License applications for the release of all genetically modified organisms in Australia is overseen by the  Office of the Gene Technology Regulator , while regulation is provided by the  Therapeutic Goods Administration  for GM medicines or  Food Standards Australia New Zealand  for GM food. The individual state governments can then assess the impact of release on markets and trade and apply further legislation to control approved genetically modified products. [ 82 ] [ 83 ]  The Australian  Parliament  relaxed the definition of GMOs, in 2019, to exclude certain GMOs from GMO  regulation  and government oversight. [ 84 ]


In  Singapore ,  synthetic biology  products are regulated as if they were  genetically modified organisms  under the  Biological Agents and Toxins Act . For further review see Trump 2017. [ 85 ]


In Saudi Arabia's  Neom  project genetically engineered agriculture is legal, encouraged, and is funded by the government as an integral part of the project. [ 86 ]


One of the key issues concerning regulators is whether GM products should be labeled. Labeling can be mandatory up to a threshold GM content level (which varies between countries) or voluntary. A study investigating voluntary labeling in South Africa found that 31% of products labeled as GMO-free had a GM content above 1.0%. [ 4 ]  In Canada and the United States labeling of GM food is voluntary, [ 5 ]  while in Europe all food (including  processed food ) or  feed  which contains greater than 0.9% of approved GMOs must be labelled. [ 6 ]  In the US state of Oregon., voters rejected Measure 27, which would have required labeling of all genetically modified foods. [ 91 ]  Japan, Malaysia, New Zealand, and Australia require labeling so consumers can exercise choice between foods that have genetically modified, conventional or organic origins. [ 92 ]


The Cartagena Protocol sets the requirements for the international trade of GMOs between countries that are signatories to it.  Any shipments contain genetically modified organisms that are intended to be used as feed, food or for processing must be identified and a list of the transgenic events be available.


"Substantial equivalence" is a starting point for the safety assessment for GM foods that is widely used by national and international agencies—including the Canadian Food Inspection Agency, Japan's Ministry of Health and Welfare and the U.S. Food and Drug Administration, the United Nation’s Food and Agriculture Organization, the  World Health Organization  and the OECD. [ 93 ]


A quote from FAO, one of the agencies that developed the concept, is useful for defining it: "Substantial equivalence embodies the concept that if a new food or food component is found to be substantially equivalent to an existing food or food component, it can be treated in the same manner with respect to safety (i.e., the food or food component can be concluded to be as safe as the conventional food or food component)". [ 94 ]   The concept of substantial equivalence also recognises the fact that existing foods often contain toxic components (usually called  antinutrients ) and are still able to be consumed safely—in practice there is some tolerable chemical risk taken with all foods, so a comparative method for assessing safety needs to be adopted. For instance, potatoes and tomatoes can contain toxic levels of respectively, solanine and alpha-tomatine alkaloids. [ 95 ] [ 96 ]


To decide if a modified product is substantially equivalent, the product is tested by the manufacturer for unexpected changes in a limited set of components such as toxins, nutrients, or allergens that are present in the unmodified food.  The manufacturer's data is then assessed by a regulatory agency, such as the U.S.  Food and Drug Administration . That data, along with data on the genetic modification itself and resulting proteins (or lack of protein), is submitted to regulators.  If regulators determine that the submitted data show no significant difference between the modified and unmodified products, then the regulators will generally not require further food safety testing.  However, if the product has no natural equivalent, or shows significant differences from the unmodified food, or for other reasons that regulators may have (for instance, if a gene produces a protein that had not been a food component before), the regulators may require that further safety testing be carried out. [ 39 ]


A 2003 review in  Trends in Biotechnology  identified seven main parts of a standard safety test: [ 97 ]


There has been discussion about applying new  biochemical  concepts and methods in evaluating substantial equivalence, such as metabolic profiling and protein profiling.  These concepts refer, respectively, to the complete measured biochemical spectrum (total fingerprint) of compounds (metabolites) or of proteins present in a food or crop.  The goal would be to compare overall the biochemical profile of a new food to an existing food to see if the new food's profile falls within the range of natural variation already exhibited by the profile of existing foods or crops.  However, these techniques are not considered sufficiently evaluated, and standards have not yet been developed, to apply them. [ 98 ]


Transgenic animals  have genetically modified DNA. Animals are different from plants in a variety of ways—biology, life cycles, or potential environmental impacts. [ 99 ]  GM plants and animals were being developed around the same time, but due to the complexity of their biology and inefficiency with laboratory equipment use, their appearance in the market was delayed. [ 100 ]


There are six categories that genetically engineered (GE) animals are approved for: [ 101 ]


Domingo, José L.; Bordonaba, Jordi Giné (2011).  "A literature review on the safety assessment of genetically modified plants"   (PDF) .  Environment International .  37  (4):  734– 742.  doi : 10.1016/j.envint.2011.01.003 .  PMID   21296423 .  In spite of this, the number of studies specifically focused on safety assessment of GM plants is still limited. However, it is important to remark that for the first time, a certain equilibrium in the number of research groups suggesting, on the basis of their studies, that a number of varieties of GM products (mainly maize and soybeans) are as safe and nutritious as the respective conventional non-GM plant, and those raising still serious concerns, was observed. Moreover, it is worth mentioning that most of the studies demonstrating that GM foods are as nutritional and safe as those obtained by conventional breeding, have been performed by biotechnology companies or associates, which are also responsible of commercializing these GM plants. Anyhow, this represents a notable advance in comparison with the lack of studies published in recent years in scientific journals by those companies.


Krimsky, Sheldon (2015).  "An Illusory Consensus behind GMO Health Assessment"   (PDF) .  Science, Technology, & Human Values .  40  (6):  883– 914.  doi : 10.1177/0162243915598381 . Archived from  the original   (PDF)  on 2019-08-31.  I began this article with the testimonials from respected scientists that there is literally no scientific controversy over the health effects of GMOs. My investigation into the scientific literature tells another story.


Panchin, Alexander Y.; Tuzhikov, Alexander I. (January 14, 2016). "Published GMO studies find no evidence of harm when corrected for multiple comparisons".  Critical Reviews in Biotechnology .  37  (2):  213– 217.  doi : 10.3109/07388551.2015.1130684 .  ISSN   0738-8551 .  PMID   26767435 .  Here, we show that a number of articles some of which have strongly and negatively influenced the public opinion on GM crops and even provoked political actions, such as GMO embargo, share common flaws in the statistical evaluation of the data. Having accounted for these flaws, we conclude that the data presented in these articles does not provide any substantial evidence of GMO harm.   The presented articles suggesting possible harm of GMOs received high public attention. However, despite their claims, they actually weaken the evidence for the harm and lack of substantial equivalency of studied GMOs. We emphasize that with over 1783 published articles on GMOs over the last 10 years it is expected that some of them should have reported undesired differences between GMOs and conventional crops even if no such differences exist in reality.


Reverse genetics  is a method in  molecular genetics  that is used to help understand the function(s) of a  gene  by analysing the  phenotypic  effects caused by  genetically engineering  specific  nucleic acid sequences  within the gene. The process proceeds in the opposite direction to  forward genetic  screens of  classical genetics . While forward genetics seeks to find the genetic basis of a  phenotype  or trait, reverse genetics seeks to find what phenotypes are controlled by particular genetic sequences.


Automated  DNA sequencing  generates large volumes of  genomic  sequence  data  relatively rapidly. Many genetic sequences are discovered in advance of other, less easily obtained, biological information. Reverse genetics attempts to connect a given genetic sequence with specific effects on the organism. [ 1 ]  Reverse genetics systems can also allow the recovery and generation of infectious or defective viruses with desired mutations. [ 2 ]  This allows the ability to study the virus  in vitro  and  in vivo .


