochemical processes and human diseases, or used to produce pharmaceuticals and other proteins. They may also be engineered to have advantageous or useful traits.  Green fluorescent protein  is sometimes used as tags which results in animal that can fluoresce, and this have been exploited commercially to produce the fluorescent  GloFish .


Gene therapy  is a promising treatment for a number of diseases where a "normal" gene carried by the vector is inserted into the genome, to replace an "abnormal" gene or supplement the expression of particular gene. Viral vectors are generally used but other nonviral methods of delivery are being developed. The treatment is still a risky option due to the viral vector used which can cause ill-effects, for example giving rise to  insertional mutation  that can result in cancer. [ 48 ] [ 49 ]  However, there have been promising results. [ 50 ] [ 51 ]


In genetic engineering,  floxing  refers to the  insertion  of a  DNA  sequence (which is then said to be  floxed ) between two  LoxP  sequences, creating an artificial  gene cassette  which can then be conditionally  deleted  (knocked out),  translocated , or  inverted  in a process called  Cre-Lox recombination . [ 1 ]   Recombination  between LoxP sites is catalysed by  Cre recombinase . The term "floxing" is a portmanteau constructed from the phrase "flanking/flanked by LoxP".


The floxing method is essential in the development of scientific model systems as it allows researchers to have spatial and temporal alteration of gene expression. [ 2 ]  The Cre-Lox system is widely used to manipulate gene expression in model organisms such as mice in order to study human diseases and drug development. [ 3 ]  For example, using the Cre-Lox system, researchers are able to study  oncogenes  and  tumor suppressor  genes and their role in the development and progression of  cancer  in mouse models. [ 4 ]


Floxing a gene allows it to be deleted (knocked out), [ 5 ] [ 6 ]  translocated or inserted [ 7 ]  (through various mechanisms in Cre-Lox recombination).


The floxing of genes is essential in the development of scientific model systems as it allows spatial and temporal alteration of gene expression. In layman's terms, the gene can be knocked-out (inactivated) in a specific tissue  in vivo , at a specific time chosen by the scientist. The scientist can then evaluate the effects of the knocked-out gene and identify the gene's normal function. [ 8 ]  This is different from having the gene absent starting from conception, whereby inactivation or loss of genes that are essential for the development of the organism may interfere with the normal function of cells and prevent the production of viable offspring. [ 9 ]


Deletion events are useful for performing  gene editing  experiments through precisely removing segments of or even whole genes. Deletion requires floxing of the segment of interest with loxP sites which face the same direction. The Cre recombinase will detect the unidirectional loxP sites and excise the floxed segment of DNA. [ 10 ]  The successfully edited clones can be selected using a selection marker which can be removed using the same Cre-LoxP system. [ 10 ]  The same mechanism can be used to create  conditional alleles  by introducing an  FRT/Flp  site which accomplishes the same mechanism but with a different enzyme.


Inversion events are useful for inactivating a gene or DNA sequence without actually removing it, and thereby maintaining a consistent amount of genetic material. The inverted genes are not often associated with abnormal  phenotypes , meaning the inverted genes are generally viable. [ 11 ]  Cre-LoxP recombination that results in inversion requires loxP sites flanking the gene of interest, with the loxP sites oriented towards each other as  inverted repeats . By undergoing Cre recombination, the region flanked by the loxP sites will become inverted, i.e. re-inserted in the same position but in reverse orientation; [ 12 ]  this process is not permanent and can be reversed. [ 13 ]


Translocation events occur when the loxP sites flank genes on two different DNA molecules in a unidirectional orientation. Cre recombinase is then used to generate a translocation between the two DNA molecules, exchanging the genetic material from one DNA molecule to the other, forming a simultaneous translocation of both floxed genes. [ 12 ] [ 14 ]


Cardiomyocytes  (heart muscle tissue) have been shown to express a type of Cre recombinase that is highly specific to cardiomyocytes and can be used by researchers to perform highly efficient recombinations. This is achieved by using a type of Cre whose expression is driven by the

 α

 {\displaystyle \alpha }

 -myosin heavy chain promoter (

 α

 {\displaystyle \alpha }

 -MyHC) . These recombinations are capable of disrupting genes in a manner that is specific to heart tissue  in vivo  and allows for the creation of conditional knockouts of the heart, mostly for use as controls. [ 15 ]  For example, using the  Cre recombinase  with the