In order to learn the influence a sequence has on phenotype, or to discover its biological function, researchers can engineer a change or disrupt the  DNA .  After this change has been made a researcher can look for the effect of such alterations in the whole  organism . There are several different methods of reverse genetics:


Site-directed mutagenesis  is a sophisticated technique that can either change regulatory regions in the  promoter  of a gene or make subtle  codon  changes in the  open reading frame  to identify important amino residues for  protein  function. [ citation needed ]


Alternatively, the technique can be used to create  null alleles  so that the gene is not functional.  For example, deletion of a gene by  gene targeting  ( gene knockout ) can be done in some organisms, such as  yeast ,  mice  and  moss . Unique among plants, in  Physcomitrella patens , gene knockout via  homologous recombination  to create  knockout moss  (see figure) is nearly as efficient as in yeast. [ 4 ]  In the case of the yeast model system directed deletions have been created in every non-essential gene in the yeast genome. [ 5 ]  In the case of the plant  model system  huge mutant libraries have been created based on gene disruption constructs. [ 6 ]  In  gene knock-in , the endogenous exon is replaced by an altered sequence of interest. [ 7 ]


In some cases conditional alleles can be used so that the gene has normal function until the conditional allele is activated.  This might entail 'knocking in'  recombinase  sites (such as lox or frt sites) that will cause a deletion at the gene of interest when a specific recombinase (such as CRE, FLP) is induced.  Cre or Flp recombinases can be induced with chemical treatments, heat shock treatments or be restricted to a specific subset of tissues. [ citation needed ]


Another technique that can be used is  TILLING . This is a method that combines a standard and efficient technique of mutagenesis with a chemical mutagen such as  ethyl methanesulfonate (EMS)  with a sensitive DNA-screening technique that identifies  point mutations  in a target gene. [ citation needed ]


In the field of virology, reverse-genetics techniques can be used to recover full-length infectious viruses with desired mutations or insertions in the viral genomes or in specific virus genes. Technologies that allow these manipulations include circular polymerase extension reaction (CPER) which was first used to generate infectious cDNA for Kunjin virus a close relative of West Nile virus. [ 8 ]  CPER has also been successfully utilised to generate a range of positive-sense RNA viruses such as SARS-CoV-2, [ 9 ]  the causative agent of COVID-19.


The discovery of  gene silencing  using double stranded RNA, also known as  RNA interference  (RNAi), and the development of gene knockdown using  Morpholino  oligos, have made disrupting gene expression an accessible technique for many more investigators. This method is often referred to as a  gene knockdown  since the effects of these reagents are generally temporary, in contrast to  gene knockouts  which are permanent. [ citation needed ]


RNAi creates a specific knockout effect without actually mutating the DNA of interest. In  C. elegans , RNAi has been used to systematically interfere with the expression of most genes in the genome. RNAi acts by directing cellular systems to degrade target messenger RNA (mRNA). [ citation needed ]


RNAi interference, specifically gene silencing, has become a useful tool to silence the expression of genes and identify and analyze their loss-of-function phenotype. When mutations occur in alleles, the function which it represents and encodes also is mutated and lost; this is generally called a loss-of-function mutation. [ 10 ]  The ability to analyze the loss-of-function phenotype allows analysis of gene function when there is no access to mutant alleles. [ 11 ]


While  RNA interference  relies on cellular components for efficacy (e.g. the Dicer proteins, the RISC complex) a simple alternative for gene knockdown is  Morpholino  antisense oligos. Morpholinos bind and block access to the target mRNA without requiring the activity of cellular proteins and without necessarily accelerating mRNA degradation.  Morpholinos are effective in systems ranging in complexity from cell-free translation in a test tube to  in vivo  studies in large animal models. [ citation needed ]


A  molecular genetic  approach is the creation of  transgenic  organisms that  overexpress  a normal gene of interest. The resulting phenotype may reflect the normal function of the gene.


Alternatively it is possible to overexpress mutant forms of a gene that interfere with the normal ( wildtype ) gene's function. For example, over-expression of a mutant gene may result in high levels of a non-functional protein resulting in a  dominant negative  interaction with the wildtype protein.  In this case the mutant version will out compete for the wildtype proteins partners resulting in a mutant phenotype.


Other mutant forms can result in a protein that is abnormally regulated and constitutively active ('on' all the time).  This might be due to removing a regulatory domain or mutating a specific amino residue that is reversibly modified (by  phosphorylation ,  methylation , or  ubiquitination ).  Either change is critical for modulating protein function and often result in informative phenotypes.


Reverse genetics plays a large role in  vaccine  synthesis. Vaccines can be created by engineering novel genotypes of infectious viral strains which diminish their pathogenic potency enough to facilitate immunity in a host. The reverse genetics approach to vaccine synthesis utilizes known viral genetic sequences to create a desired phenotype: a virus with both a weakened pathological potency and a similarity to the current circulating virus strain. Reverse genetics provides a convenient alternative to the traditional method of creating  inactivated vaccines , viruses which have been killed using heat or other chemical methods.


Vaccines created through reverse genetics methods are known as  attenuated vaccines , named because they contain weakened (attenuated) live viruses. Attenuated vaccines are created by combining genes from a novel or current virus strain with previously attenuated viruses of the same species. [ 12 ]  Attenuated viruses are created by propagating a live virus under novel conditions, such as a chicken's egg. This produces a viral strain that is still live, but not pathogenic to humans, [ 13 ]  as these viruses are rendered defective in that they cannot replicate their genome enough to propagate and sufficiently infect a host. However, the viral genes are still expressed in the host's cell through a single replication cycle, allowing for the development of an immunity. [ 14 ]


A common way to create a vaccine using reverse genetic techniques is to utilize plasmids to synthesize attenuated viruses. This technique is most commonly used in the yearly production of  influenza vaccines , where an eight  plasmid  system can rapidly produce an effective vaccine. The entire genome of the  influenza  A virus consists of eight RNA segments, so the combination of six attenuated viral  cDNA  plasmids with two wild-type plasmids allow for an attenuated vaccine strain to be constructed. For the development of influenza vaccines, the fourth and sixth RNA segments, encoding for the  hemagglutinin  and  neuraminidase  proteins respectively, are taken from the circulating virus, while the other six segments are derived from a previously attenuated master strain. The HA and NA proteins exhibit high  antigen  variety, and therefore are taken from the current strain for which the vaccine is being produced to create a well matching vaccine. [ 12 ]


The plasmid used in this eight-plasmid system contains three major components that allow for vaccine development. Firstly, the plasmid contains  restriction sites  that will enable the incorporation of influenza genes into the plasmid. Secondly, the plasmid contains an antibiotic resistance gene, allowing the selection of merely plasmids containing the correct gene. Lastly, the plasmid contains two promotors, human pol 1 and pol 2 promotor that transcribe genes in opposite directions. [ 15 ]


cDNA sequences of viral RNA are synthesized from attenuated master strains by using  RT-PCR . [ 12 ]  This cDNA can then be inserted between an RNA polymerase I (Pol I) promoter and terminator sequence through restriction enzyme digestion. The cDNA and pol I sequence is then, in turn, surrounded by an RNA polymerase II (Pol II) promoter and a  polyadenylation  site. [ 16 ]  This entire sequence is then inserted into a plasmid. Six plasmids derived from attenuated master strain cDNA are cotransfected into a target cell, often a chicken egg, alongside two plasmids of the currently circulating wild-type influenza strain. Inside the target cell, the two "stacked" Pol I and Pol II enzymes transcribe the viral cDNA to synthesize both negative-sense viral RNA and positive-sense mRNA, effectively creating an attenuated virus. [ 12 ]  The result is a defective vaccine strain that is similar to the current virus strain, allowing a host to build immunity. This synthesized vaccine strain can then be used as a seed virus to create further vaccines.


Vaccines engineered from reverse genetics carry several advantages over traditional vaccine designs. Most notable is speed of production. Due to the high  antigenic variation  in the HA and NA  glycoproteins , a reverse-genetic approach allows for the necessary genotype (i.e. one containing HA and NA proteins taken from currently circulating virus strains) to be formulated rapidly. [ 12 ]  Additionally, since the final product of a reverse genetics attenuated vaccine production is a live virus, a higher  immunogenicity  is exhibited than in traditional inactivated vaccines, [ 17 ]  which must be killed using chemical procedures before being transferred as a vaccine. However, due to the live nature of attenuated viruses, complications may arise in  immunodeficient  patients. [ 18 ]  There is also the possibility that a mutation in the virus could result the vaccine to turning back into a live unattenuated virus. [ 19 ]


A  Riboprobe,  abbreviation of RNA probe ,  is a segment of labelled RNA that can be used to detect a target mRNA or DNA during  in situ hybridization . [ 1 ]  RNA  probes  can be produced by  in vitro  transcription of cloned  DNA  inserted in a suitable  plasmid  downstream of a viral promoter. Some bacterial  viruses  code for their own  RNA polymerases , which are highly specific for the viral promoters. Using these  enzymes , labeled  NTPs , and inserts inserted in both forward and reverse orientations, both sense and antisense riboprobes can be generated from a cloned  gene .