 α

 {\displaystyle \alpha }

 -MyHC promoter causes the floxed gene to be inactivated in the heart alone. Further, these knockouts can be made inducible. In several mouse studies,  tamoxifen  is used to induce the expression of  Cre recombinase . [ 2 ]  In this case,  Cre recombinase  is fused to a portion of the mouse  estrogen receptor  (ER) which contains a mutation within its  ligand binding domain (LBD) . The mutation renders the receptor inactive, which leads to incorrect localization through its interactions with chaperone proteins such as heat shock protein 70 and 90 ( Hsp70  and  Hsp90 ). Tamoxifen binds to Cre-ER and disrupts its interactions with the chaperones, which allows the Cre-ER fusion protein to enter the nucleus and perform recombination on the floxed gene. [ 16 ] [ 17 ]  Additionally,  Cre recombinase  can be induced by heat when under the control of specific heat shock elements (HSEs). [ 18 ] [ 19 ]


In  genetics ,  Flp- FRT  recombination  is a  site-directed recombination  technology,  increasingly used to manipulate an organism's DNA under controlled conditions  in vivo . It is analogous to  Cre- lox  recombination  but involves the recombination of sequences between short flippase recognition target ( FRT ) sites by the  recombinase  flippase ( Flp ) derived from the 2 μ plasmid of  baker's yeast   Saccharomyces cerevisiae .


The 34bp minimal FRT site sequence has the sequence


for which flippase (Flp) binds to both 13-bp 5'-GAAGTTCCTATTC-3' arms flanking the 8 bp spacer, i.e. the  site-specific recombination  (region of crossover) in reverse orientation.  FRT -mediated cleavage occurs just ahead from the asymmetric 8bp core region ( 5' tctagaaa 3' ) on the top strand and behind this sequence on the bottom strand. [ 1 ]  Several variant  FRT  sites exist, but recombination can usually occur only between two  identical   FRT s but generally not among  non-identical  ("heterospecific")  FRT s. [ 2 ] [ 3 ]


In yeast, this enzyme corrects decreases in 2 μ plasmid copy number caused by rare missegregation events. It does so by causing recombination between the two inverted repetitions on the 2 μ plasmid during  DNA replication . This changes the direction of one replication fork, causing multiple rounds of copying in a single initiation. [ 4 ]


Senecoff  et al.  (1987) investigated how nucleotide substitutions within the FRT affected the efficacy of the FLP-mediated recombination. The authors induced base substitutions in either one or both of the FRT sites and tested the concentration of FLP required to observe site-specific recombinations. Every base substitution was performed on each of the thirteen nucleotides within the FRT site (example G to A, T, and C). First, the authors showed that most mutations within the FRT sequence cause minimal effects if present within only one of the two sites. If mutations occurred within both sites, the efficiency of FLP is dramatically reduced. Second, the authors provided data for which nucleotides are most crucial for the binding of FLP and efficacy of the site-specific recombination. If the first nucleotide in both FRT sites is substituted to a cytosine (G to C), the third nucleotide is substituted for a thymine (A to T), or the seventh nucleotide is substituted for an adenosine (G to A), then the efficacy of the FLP-mediated site-specific recombination is reduced more than 100-fold. [ 5 ]  While a base substitution of any of the aforementioned nucleotides in only one of the FRT sites led to a ten-fold, ten-fold, and five-fold reduction of efficacy, respectively. [ 5 ]


Base Substitutions in the capitalized nucleotides led to the greatest reduction in FLP-mediated site-specific recombination (Wildtype x mutant and mutant x mutant):


Many available constructs include an additional arm sequences (5'- GAAGTTCCTATTCC -3') one base pair away from the upstream element and in the same orientation:


This segment is dispensable for excision but essential for integration, including  Recombinase-mediated cassette exchange . [ 6 ]


Because the recombination activity can be targeted to a selected organ, or a low level of recombination activity can be used to consistently alter the DNA of only a subset of cells, Flp- FRT  can be used to construct  genetic mosaics  in multicellular organisms.  Using  this technology , the loss or alteration of a gene can be studied in a given target organ of interest, even in cases where experimental animals would not survive the loss of this gene in other organs ( spatial control ). The effect of altering a gene can also be studied over time, by using an  inducible promoter  to trigger the recombination activity late in development ( temporal control ) - this prevents the alteration.


The Flp protein, much like Cre, is a tyrosine family site-specific recombinase. [ 7 ]  This family of recombinases performs its function via a type IB topoisomerase mechanism causing the recombination of two separate strands of DNA. [ 7 ]  Recombination is carried out by a repeated two-step process. The initial step causes the creation of a  Holliday junction  intermediate. The second step promotes the resulting recombination of the two complementary strands. As their family name suggests, a highly conserved tyrosine nucleophile cleaves the DNA strands. [ 7 ]  The nucleophilic properties of the tyrosine attack and bind to the 3'-phosphate at the point of DNA cleavage. [ 7 ]  The resulting 5'-hydroxyl group of the cleaved DNA acts as the nucleophile and attacks the 3'-phosphate on the complementarily cleaved DNA strand, resulting in successful recombination. [ 7 ]  Outside of the tyrosine residue, there is a conserved catalytic pentad. This pentad is made up of a lysine (Lysβ), two arginines (Arg I and II), a histidine (His-II), and a histidine/tryptophan (His/Trp-III) that comprises a mandatory and highly conserved constellation of residues for the active site of Flp and Cre (along with other IB topiosomerases). [ 7 ]  These other residues are crucial to the correct orientation of Flp binding and positioning on the DNA strands.


Initial application of the FLP-FRT recombinase did not work in mammals. The FLP protein was  thermolabile  (denatured at elevated temperatures) and therefore was not useful in the mammalian model due to elevated body temperatures of these model systems. However, due to patents and restrictions on the use of Cre-Lox recombination, great interest was taken to produce a more thermostable FLP-FRT cassette. Some of the first results were produced by Buchholz  et al.  (1997) by utilizing cycling  mutagenesis  in  Escherichia coli  . In their research, the authors transfected  E. coli c ells with two plasmids: one coding for randomly mutated FLP proteins downstream of an arabinose promoter and another containing a lacZ gene promoter within a FRT cassette. The  E. coli  were grown on arabinose plates at 37 °C and 40 °C, and if recombination occurred, the lacZ expression would be attenuated, and the colonies would appear white. White colonies were selected from each generation and grown on a new arabinose plates at same previous temperatures for eight generations. [ 8 ]  After recombination was confirmed by western-blotting and the mutated FLP genes were sequenced, this  e ighth generation  FLP  protein (FLPe) was transfected into mammalian cell culture, and recombination in mammalian cells was confirmed. [ 8 ]  This variant of FLP only has 4 amino acid substitutions: P2S, L33S, Y108N, and S294P.


Genetic mosaicism  occurs within an organism when similar cell types express different phenotypes due to dissimilar genotypes at specific loci. Simply put, this occurs when one organism contains different genotypes, which is usually rare in nature. However, this can be easily (and problematically) produced using FLP-FRT recombination. If two different FRT sites are present within a cell, and FLP is present in appropriate concentrations, the FRT cassette will continue to be excised and inserted between the two FRT sites. This process will continue until the FLP proteins fall below the required concentrations resulting in cells within an organism possessing different genotypes. [ 9 ]  This has been seen from fruit flies to mice and is indiscriminate against specific chromosomes (somatic and sex) or cell types (somatic and germline). [ 9 ] [ 10 ]


Before the publication of Dymecki  et al . (1998), Cre recombinase had been used for cell-fate mapping of neuronal progenitors in mice using the  En2   promoter. [ 11 ]  Thus, the authors of Dymecki  et al . (1998) theorized that FLP recombinase could be utilized in a similar fashion with a similar efficacy as Cre recombinase in mice. The authors created a two transgenic mouse lines: a neuronal Wnt1::Flp fusion line and a line that possessed the FRT cassette flanking the 18th exon of  tm1Cwr . [ 9 ]  The authors chose this exon for excision because if it is excised, its results in a null phenotype. The authors mated the two lines and allowed the progeny to reach adulthood before the progeny were sacrificed. RNA extraction was performed on neuronal, muscular, enteric, and tail tissue.  Reverse-transcription PCR  and  northern blotting  confirmed the excision of the 18th exon of  tm1Cwr  abundantly in the brain tissue and moderately in muscular tissue (due to  Scwhann cells  within the muscle). As expected, excision was not seen in other tissues. The authors saw equal, if not better, efficiency of the FLP recombinase in cell-fate determination than Cre recombinase.