Since  James Watson  and  Francis Crick  revealed the  double helix  nature of DNA molecule (Watson & Crick, 1953 [ 2 ] ), the hydrogen bonds between the four bases are well known: adenine always binds to thymine and cytosine always binds to guanine. This binding pattern is the basic principle of modern genetic technologies. Joseph Gall and Mary Lou Pardue published a paper in 1969 demonstrating that radioactive marked ribosomal DNA can be used to detect its complementary DNA sequence in a frog egg, [ 3 ]  known as the first researchers who use DNA probes to perform in situ hybridization. RNA probes were proved to be able to perform the same function and also used with  in situ hybridization . Fluorescent dyed probes replaced radio labeled probes due to the consideration of safety, stability, and ease of detection. [ 4 ]  Detecting a DNA sequence is similar to "looking for a needle in a haystack, with the needle being the DNA sequence of interest and the haystack being a set of chromosomes [ 5 ] ". The ability of the DNA helix to disassociate, re-anneal and the remarkable accuracy of base-pairing grants riboprobes the ability to locate its complementary DNA sequence on chromosomes.


It is found by two independent riboprobes, that the most important  cell  populations of the  hippocampus , are expressing high levels of  diacylglycerol lipase  α (DGL-α), an  enzyme  involved in the production of the  endocannabinoid   2-arachidonoyl-glycerol  (2-AG:  C 23 H 38 O 4 ; 20: 4 ,  ω-6 ). [ 6 ]


There are two kinds of probes used during in situ hybridization: Riboprobes and DNA oligonucleotides probes. [ 7 ]  Riboprobes are essential in the study of embryo development in which DNA probes are insufficient. With labelled (fluoresce-dyed, for instance) antisense RNA probes hybridized with developing embryo's mRNA, tracking the expression of genes in different stages of development is possible. RNA probes can be used in detecting either whole embryo's development or just on tissue sections of interest. The ability of riboprobes to bind to transcribed mRNA makes RNA probes important in research on  model organisms : Drosophila, zebrafish, chick,  Xenopus  and mouse. [ 8 ]  RNA probes can also be utilized in  immunohistochemistry  to identify tissue infection in embryos. [ 9 ]  Viral mRNA can be targeted by its antisense RNA probes, while infected tissues don't have complementary mRNA that can hybridize with probes; the unique mRNA sequence of each organism makes the detection of expression of certain gene highly effective and accurate.


Fluorescence in situ hybridization  (FISH)is the most widely used riboprobe technique. A target sequence and a probe are essential in FISH. First, the probe is labeled with either direct or indirect labeling strategy: hapten-modified nucleotides are used in indirect labeling, and fluorophore-modified nucleotides are used in direct labeling. The target DNA and probes are denatured and mixed, allows the re-annealing of DNA sequences. Indirect labeling requires an extra step to produce visualized signals that require use of enzymatic or immunological system, but provides greater signal amplification than direct labeling. [ 10 ]


FISH probes can also be used in  karyotype  studies. DNA probes can be labeled with various fluorochromes  which produce a unique color for each chromosomes. The probes are then hybridized with metaphase chromosomes, producing unique patterns on each chromosomes. This method is useful when people want to study the translocation, deletion and duplication of chromosomes on a larger scale comparing to site-specific FISH. [ 11 ]


Baynes, John W.; Marek H. Dominiczak (2005).  Medical Biochemistry 2nd. Edition .   Elsevier Mosby . p.  477 .  ISBN   978-0-7234-3341-5 .


YouTube video:in situ hybridization


A detailed description of Fluorescence In Situ Hybridization:


Riboprobe In Vitro Transcription Systems Technical Manual:


An  RNA timestamp  is a technology that enables the age of any given RNA transcript to be inferred by exploiting  RNA editing . [ 1 ]  In this technique, the RNA of interest is tagged to an  adenosine  rich reporter motif that consists of multiple MS2 binding sites. These MS2 binding sites recruit a complex composed of ADAR2 (adenosine deaminase acting on RNA catalytic 2 domain) and MCP (MS2 capsid protein). The binding of the ADAR2 enzyme to the RNA timestamp initiates the gradual conversion of adenosine to  inosine  molecules. Over time, these edits accumulate and are then read through RNA-seq. This technology allows us to glean cell-type specific temporal information associated with RNA-seq data, that until now, has not been accessible. [ 1 ]


The advent of  RNA-sequencing  (RNA-seq) in 2009 allowed for a deeper look into the biology of unique cell types by allowing researchers to examine the presence and quantity of RNA in a sample at any given time. [ 2 ]  The ability for analysis of the  transcriptome has revealed valuable information about cellular differences and transcriptional changes between cell types. Further, RNA-seq has provided insights into examining alternative gene  splicing ,  post-transcriptional modification  and  fusion genes   – all of which would go undetected with  genome analysis  alone. [ 3 ] [ 4 ]  The missing piece of the puzzle is understanding temporally when genes are expressed in a cell. RNA-seq requires the destruction of the cell, thus only revealing the transcriptome at a single moment. [ 1 ]  Understanding expression times and patterns of genes transcription would create a deeper understanding of the roles of genes and how regulation of expression timing could be affecting cellular processes and possible dysregulation could be contributing to disease development.


In recent years, there have been other technologies created with the end goal of determining the age of RNA transcripts within a given cell. For example, TimeLapse-seq, [ 5 ]   SLAM-seq [ 6 ]  as well as measuring RNA velocity which is the instantaneous change in cell state from unspliced transcripts. [ 7 ]  However, these methods were only able to reveal the age of transcripts at a fixed time point and failed to measure transcript age from more than one transcriptional pulse. While this information has importance, the ability to capture the dynamic transcriptional changes within a cell remained elusive.


The research groups of Edward Boyden at  MIT  and Fei Chen at  Harvard , both in Cambridge, Massachusetts, U.S.A, developed RNA timestamps, a new (2020) method that allows the age chosen transcripts in a cell to be inferred. [ 1 ]  This technology provides the means to understand cellular biology at a new level and deepen our understanding of cellular processes  and transcriptional regulation. Further, the use of RNA timestamps does not require lysis of the cell which is an undeniable advantage. This allows timestamps to have the unique availability to suggest information from multiple transcriptional pulses throughout the cellular development rather than being limited to a fixed time point. [ 1 ]


Since multiple timestamped RNA transcripts are produced in the cells by a specific promoter, a transcriptional program was developed that uses a gradient descent algorithm to describe the number of timestamps generated as a function of time. [ 1 ]  This helps with determining the source of the time point for the RNA timestamps and the accuracy of the algorithm increased with the number of RNAs. Further, this reveals the time point at which the specific promoter was active. [ 1 ]


RNA timestamp experiments were first experimentally validated in HEK293T cells that expressed the ADAR variant along with time stamped RNA under the control of tetracycline response element (TRE) induced by doxycycline. From these experiments, it was shown that the age of multiple RNA transcripts can be accurately determined (with a 95% confidence interval of 2.7±0.4h). [ 1 ]


As part of their proof of concept experiments, the researchers showed that timestamps can be used in primary  hippocampal   neuron  cell culture to infer the c-fos response from KCl activation of neural activity. [ 1 ]


This technology was also shown to have the potential to determine transcriptional events in individual cells. Timestamps can be read out when used in combination with high throughput  single-cell  droplet based methods thus allowing RNA timestamps to be used for applications like ordering of mRNAs based on the timing of transcriptional processes based on cell type, and determining whether a specific promoter was active. [ 1 ]


RNA timestamp technology provides a novel means to understand cellular transcription. Only a few months after this technology was described, Dr. Michael Gilhooey and colleagues in England discussed RNA timestamps as an interesting new perspective to apply to their research on inherited optic neuropathies. [ 12 ]  This suggests diverse potential applications of RNA timestamps, and perhaps it will be beneficial to better understanding transcriptional changes in human diseases, as alluded to by Dr. Gilhooey.


Further, the researchers suggested that perhaps timestamps could be calibrated to be used in  in vivo  experiments. The researchers have also suggested a potential mechanism using a virus to deliver the RNA reporter to target cells   in vivo . [ 13 ]  This could provide a revolutionary mechanism to track  expression dynamics during development or recording responses to stimuli  in vivo . However, for now, timestamps are limited to  in vitro  experiments.