To date, Flp recombinase has been utilized many times in  D. melanogaster.  A comparison of Flp recombinase to Cre recombinase in  D. melanogaster  was published by Frickenhaus  et al.  (2015) [ 12 ] .  The authors of Frickenhaus  et al.  (2015) had a two-fold objective: characterize and compare the efficacy of Flp recombinase "knock-out" to Cre recombinase "knock-out" and RNAi knockdown and reveal the function of  cabeza  (caz), the fly ortholog to  FUS , in the neurons and muscle tissue of  D. melanogaster .  FUS  has been strongly implicated in  amyotrophic lateral sclerosis  (ALS) and  frontotemporal dementia  in humans [ 12 ]   .  The authors used an  elav - Gal4/UAS -Flp or Cre system to express the recombinase specifically in neurons and a  Mef2 -Gal4/UAS-Flp or Cre system to express it specifically in muscles. [ 12 ]  The authors conclude that the Flp recombinase "knock-out" tool is more effective than both RNAi and Cre recombinase for the purpose of knocking out specific genes in specific tissues or cell lines due to the lack of the leaky expression seen in both the Cre protein and the RNAi transcript. Also, the authors witnessed a toxicity to the Cre protein that is not seen with the Flp protein. [ 12 ]


The efficacy of the FLPe recombinase system was evaluated in zebrafish by Wong  et al . (2009). [ 13 ]  Embryos, that were hemizygous for a FRT-flanked enhanced green fluorescent protein (EGFP) downstream of a muscle-specific promoter, were injected with the FLPe protein. Without FLPe, these embryoes should express  EGFP  in all muscle tissue, and if crossed with a wild type strand, 50% of the resulting progeny should also express EGFP in muscle tissue. Embryoes, injected with FLPe, had significantly reduced expression of EGFP in muscle tissue, and mosaicism was seen also. [ 13 ]  When these embryoes reached adulthood, they were mated with a wild type strain, and the resulting clutches had significantly less progeny that expressed the EGFP in the muscle tissue (0-4%). [ 13 ]  These results show, that not only is FLPe highly effective in somatic cells, it is also highly effective in the germline of zebrafish.


Phytosensors are genetically modified plants that can report the presence of biotic or abiotic contaminants. [ 14 ]  The production of these engineered plants have great agricultural and laboratory promise. However, creating an appropriate reporter vector has shown to be problematic.  cis -regulatory elements play a major role in the transcriptional activation of genes in plants, and many are not well understood. Many phytosensors either under express their reporter genes or report false-positives due to synthetic promoters. [ 14 ]  The authors of Rao  et al.  (2010) utilized the FLP recombinase tool for the production of a highly efficient phytosensors. The authors used a heat shock promoter to induce the production of FLP while a FRT-flanked vector separated the CaMV 35S promoter from the beta-glucuronidase gene (GUS). When the plants were exposed to heat-shock, FLP-induction led to the excision of the FRT-flanked vector, effectively moving the GUS gene directly downstream of the CaMV 35S promoter. The activation of GUS led to the leaves of the plants to change from green to blue; thus, the phytosensor effectively reported stress to the model system! [ 14 ]


RNA interference (RNAi) has caused a paradigm shift in the expression of genes and potential gene knockouts in Eukaryotes. Before the production of the GRIM expression system, the creation of RNAi vectors was expensive and time consuming. The vectors were produced by the traditional copy-and-paste molecular cloning method. [ 15 ]  Garwick-Coppens  et al . (2011) developed a much more efficient method for the production of RNAi vectors in which expression of the RNAi can be knocked-in using Cre-recombinase and knocked-out using Flp-recombinase. The novel GRIM Expression System allows for the much faster generation of expression vectors containing artificial RNAi constructs. [ 15 ]  The authors went on to show that their expression system works quite effectively in human embryonic kidneys (HEK) cells, a common human immortalized cell line in molecular research.


Functional cloning  is a  molecular cloning  technique that relies on prior knowledge of the encoded  protein ’s  sequence  or  function  for  gene  identification. [ 1 ] [ 2 ] [ 3 ]  In this assay, a  genomic  or  cDNA library  is screened to identify the genetic sequence of a protein of interest.  Expression  cDNA libraries may be screened with  antibodies  specific for the protein of interest or may rely on selection via the protein function. [ 1 ]  Historically, the amino acid sequence of a protein was used to prepare degenerate oligonucleotides which were then  probed  against the library to identify the gene encoding the protein of interest. [ 2 ] [ 3 ]  Once candidate clones carrying the gene of interest are identified, they are  sequenced  and their identity is confirmed. This method of cloning allows researchers to screen entire  genomes  without prior knowledge of the location of the gene or the genetic sequence. [ 1 ]


This technique can be used to identify genes that encode similar proteins from one organism to another. [ 4 ]  Similarly, this technique can be paired with  metagenomic  libraries to identify novel genes and proteins that perform similar functions, such as the identification of novel  antibiotics  by screening for  beta-lactamase  activity or selecting for growth in the presence of  penicillin . [ 5 ]


The workflow of a functional cloning experiment varies depending on the source of genetic material, the extent of prior knowledge of the protein or gene of interest and the ability to screen for the protein function. In general, a functional cloning experiment consists of four steps: 1) sample collection, 2) library preparation, 3) screening or selection and 4)  sequencing .


Genetic material is collected from a particular cell type, organism or environmental sample relevant to the biological question. In functional cloning,  mRNA  is commonly isolated and  cDNA  is prepared from the isolated mRNA ( RNA extraction ). [ 6 ]  In certain circumstances  genomic DNA  may be isolated, particularly when environmental samples are used as the source of genetic material. [ 1 ]


If the starting material is  genomic DNA , the DNA is sheared to produce fragments of appropriate length for the  vector  of choice. The DNA fragments or cDNA are then treated with  restriction endonucleases  and  ligated  to a  plasmid  or chromosomal vectors. In the case of assays that screen for the protein or for its function, an  expression vector  is used to ensure that the gene product is expressed. The vector choice will depend on the origin of the DNA or cDNA to ensure proper expression and to ensure that the encoded gene will fall within the limits of the vector's insert size. [ 7 ]


The choice of host is important to ensure that the  codon usage  will be similar to the donor organism. The host will also need to guarantee that the proper  post-translational modifications  and  protein folding  will occur to enable proper functioning of the expressed proteins. [ 7 ]


The method of screening the prepared genomic or cDNA libraries for the gene of interest is highly variable depending on the experimental design and biological question. One method of screening is to  probe  colonies via  Southern blotting  with degenerate oligonucleotides prepared from the amino acid sequence of the query protein. [ 8 ]  In expression libraries, the protein of interest can be identified by screening with an antibody specific for the query protein via  Western blotting  to identify colonies carrying the gene of interest. In other circumstances, a specific assay can be used to screen or select for the protein's activity. [ 1 ]  For example, genes conferring  antibiotic resistance  can be selected by growing the colonies of the library on media containing a specified  antibiotic . [ 5 ]  Another example is screening for  enzymatic activity  by incubating with a substrate that is catalyzed to a colorimetric compound that can easily be visualized. [ 9 ]


The final step of functional cloning is to  sequence  the DNA or cDNA from the clones that were successfully identified in the screen or selection step. The sequence can then be annotated and used for downstream applications, such as  protein expression  and  purification  for industrial applications. [ 10 ]


The advantages of functional cloning include the ability to screen for novel genes with desired applications in organisms that cannot be cultured, particularly from bacterial or viral specimens. [ 1 ]  Additionally, genes encoding proteins with related functions can be identified when there is low sequence similarity due to the ability to screen for the protein function alone. Functional cloning allows for gene identification without prior knowledge of the organism's genome sequence or position of the gene within the genome. [ 1 ]