Despite the many advantages of this method, there are some caveats of RNA timestamping worth mentioning:


Self-complementary adeno-associated virus  ( scAAV ) is a  viral vector  engineered from the naturally occurring  adeno-associated virus  (AAV) to be used as a tool for  gene therapy . [ 1 ]  Use of  recombinant  AAV (rAAV) has been successful in  clinical trials  addressing a variety of diseases. [ 2 ]  This lab-made progeny of rAAV is termed "self-complementary" because the  coding region  has been designed to form an intra-molecular double-stranded DNA template. A  rate-limiting step  for the standard AAV genome involves the second-strand synthesis since the typical AAV  genome  is a single-stranded  DNA  template. [ 3 ] [ 4 ]  However, this is not the case for scAAV genomes. Upon infection, rather than waiting for cell mediated  synthesis  of the second strand, the two complementary halves of scAAV will associate to form one double stranded DNA (dsDNA) unit that is ready for immediate  replication  and  transcription . The caveat of this construct is that instead of the full coding capacity found in rAAV (4.7–6 kb ) [ 5 ]  scAAV can only hold about half of that amount (≈2.4kb). [ 6 ]


In gene therapy application utilizing rAAV, the virus  transduces  the cell with a single stranded DNA (ssDNA) flanked by two  inverted terminal repeats  (ITRs). These ITRs form hairpins at the end of the sequence to serve as  primers  to initiate synthesis of the second strand before subsequent steps of  infection  can begin. The second strand synthesis is considered to be one of several blocks to efficient infection. [ 7 ]  Additional advantages of scAAV include increased and prolonged  transgene   expression   in vitro  and  in vivo , as well as "higher in vivo DNA stability and more effective circularization." [ 8 ]


scAAV is an attractive vector for use in  gene therapy  for many reasons. Its parent vector, AAV, is already being used in clinical trials. [ 9 ]  Due to a variety of scAAV  serotypes  available, scientists can choose a serotype which has properties desirable for their therapy. Selecting only a subset of cells improves specificity and lowers the risk of being inhibited by the  immune system . Different scAAV and AAV serotypes can efficiently transfect a variety of cellular targets. [ 10 ] [ 11 ]  Like all vector-based approaches to gene therapy, one obstacle in translating therapies from pre-clinical trials into a human  clinical  application will be the production of large quantities of highly concentrated virus. [ 12 ]  One disadvantage that scAAV faces is that due to robust gene expression, transgene products delivered via scAAV elicit a stronger immune response than those same transgenes delivered via a single-stranded AAV vector. [ 13 ]


Like AAV, scAAV is a member of the family  Parvoviridae , commonly known as  parvoviruses . These viruses are  nonenveloped , single-strand DNA (ssDNA) viruses. Within Parvoviridae, scAAV further belongs to the  Dependovirus  genus, characteristically defined by an inability to replicate on their own. In nature, these viruses depend on another virus to provide replication machinery; adeno-associated virus can only replicate during an active infection of  adenovirus  or some types of  herpesvirus . In lab use, this obstacle is overcome by addition of the helper  plasmids , which  exogenously  expresses replication genes which AAV itself lacks. [ 14 ]


As a dependovirus, scAAV remains in a  latent  state within the cell until the cell experiences certain permissive conditions. These can include presence of a  helper virus  infection (such as adenovirus) or other toxic events such as exposure to  UV  light or  carcinogens . [ 14 ]  Because the endogenous rep  ORF  has been replaced with transgene, exogenously provided rep genes encode the proteins required for genome replication and other viral life cycle components. The ITRs located 5' and 3' of the viral genome serve as the origin of replication. [ 15 ]


Like the rep ORF, scAAV's cap ORF has been replaced by transgene and therefore is provided exogenously in a lab environment. The genes encoded in this ORF build capsid proteins and are responsible (along with intracellular processing) for conveying target specificity. Rep proteins participate in the integration of the genome into preformed capsids. [ 15 ]  Despite the fact that scAAV is designed to form dsDNA upon infection, the two complementary strands are not packaged in a double stranded manner. Parvoviruses package their viral genome such that the ssDNA bases come in contact with the amino acids on the inside of the viral capsid. Thus the sequence of scAAV is likely unwound by a virally encoded DNA helicase before being packaged into viral protein capsid. [ 7 ]


Site-specific recombinase technologies  are  genome engineering  tools that depend on  recombinase enzymes  to replace targeted sections of DNA.


In the late 1980s gene targeting in murine  embryonic stem cells  (ESCs) enabled the transmission of mutations into the mouse germ line, and emerged as a novel option to study the genetic basis of regulatory networks as they exist in the genome. Still, classical  gene targeting  proved to be limited in several ways as gene functions became irreversibly destroyed by the marker gene that had to be introduced for selecting recombinant ESCs. These early steps led to animals in which the mutation was present in all cells of the body from the beginning leading to complex phenotypes and/or early lethality. There was a clear need for methods to restrict these mutations to specific points in development and specific cell types. This dream became reality when groups in the USA were able to introduce bacteriophage and yeast-derived site-specific recombination (SSR-) systems into mammalian cells as well as into the mouse. [ 1 ] [ 2 ] [ 3 ]


Common genetic engineering strategies require a permanent modification of the target genome. To this end great sophistication has to be invested in the design of routes applied for the delivery of transgenes. Although for biotechnological purposes random integration is still common, it may result in unpredictable gene expression due to variable transgene copy numbers, lack of control about integration sites and associated mutations.  The molecular requirements in the stem cell field are much more stringent. Here,  homologous recombination  (HR) can, in principle, provide specificity to the integration process, but for eukaryotes it is compromised by an extremely low efficiency. Although meganucleases, zinc-finger- and transcription activator-like effector nucleases (ZFNs and TALENs) are actual tools supporting HR, it was the availability of site-specific recombinases (SSRs) which triggered the rational construction of cell lines with predictable properties. Nowadays both technologies, HR and SSR can be combined in highly efficient "tag-and-exchange technologies". [ 4 ]


Many  site-specific recombination  systems have been identified to perform these DNA rearrangements for a variety of purposes, but nearly all of these belong to either of two families, tyrosine recombinases (YR) and serine recombinases (SR), depending on their  mechanism . These two families can mediate up to three types of DNA rearrangements (integration, excision/resolution, and inversion) along different reaction routes based on their origin and architecture. [ 5 ]


The founding member of the YR family is the  lambda integrase , encoded by   bacteriophage λ , enabling the integration of phage DNA into the bacterial genome. A common feature of this class is a conserved tyrosine nucleophile attacking the scissile DNA-phosphate to form a 3'-phosphotyrosine linkage. Early members of the SR family are closely related  resolvase  /  DNA invertases  from the bacterial transposons Tn3 and γδ, which rely on a catalytic serine responsible for attacking the scissile phosphate to form a 5'-phosphoserine linkage. These undisputed facts, however, were compromised by a good deal of confusion at the time other members entered the scene, for instance the YR recombinases  Cre  and  Flp  (capable of integration, excision/resolution as well as inversion), which were nevertheless welcomed as new members of the "integrase family". The converse examples are PhiC31 and related SRs, which were originally introduced as resolvase/invertases although, in the absence of auxiliary factors, integration is their only function. Nowadays the standard activity of each enzyme determines its classification reserving the general term "recombinase" for family members which, per se, comprise all three routes, INT, RES and INV:


Our table extends the selection of the conventional SSR systems and groups these according to their performance. All of these enzymes recombine two target sites, which are either identical (subfamily A1) or distinct (phage-derived enzymes in  A2, B1 and B2). [ 6 ]  Whereas for A1 these sites have individual designations (" FRT " in case of Flp-recombinase,  loxP  for Cre-recombinase), the terms " att P" and " att B" (attachment sites on the phage and bacterial part, respectively) are valid in the other cases. In case of subfamily A1 we have to deal with short (usually 34 bp-) sites consisting of two (near-)identical 13 bp arms (arrows) flanking an 8 bp spacer (the crossover region, indicated by red line doublets). [ 7 ]  Note that for Flp there is an alternative, 48 bp site available with three arms, each accommodating a Flp unit (a so-called "protomer").  att P- and  att B-sites follow similar architectural rules, but here the arms show only partial identity (indicated by the broken lines) and differ in both cases. These features account for relevant differences:


In order  to streamline this chapter the following implementations will be focused on two recombinases (Flp and Cre) and just one integrase (PhiC31) since their spectrum covers the tools which, at present, are mostly used for directed genome modifications. This will be done in the framework of the following overview.


The mode integration/resolution and inversion (INT/RES and INV) depend on the orientation of recombinase target sites (RTS), among these pairs of  att P and  att B. Section C indicates, in a streamlined fashion, the way  recombinase-mediated cassette exchange  (RMCE) can be reached by synchronous double-reciprocal crossovers (rather than integration, followed by resolution). [ 8 ] [ 9 ]


Tyr-Recombinases are reversible, while the Ser-Integrase is unidirectional. Of note is the way reversible Flp (a Tyr recombinase) integration/resolution is modulated by 48 bp (in place of 34 bp minimal)  FRT  versions: the extra 13 bp arm serves as a Flp "landing path" contributing to the formation of the synaptic complex, both in the context of Flp-INT and Flp-RMCE functions (see the respective equilibrium situations). While it is barely possible to prevent the (entropy-driven) reversion of integration in section A for Cre and hard to achieve for Flp, RMCE can be completed if the donor plasmid is provided at an excess due to the bimolecular character of both the forward-  and the reverse reaction. Posing both  FRT  sites in an inverse manner will lead to an equilibrium of both orientations for the insert (green arrow). In contrast to Flp, the Ser integrase PhiC31 (bottom representations) leads to unidirectional integration, at least in the absence of an recombinase-directionality (RDF-)factor. [ 10 ]   Relative to Flp-RMCE, which requires two different ("heterospecific")  FRT -spacer mutants, the reaction partner ( att B) of the first reacting  att P site is hit arbitrarily, such that there is no control over the direction the donor cassette enters the target (cf. the alternative products). Also different from  Flp-RMCE , several distinct RMCE targets cannot be mounted in parallel, owing to the lack of heterospecific (non-crossinteracting)  att P/ att B combinations.


Cre recombinase  (Cre) is able to recombine specific sequences of DNA without the need for  cofactors . The enzyme recognizes 34 base pair DNA sequences called   loxP  ("locus of crossover in phage P1"). Depending on the orientation of target sites with respect to one another, Cre will integrate/excise or invert DNA sequences. Upon the excision (called "resolution" in case of a circular substrate) of a particular DNA region, normal gene expression is considerably compromised or terminated. [ 11 ]


Due to the pronounced resolution activity of Cre, one of its initial applications was the excision of  lox P-flanked ("floxed") genes leading to cell-specific gene knockout of such a floxed gene after Cre becomes expressed in the tissue of interest. Current technologies incorporate methods, which allow for both the spatial and temporal control of Cre activity.  A common method facilitating the spatial control of genetic alteration involves the selection of a tissue-specific  promoter  to drive Cre expression. Placement of Cre under control of such a promoter results in localized, tissue-specific expression. As an example, Leone et al. have placed the transcription unit under the control of the regulatory sequences of the  myelin  proteolipid protein (PLP) gene, leading to induced removal of targeted gene sequences in  oligodendrocytes  and  Schwann cells . [ 12 ]  The specific DNA fragment recognized by Cre remains intact in cells, which do not express the PLP gene; this in turn facilitates empirical observation of the localized effects of genome alterations in the myelin sheath that surround nerve fibers in the  central nervous system  (CNS) and the  peripheral nervous system  (PNS). [ 13 ]  Selective Cre expression has been achieved in many other cell types and tissues as well.


In order to control temporal activity of the excision reaction, forms of Cre which take advantage of various  ligand  binding domains have been developed. One successful strategy for inducing specific temporal Cre activity involves fusing the enzyme with a mutated ligand-binding domain for the human  estrogen receptor  (ERt).  Upon the introduction of  tamoxifen  (an estrogen  receptor antagonist ), the Cre-ERt construct is able to penetrate the nucleus and induce targeted mutation. ERt binds tamoxifen with greater affinity than  endogenous   estrogens , which allows Cre-ERt to remain  cytoplasmic  in animals untreated with tamoxifen. The temporal control of SSR activity by tamoxifen permits genetic changes to be induced later in  embryogenesis  and/or in adult tissues. [ 12 ]  This allows researchers to bypass embryonic lethality while still investigating the function of targeted genes.


Recent extensions of these general concepts led to generating the "Cre-zoo", i.e. collections of hundreds of mouse strains for which defined genes can be deleted by targeted Cre expression. [ 3 ]


In its natural host (S. cerevisiae) the  Flp/ FRT  system enables replication of a "2μ plasmid" by the inversion of a segment that is flanked by two identical, but oppositely oriented  FRT  sites ("flippase" activity). This inversion changes the relative orientation of replication forks within the plasmid enabling "rolling circle"—amplification of the circular 2μ entity before the multimeric intermediates are resolved to release multiple monomeric products. Whereas 34 bp minimal  FRT  sites favor excision/resolution to a similar extent as the analogue  lox P sites for Cre, the natural, more extended 48 bp FRT variants enable a higher degree of integration, while overcoming certain promiscuous interactions as described for phage enzymes like Cre-  [ 5 ]  and PhiC31. [ 6 ]  An additional advantage is the fact, that simple rules can be applied to generate heterospecific  FRT  sites which undergo crossovers with equal partners but nor with wild type  FRT s. These facts have enabled, since 1994, the development and continuous refinements of  recombinase-mediated cassette exchange  (RMCE-)strategies permitting the clean exchange of a target cassette for an incoming donor cassette. [ 6 ]


Based on the RMCE technology, a particular resource of pre-characterized ES-strains that lends itself to further elaboration has evolved in the framework of the EUCOMM (European Conditional Mouse Mutagenesis) program, based on the now established Cre- and/or Flp-based "FlExing" (Flp-mediated excision/inversion) setups, [ 6 ]  involving the excision and inversion activities. Initiated in 2005, this project focused first on saturation mutagenesis to enable complete functional annotation of the mouse genome (coordinated by the International Knockout-Mouse Consortium, IKMC) with the ultimate goal to have all protein genes mutated via gene trapping and -targeting in murine ES cells. [ 14 ]  These efforts mark the top of various "tag-and-exchange" strategies, which are dedicated to tagging a distinct genomic site such that the "tag" can serve as an address to introduce novel (or alter existing) genetic information. The tagging step  per se  may address certain classes of integration sites by exploiting integration preferences of retroviruses or even site specific integrases like PhiC31, both of which act in an essentially unidirectional fashion.


The traditional, laborious "tag-and-exchange" procedures relied on two successive homologous recombination (HR-)steps, the first one ("HR1") to introduce a tag consisting of a selection marker gene. "HR2" was then used to replace the marker by the "GOI. In the first ("knock-out"-) reaction the gene was tagged with a selectable marker, typically by insertion of a hygtk ([+/-]) cassette providing G418 resistance. In the following "knock-in" step, the tagged genomic sequence was replaced by homologous genomic sequences with certain mutations. Cell clones could then be isolated by their resistance to ganciclovir due to loss of the HSV-tk gene, i.e. ("negative selection"). This conventional two-step tag-and-exchange procedure  [ 15 ]   could be streamlined after the advent of RMCE, which could take over and add efficiency to the knock-in step.


Without much doubt, Ser  integrases  are the current tools of choice for integrating transgenes into a restricted number of well-understood genomic acceptor sites that mostly (but not always) mimic the phage  att P site in that they attract an  att B-containing donor vector. At this time the most prominent member is PhiC31-INT with proven potential in the context of human and mouse genomes.


Contrary to the above Tyr recombinases, PhiC31-INT as such acts in a unidirectional manner, firmly locking in the donor vector at a genomically anchored target. An obvious advantage of this system is that it can rely on unmodified, native  att P (acceptor) and  att B donor sites. Additional benefits (together with certain complications) may arise from the fact that mouse and human genomes per se contain a limited number of endogenous targets (so called " att P-pseudosites"). Available information suggests that considerable DNA sequence requirements let the integrase recognize fewer sites than retroviral or even transposase-based integration systems opening its career as a superior carrier vehicle for the transport and insertion at a number of well established genomic sites, some of which with so called "safe-harbor" properties. [ 10 ]


Exploiting the fact of specific ( att P x  att B) recombination routes, RMCE becomes possible without requirements for synthetic, heterospecific  att -sites. This obvious advantage, however comes at the expense of certain shortcomings, such as lack of control about the kind or directionality of the entering (donor-) cassette. [ 6 ]  Further restrictions are imposed by the fact that irreversibility does not permit standard  multiplexing-RMCE  setups including "serial RMCE" reactions, i.e.,  repeated cassette exchanges at a given genomic  locus .