As with other cloning techniques, vector and host choice affect the success of gene identification via functional cloning due to cloning bias. The vector must have an insert size that will accommodate the entire DNA sequence of the expressed protein. Additionally, in expression vectors the  promoters  and  terminators  must function within the chosen host organism. The host choice may affect  transcription  and  translation  due to differing  codon usage , transcriptional and translational machinery or  post-translational modifications  within the host. [ 7 ] [ 1 ]


Other limitations include the labour-intensive library preparation and potential screens which can be both expensive and time-consuming. [ 7 ]


Positional cloning  is another molecular cloning technique for identification of a gene of interest. This method uses exact chromosomal location instead of function to guide gene identification. [ 11 ]  Because of this, this method focuses on all the genetic material at a chromosomal  locus  and makes no assumptions about function. [ 11 ]  In  model organisms  such as  mice  or  yeast , this method is used more frequently as the information about the position of a gene of interest can be obtained from the  sequenced genome . However, this method becomes much more cumbersome when sequence information is not available. In this case,  linkage analysis  can also be used. [ 11 ]  Functional cloning on the other hand is more readily used in organisms such as  bacterial pathogens  that are  viable but nonculturable  and where sequence data is not available but gene homology or protein function is still of interest. [ 11 ]


A way to differentiate between functional and positional cloning is to visualize genes as words. Functional cloning is like using a thesaurus to look up words and selecting for new words that have the same meanings (or functions). [ 12 ]  Positional cloning is more like picking a specific page of a dictionary and then browsing only that page for any words of interest. [ 12 ]


With the advent of  sequencing  technology becoming cheaper and cheaper, it is now more feasible to sequence an unknown genome and then computationally determine  homology  instead of screening. [ 13 ]  This brings the added benefit of being able to screen for multiple genes of interest at the same time and reduces experimental time. It also allows one to avoid labour-intensive cloning procedures as well. [ 14 ]  However, if this route is taken, there are other biases and hurdles one must consider. By using sequenced data, one is able to screen based on homology alone. [ 1 ]  A function-based approach thus allows for discovery of novel enzymes whose functions would not have been predicted based on DNA sequence alone. [ 1 ]  Therefore, while sequencing is less labour-intensive experimentally, it can also lead to missed genes of interest due to differing sequence homology in genes of related function.


Gibson assembly  is a quick cloning method that uses three primary enzymes; 5'  exonuclease ,  polymerase  and  ligase . [ 15 ]  The exonuclease digests the 5' end of DNA fragments leaving a 3' overhang. [ 15 ]  If there is significant homology (20-40 bp) on each end of the DNA insert, it can anneal with a complementary backbone. [ 15 ]  Afterwards the polymerase can fill in the gaps while ligase fuses the nicks at the end. [ 15 ]  This method greatly increases the rate of cloning and success rate of cloning into a vector backbone. [ 15 ]  However, it requires the DNA fragment to have significant homology with the plasmid. [ 15 ]  For this reason, knowledge of the sequence being cloned must be known beforehand. This is not a requirement with functional cloning.


TOPO Cloning  is a cloning method that uses  Taq polymerase . [ 16 ]  This is because Taq leaves a single  adenosine  overhang on the 3’ end of  PCR  reaction products. [ 16 ]  Utilizing this knowledge, backbones with a 5’  thymine  overhang can be used for cloning purposes. [ 16 ]  In this case knowledge of the fragment being cloned must be known to be able to make PCR primers for it and the number of TOPO Cloning compatible vectors is relatively small. However, it provides the advantage that reactions only take about 5 minutes to do. [ 16 ]


Gateway recombination cloning  is a cloning method in which a DNA fragment is moved from one plasmid backbone to another via a single  homologous recombination  event. [ 17 ]  However, for this method to work, the DNA fragment of interest must be flanked by recombination sites. [ 17 ]  While this method isn't strictly an alternative, it does allow the movement of DNA fragments from one plasmid to another quicker than creating a whole new genomic library. The reason this method may be used in conjunction with functional cloning is to put a library under a different promoter or on a backbone with a different selection marker. [ 17 ]  This can come in handy if an individual wants to try functional cloning in a wide range of bacteria to try to combat the issue with  codon bias . [ 17 ] [ 7 ]