Annotation of the human and mouse genomes has led to the identification of >20 000 protein-coding  genes and  >3 000 noncoding RNA genes, which guide the development of the organism from fertilization through embryogenesis to adult life. Although dramatic progress is noted, the relevance of rare gene variants has remained a central topic of research.


As one of the most important platforms for dealing with vertebrate gene functions on a large scale, genome-wide genetic resources of mutant murine ES cells have been established. To this end four international programs aimed at saturation mutagenesis of the mouse genome have been founded in Europe and North America (EUCOMM, KOMP, NorCOMM, and TIGM). Coordinated by  the International Knockout Mouse Consortium (IKSC) these  ES-cell repositories are available for exchange between international research units. Present resources comprise mutations in 11 539 unique genes, 4 414 of these conditional. [ 14 ]


The relevant technologies have now reached a level permitting their extension to other mammalian species and to human stem cells, most prominently those with an  iPS (induced pluripotent)  status.


Somatic fusion , also called  protoplast fusion , is a type of  genetic modification  in plants by which two distinct species of plants are fused together to form a new  hybrid  plant with the characteristics of both, a  somatic hybrid . [ 1 ]  Hybrids have been produced either between different varieties of the same species (e.g. between non-flowering  potato  plants and flowering potato plants) or between two different species (e.g. between wheat  Triticum  and rye  Secale  to produce  Triticale ).


Uses of somatic fusion include developing plants resistant to disease, such as making potato plants resistant to  potato leaf roll disease . [ 2 ]  Through somatic fusion, the crop potato plant  Solanum tuberosum  – the yield of which is severely reduced by a viral disease transmitted on by the aphid  vector  – is fused with the wild, non-tuber-bearing potato  Solanum brevidens , which is resistant to the disease. The resulting hybrid has the chromosomes of both plants and is thus similar to  polyploid  plants.
Somatic hybridization was first introduced by Carlson et al. in  Nicotiana glauca . [ 3 ]


The somatic fusion process occurs in four steps: [ 4 ]


The procedure for seed plants describe above, fusion of  moss  protoplasts can be initiated without electric shock but by the use of  polyethylene glycol  (PEG). Further, moss protoplasts do not need  phytohormones  for  regeneration , and they do not form a  callus . [ 5 ]  Instead, regenerating moss protoplasts behave like  germinating  moss  spores . [ 6 ]  Of further note  sodium nitrate  and calcium ion at high pH can be used, although results are variable depending on the organism. [ 7 ]


Somatic cells  of different types can be fused to obtain hybrid cells. Hybrid cells are useful in a variety of ways, e.g.,


Chromosome  mapping through somatic cell hybridization is essentially based on fusion of human and  mouse  somatic cells. Generally, human  fibrocytes  or  leucocytes  are fused with mouse continuous  cell lines .


When human and mouse cells (or cells of any two  mammalian  species or of the same species) are mixed, spontaneous  cell fusion  occurs at a very low rate (10-6). Cell fusion is enhanced 100 to 1000 times by the addition of  ultraviolet  inactivated  Sendai  (parainfluenza) virus or  polyethylene glycol  (PEG).


These agents adhere to the  plasma membranes  of cells and alter their properties in such a way that facilitates their fusion. Fusion of two cells produces a heterokaryon, i.e., a single hybrid cell with two nuclei, one from each of the cells entering fusion. Subsequently, the two nuclei also fuse to yield a hybrid cell with a single nucleus.


A generalized scheme for somatic cell hybridization may be described as follows. Appropriate human and mouse cells are selected and mixed together in the presence of inactivated Sendai virus or PEG to promote cell fusion. After a period of time, the cells (a mixture of man, mouse and 'hybrid' cells) are plated on a  selective medium , e.g.,  HAT medium , which allows the  multiplication  of hybrid cells only.


Several  clones  (each derived from a single hybrid cell) of the hybrid cells are thus isolated and subjected to both  cytogenetic  and appropriate  biochemical analyses  for the detection of  enzyme /  protein / trait  under investigation. An attempt is now made to correlate the presence and absence of the trait with the presence and absence of a  human chromosome  in the hybrid clones.


If there is a perfect  correlation  between the presence and absence of a human chromosome and that of a trait in the hybrid clones, the gene governing the trait is taken to be located in the concerned chromosome.


The HAT medium is one of the several selective media used for the selection of hybrid cells. This medium is supplemented with  hypoxanthine ,  aminopterin  and  thymidine , hence the name HAT medium.  Antimetabolite  aminopterin blocks the cellular  biosynthesis  of  purines  and  pyrimidines  from  simple sugars  and  amino acids .
However, normal human and mouse cells can still multiply as they can utilize hypoxanthine and thymidine present in the medium through a  salvage pathway , which ordinarily recycles the purines and pyrimidines produced from degradation of  nucleic acids .
Hypoxanthine is converted into  guanine  by the enzyme  hypoxanthine-guanine phosphoribosyltransferase  (HGPRT), while thymidine is  phosphorylated  by  thymidine kinase  (TK); both HGPRT and TK are enzymes of the salvage pathway.
On a HAT medium, only those cells that have active HGPRT (HGPRT+) and TK (TK+) enzymes can proliferate, while those deficient in these enzymes (HGPRr- and/or TK-) can not divide (since they cannot produce purines and pyrimidines due to the aminopterin present in the HAT medium).
For using HAT medium as a selective agent,  human cells  used for fusion must be deficient for either the enzyme HGPRT or TK, while mouse cells must be deficient for the other enzyme of this pair. Thus, one may fuse HGPRT deficient human cells (designated as TK+ HGPRr-) with TK deficient mouse cells (denoted as TK- HGPRT+).
Their fusion products (hybrid cells) will be TK+ (due to the human  gene ) and HGPRT+ (due to the mouse gene) and will multiply on the HAT medium, while the man and mouse cells will fail to do so. Experiments with other selective media can be planned in a similar fashion.


Note: The table only lists a few examples, there are many more crosses. The possibilities of this technology are great; however, not all species are easily put into protoplast culture.


Sperm-mediated gene transfer  (SMGT) is a transgenic technique that transfers  genes  based on the ability of  sperm cells  to spontaneously bind to and internalize exogenous  DNA  and transport it into an oocyte during fertilization to produce  genetically modified animals . 1  Exogenous DNA refers to DNA that originates outside of the organism. Transgenic animals have been obtained using SMGT, but the efficiency of this technique is low. Low efficiency is mainly due to low uptake of exogenous DNA by the spermatozoa, reducing the chances of fertilizing the oocytes with transfected spermatozoa. 2  In order to successfully produce transgenic animals by SMGT, the spermatozoa must attach the exogenous DNA into the head and these transfected spermatozoa must maintain their functionality to fertilize the oocyte. 2  Genetically modified animals produced by SMGT are useful for research in biomedical, agricultural, and veterinary fields of study. SMGT could also be useful in generating animals as models for human diseases or lead to future discoveries relating to human gene therapy.


The method for SMGT uses the sperm cell, a natural vector of genetic material, to transport exogenous DNA. The exogenous DNA molecules bind to the cell membrane of the head of the sperm cell. This binding and internalization of the DNA is not a random event. The exogenous DNA interacts with the DNA-binding proteins (DBPs) that are present on the surface of the sperm cell. 3   Spermatozoa  are naturally protected against the intrusion of exogenous DNA molecules by an inhibitory factor present in mammals’ seminal fluid. This factor blocks the binding of sperm cells and exogenous DNA because in the presence of the inhibitory factor, DBPs lose their ability to bind to exogenous DNA. In the absence of this inhibitory factor, DBPs on sperm cells are able to interact with DNA and can then translocate the DNA into the cell. Therefore, the seminal fluid must be removed from the sperm samples by extensive washing immediately after  ejaculation . 3  After the DNA is internalized, the exogenous DNA must be integrated into the genome. There are various mechanisms suggested for DNA integration, including integrating DNA at oocyte activation, at nucleus decondensation, or at the formation of the pronuclei, but all of these suggested mechanisms imply that the integration of DNA happens after the penetration of the sperm cell into the oocyte. 3


Sperm-mediated gene transfer is considered controversial because despite the successes, it has not yet become established as a reliable form of genetic manipulation. Skepticism arises based on the assumption that evolutionary chaos could arise if sperm cells could act as vectors for exogenous DNA. 4  Reasonable assumption tells us that because reproductive tracts contain free DNA molecules, sperm cells should be highly resistant to the risk of picking up exogenous DNA molecules. SMGT has been demonstrated experimentally and followed the assumption that nature has barriers against SMGT. These barriers are not always absolute and could explain the inconsistent experimental outcomes of SMGT. 4  If there are natural barriers against SMGT, then the successes may only represent unusual cases in which the barriers failed. Two barriers have been identified; the inhibitory factor in seminal fluid that prevents binding to foreign DNA molecules and a sperm endogenous nuclease activity that is triggered upon interaction of sperm cells with foreign DNA molecules. 4  These protections give reason to believe that unintentional interactions between sperm and exogenous genetic sequences is kept to a minimal. These barriers allow for protection against the threat that every fertilization event could become a potentially mutagenic one. 4


Transgenic animals have been produced successfully using gene transfer techniques such as sperm-mediated gene transfer. Though this production has been successful, the efficiency of the process is low. Low efficiency of SMGT in the production of transgenic animals is mainly due to poor uptake of the exogenous DNA by the sperm cells, thus reducing the number of fertilized oocytes with transfected spermatozoa. 5  From 1989 to 2004, there were over 30 claims for the production of viable transgenic animals using SMGT, but only about 25 percent of these demonstrated a transmission of the transgenes beyond the F 0  generation. 4  This transmission is required in order to claim usable animal transgenesis. According to previous studies, numerous animal species, including mammals, birds, insects, and fish, have been found susceptible to SMGT techniques, thus indicating that SMGT has broad applicability across a wide variety of Metazoan species. 4  Currently, despite the low frequency of transmission of transgenes, the frequency of phenotype modifications and overall animal transgenesis has been as high as 80 percent in some experiments. 4


The potential use of sperm-mediated gene transfer for embryo somatic gene therapy is a possibility for future research. Embryo somatic gene therapy would be advantageous because there seems to be an inverse correlation between the age of the patient and the effectiveness of gene therapy. Therefore, the possibility of gene therapy treatment before irreversible damage occurs would be ideal. 4  A majority of the experiments that report successful SMGT provide evidence of post-fertilization transfer and maintenance of transgenes. 6  SMGT has potential advantages of being a simple and cost-effective method of gene therapy, especially in contrast with pronuclear microinjection, another transgenic technique. Nevertheless, despite some successes and its potential utility, SMGT is not yet established as a reliable form of genetic modification. 6


1. Lavitrano M, Giovannoni R, Cerrito MG. 2013. Methods for sperm-mediated gene transfer. Methods Molecular Biology. 927:519-529.


2. García-Vázquez FA, Ruiz S, Grullón LA, Ondiz AD, Gutiérrez-Adán A, Gadea J. 2011. Factors affecting porcine sperm mediated gene transfer. Research in Veterinary Science. 91(3):446-53.


3. Lavitrano M, Busnelli M, Cerrito MG, Giovannoni R, Manzini S, Vargiolu A. 2006. Sperm-mediated gene transfer. Reproduction, Fertility and Development. 18:19-23.


4. Smith K, Spadafora C. 2005. Sperm-mediated gene transfer: applications and implications. BioEssays. 27(5):551-562.


5. Collares T, Campos VF, de Leon PM, Moura, Cavalcanti PV, Amaral, MG, et al. 2011. Transgene transmission in chickens by sperm-mediated gene transfer after seminal plasma removal and exogenous DNA treated with dimethylsulfoxide or N,N-dimethylacetamide. Journal of Biosciences. 36(4):613-620.


6. Smith K. 2004. Gene therapy: the potential applicability of gene transfer technology to the human germline. International Journal of Medical Sciences. 1(2):76-91.


Synthetic genome is a synthetically built genome whose formation involves either  genetic modification  on pre-existing life forms or  artificial gene synthesis  to create new DNA or entire lifeforms. [ 1 ] [ 2 ] [ 3 ]  The field that studies  synthetic genomes  is called  synthetic genomics .


Soon after the discovery of  restriction endonucleases  and  ligases , the field of genetics began using these molecular tools to assemble artificial sequences from smaller fragments of synthetic or naturally occurring  DNA . The advantage in using the recombinatory approach as opposed to continual  DNA synthesis  stems from the inverse relationship that exists between synthetic DNA length and percent purity of that synthetic length. In other words, as you synthesize longer sequences, the number of error-containing clones increases due to the inherent error rates of current technologies. [ 4 ]  Although  recombinant DNA  technology is more commonly used in the construction of  fusion proteins  and  plasmids , several techniques with larger capacities have emerged, allowing for the construction of entire genomes. [ 5 ]


Polymerase cycling assembly  (PCA) uses a series of oligonucleotides (or oligos), approximately 40 to 60 nucleotides long, that altogether constitute both strands of the DNA being synthesized. These oligos are designed such that a single oligo from one strand contains a length of approximately 20 nucleotides at each end that is complementary to sequences of two different oligos on the opposite strand, thereby creating regions of overlap. The entire set is processed through cycles of: (a)  hybridization  at 60 °C; (b) elongation via  Taq polymerase  and a standard ligase; and (c) denaturation at 95 °C, forming progressively longer contiguous strands and ultimately resulting in the final genome. [ 6 ]  PCA was used to generate the first synthetic genome in history, that of the  Phi X 174 virus . [ 7 ]


The  gibson assembly  method, designed by Daniel Gibson during his time at the  J. Craig Venter Institute , requires a set of double-stranded DNA cassettes that constitute the entire genome being synthesized. Note that cassettes differ from contigs by definition, in that these sequences contain regions of homology to other cassettes for the purposes of  recombination . In contrast to Polymerase Cycling Assembly, Gibson Assembly is a single-step, isothermal reaction with larger sequence-length capacity; ergo, it is used in place of Polymerase Cycling Assembly for genomes larger than 6 kb.


A T5  exonuclease  performs a chew-back reaction at the terminal segments, working in the 5' to 3' direction, thereby producing complementary overhangs. The overhangs hybridize to each other, a Phusion  DNA polymerase  fills in any missing nucleotides and the nicks are sealed with a ligase. However, the genomes capable of being synthesized using this method alone is limited because as DNA cassettes increase in length, they require propagation in vitro in order to continue hybridizing; accordingly, Gibson assembly is often used in conjunction with Transformation-Associated Recombination (see below) to synthesize genomes several hundred kilobases in size. [ 8 ]


The goal of transformation-associated recombination (TAR) technology in synthetic genomics is to combine DNA contigs by means of  homologous recombination  performed by the  Yeast Artificial Chromosome  (YAC). Of importance is the CEN element within the YAC  vector , which corresponds to the yeast centromere. This sequence gives the vector the ability to behave in a chromosomal manner, thereby allowing it to perform  homologous recombination . [ 9 ]


First, gap repair cloning is performed to generate regions of homology flanking the DNA contigs. Gap Repair Cloning is a particular form of the  Polymerase Chain Reaction  in which specialized  primers  with extensions beyond the sequence of the DNA target are utilized. [ 10 ]  Then, the DNA cassettes are exposed to the YAC vector, which drives the process of homologous recombination, thereby connecting the DNA cassettes. Polymerase Cycling Assembly and TAR technology were used together to construct the 600 kb  Mycoplasma genitalium  genome in 2008, the first synthetic organism ever created. [ 11 ]  Similar steps were taken in synthesizing the larger  Mycoplasma mycoides  genome a few years later. [ 12 ]


It is difficult to directly synthesize oligonucleotides larger than ~200 base pairs and maintain high fidelity. [ 13 ]  Therefore, smaller oligonucleotides (around 5-20 base pairs) are combined to create genome-size oligonucleotides. Previous methods of stitching the smaller strands involved using  T4 polynucleotide ligase . Modern techniques, like PCA/ PCR  based-methods have improved on this method, increasing speed and fidelity. To further increase fidelity, PCA-based methods can include an error-reversal step in which  nucleases  recognize and cut mismatched base pairs. [ 14 ]  Recognition is possible because errors usually cause structural budges and abnormalities in the DNA. [ 15 ]   Currently, a 4-Mb  E. coli  genome created in May 2019 holds the record for the largest synthetic genome size. [ 16 ]


Synthetic genomics  is a nascent field of  synthetic biology  that uses aspects of  genetic modification  on pre-existing life forms, or  artificial gene synthesis  to create new DNA or entire lifeforms.