Metagenomics  is one of the largest fields that commonly uses functional cloning. Metagenomics studies all the genetic material from a specific environmental sample, such as the gut microbiome or lake water. [ 1 ]   Functional libraries  are created that contain DNA fragments from the environment. [ 1 ]  As the original bacterium that a DNA sequence originated from cannot be easily detected, creating metagenomic functional libraries possesses advantages. Less than 1% of all bacteria are easily cultured in the lab, leaving a large percentage of bacteria that cannot be grown. [ 18 ]  By using functional libraries, the gene functions of unculturable bacteria can still be studied. [ 1 ]  Furthermore, these uncultured microbes provide a source for the discovery of novel enzymes with biotechnological applications. Some novel proteins that have been discovered from marine environments include enzymes such as proteases, amylases, lipases, chitinases, deoxyribonucleases and phosphatases. [ 19 ]


There are situations in which it is imperative to determine if a gene homolog from one source is present in another organism. For example, identification of novel  DNA polymerases  for  polymerase chain reaction (PCR)  reactions which synthesize DNA molecules from deoxyribonucleotides. [ 20 ]  While human polymerase optimally works at 37 °C (99 °F), DNA does not denature until 94–98 °C (201–208 °F). [ 20 ]  This poses a problem as at these temperatures the human DNA polymerase would denature during the denaturation step of the PCR reaction resulting in a non-functioning polymerase protein and a failed PCR. To combat this a DNA polymerase from a  thermophile , or bacteria that grows at high temperatures, could be used. An example is  Taq polymerase  which comes from the thermophilic bacterium  Thermus aquaticus . [ 20 ]  One could set up a functional cloning screen to find homologous polymerases that have the added advantage of being thermostable at high temperatures.


With this in mind, 3173 Polymerase, another polymerase enzyme, now commonly used in  RT-PCR  reactions was discovered using the above theory. [ 21 ]  In RT-PCR reactions, two separate enzymes are commonly used. The first is a retroviral  reverse transcriptase  to convert  RNA  to  cDNA . [ 21 ]  The second is a thermostable DNA polymerase to amplify the target sequence. [ 21 ]  3173 Polymerase is able to perform both enzymatic functions resulting in a better option for RT-PCR. [ 21 ]  The enzyme was discovered using functional cloning from a viral host originally found in Octopus hot springs (93 °C) in Yellowstone National Park. [ 21 ]


One of the ongoing challenges of treating bacterial infections is  antibiotic resistance  which commonly arises when patients do not take their full treatment of medication and hence allow bacteria to develop resistance to antibiotics over time. [ 22 ]  To understand how to combat antibiotic resistance it is important to understand how the bacterial genome is evolving and changing in healthy individuals with no recent usage of antibiotics to provide a baseline. [ 22 ]  Using a functional cloning-based technique, DNA isolated from human  microflora  were cloned into expression vectors in  Escherichia coli . [ 22 ]  Afterwards, antibiotics were applied as a screen. [ 22 ]  If a plasmid contained a gene insert that provided antibiotic resistance the cell survived and was selected on the plate. [ 22 ]  If the insert provided no resistance, the cell died and did not form a colony. [ 22 ]  Based on selection of cell colonies that survived, a better picture of genetic factors contributing to antibiotic resistance were pieced together. Most of the resistance genes that were identified were previously unknown. [ 22 ]  By using a functional cloning-based technique one is able to elucidate genes giving rise to antibiotic resistance to better understand treatment for bacterial infections.


A  gene drive  is a natural process [ 1 ]  and technology of  genetic engineering  that propagates a particular suite of genes throughout a population [ 2 ]  by altering the probability that a specific allele will be transmitted to offspring (instead of the  Mendelian 50%  probability). Gene drives can arise through a variety of mechanisms. [ 3 ] [ 4 ]  They have been proposed to provide an effective means of genetically modifying specific populations and entire species.


The technique can employ adding, deleting, disrupting, or modifying genes. [ 5 ] [ 6 ]


Proposed applications include exterminating insects that carry pathogens (notably mosquitoes that transmit  malaria ,  dengue , and  zika  pathogens), controlling  invasive species , or eliminating  herbicide  or  pesticide resistance . [ 7 ] [ 5 ] [ 8 ] [ 9 ]