Synthetic genomics is unlike  genetic modification  in the sense that it does not use naturally occurring genes in its life forms. It may make use of  custom designed base pair series , though in a more expanded and presently unrealized sense synthetic genomics could utilize genetic codes that are not composed of the  two base pairs  of  DNA  that are currently used by life.


The development of synthetic genomics is related to certain recent technical abilities and technologies in the field of genetics. The ability to construct long  base pair  chains cheaply and accurately on a large scale has allowed researchers to perform experiments on genomes that do not exist in nature. Coupled with the developments in  protein folding  models and decreasing computational costs the field of synthetic genomics is beginning to enter a productive stage of vitality.


Researchers were able to create a synthetic organism for the first time in 2010. [ 1 ]  This breakthrough was undertaken by  Synthetic Genomics, Inc. , which continues to specialize in the research and commercialization of custom designed genomes. [ 2 ]  It was accomplished by synthesizing a 600 kbp genome (resembling that of  Mycoplasma genitalium , save the insertion of a few watermarks) via the  Gibson Assembly  method and Transformation Associated Recombination. [ 3 ]


Soon after the discovery of  restriction endonucleases  and  ligases , the field of genetics began using these molecular tools to assemble artificial sequences from smaller fragments of synthetic or naturally-occurring DNA. The advantage in using the recombinatory approach as opposed to continual DNA synthesis stems from the inverse relationship that exists between synthetic DNA length and percent purity of that synthetic length. In other words, as you synthesize longer sequences, the number of error-containing clones increases due to the inherent error rates of current technologies. [ 4 ]  Although recombinant DNA technology is more commonly used in the construction of  fusion proteins  and  plasmids , several techniques with larger capacities have emerged, allowing for the construction of entire genomes. [ 5 ]


Polymerase cycling assembly  (PCA) uses a series of oligonucleotides (or oligos), approximately 40 to 60 nucleotides long, that altogether constitute both strands of the DNA being synthesized. These oligos are designed such that a single oligo from one strand contains a length of approximately 20 nucleotides at each end that is complementary to sequences of two different oligos on the opposite strand, thereby creating regions of overlap. The entire set is processed through cycles of: (a) hybridization at 60 °C; (b) elongation via  Taq polymerase  and a standard ligase; and (c) denaturation at 95 °C, forming progressively longer contiguous strands and ultimately resulting in the final genome. [ 6 ]  PCA was used to generate the first synthetic genome in history, that of the  Phi X 174 virus . [ 7 ]


The  Gibson assembly method , designed by Daniel Gibson during his time at the  J. Craig Venter Institute , requires a set of double-stranded DNA cassettes that constitute the entire genome being synthesized. Note that cassettes differ from contigs by definition, in that these sequences contain regions of homology to other cassettes for the purposes of  recombination . In contrast to Polymerase Cycling Assembly, Gibson Assembly is a single-step, isothermal reaction with larger sequence-length capacity; ergo, it is used in place of Polymerase Cycling Assembly for genomes larger than 6 kb.


A  T5 exonuclease  performs a chew-back reaction at the terminal segments, working in the 5' to 3' direction, thereby producing complementary overhangs. The overhangs hybridize to each other, a  Phusion DNA polymerase  fills in any missing nucleotides and the nicks are sealed with a ligase. However, the genomes capable of being synthesized using this method alone is limited because as DNA cassettes increase in length, they require propagation in vitro in order to continue hybridizing; accordingly, Gibson assembly is often used in conjunction with  transformation-associated recombination  (see below) to synthesize genomes several hundred kilobases in size. [ 8 ]


The goal of  transformation-associated recombination  (TAR) technology in synthetic genomics is to combine DNA contigs by means of  homologous recombination  performed by the  yeast artificial chromosome  (YAC). Of importance is the CEN element within the YAC vector, which corresponds to the yeast centromere. This sequence gives the vector the ability to behave in a chromosomal manner, thereby allowing it to perform homologous recombination. [ 9 ]


First,  gap repair cloning  is performed to generate regions of homology flanking the DNA contigs. Gap Repair Cloning is a particular form of the  polymerase chain reaction  in which specialized  primers  with extensions beyond the sequence of the DNA target are utilized. [ 10 ]  Then, the DNA cassettes are exposed to the YAC vector, which drives the process of homologous recombination, thereby connecting the DNA cassettes. Polymerase Cycling Assembly and TAR technology were used together to construct the 600 kb  Mycoplasma genitalium  genome in 2008, the first synthetic organism ever created. [ 11 ]  Similar steps were taken in synthesizing the larger  Mycoplasma mycoides  genome a few years later. [ 12 ]


An unnatural base pair (UBP) is a designed subunit (or  nucleobase ) of  DNA  which is created in a laboratory and does not occur in nature. In 2012, a group of American scientists led by  Floyd E. Romesberg , a chemical biologist at the  Scripps Research Institute  in San Diego, California, published that his team designed an unnatural base pair (UBP). [ 13 ]   The two new artificial nucleotides or  Unnatural Base Pair  (UBP) were named  d5SICS  and  dNaM . More technically, these artificial  nucleotides  bearing hydrophobic  nucleobases , feature two fused  aromatic rings  that form a (d5SICS–dNaM) complex or base pair in DNA. [ 14 ] [ 15 ]  In 2014 the same team from the Scripps Research Institute reported that they synthesized a stretch of circular DNA known as a  plasmid  containing natural T-A and C-G base pairs along with the best-performing UBP Romesberg's laboratory had designed, and inserted it into cells of the common bacterium  E. coli  that successfully replicated the unnatural base pairs through multiple generations. [ 16 ]  This is the first known example of a living organism passing along an expanded genetic code to subsequent generations. [ 14 ] [ 17 ]   This was in part achieved by the addition of a supportive algal gene that expresses a  nucleotide triphosphate  transporter which efficiently imports the triphosphates of both d5SICSTP and dNaMTP into  E. coli  bacteria. [ 14 ]  Then, the natural bacterial replication pathways use them to accurately replicate the  plasmid  containing d5SICS–dNaM.


The successful incorporation of a third base pair is a significant breakthrough toward the goal of greatly expanding the number of  amino acids  which can be encoded by DNA, from the existing 20 amino acids to a theoretically possible 172, thereby expanding the potential for living organisms to produce novel  proteins . [ 16 ]  The artificial strings of DNA do not encode for anything yet, but scientists speculate they could be designed to manufacture new proteins which could have industrial or pharmaceutical uses. [ 18 ]


In April 2019, scientists at  ETH Zurich  reported the creation of the world's first  bacterial genome , named  Caulobacter ethensis-2.0 , made entirely by a computer, although a related viable form of  C. ethensis-2.0  does not yet exist. [ 19 ] [ 20 ]


The  tetraploid complementation assay  is a technique in biology in which cells of two  mammalian   embryos  are combined to form a new embryo. [ 1 ]  It is used to construct  genetically modified organisms , to study the consequences of certain mutations on embryonal development, and in the study of  pluripotent   stem cells .


Normal mammalian  somatic cells  are  diploid : each  chromosome  (and thus every  gene ) is present in duplicate (excluding genes from X chromosome absent in Y chromosome). The assay starts with producing a  tetraploid  cell in which every chromosome exists fourfold. This is done by taking an embryo at the two-cell stage and fusing the two cells by applying an electrical current. The resulting tetraploid cell will continue to divide, and all daughter cells will also be tetraploid.


Such a tetraploid embryo can develop normally to the  blastocyst  stage and will  implant  in the wall of the  uterus . The tetraploid cells can form the extra-embryonic tissue ( placenta , etc.); however, a proper  fetus  will rarely develop.


In the tetraploid complementation assay, one now combines such a tetraploid embryo (either at the  morula  or  blastocyst  stage) with normal diploid  embryonic stem cells  (ES) from a different organism. The embryo will then develop normally; the fetus is exclusively derived from the ES cell, while the extra-embryonic tissues are exclusively derived from the tetraploid cells.


A  therapeutic interfering particle  is an antiviral preparation that reduces the replication rate and pathogenesis of a particular viral infectious disease. A therapeutic interfering particle is typically a biological agent (i.e., nucleic acid) engineered from portions of the viral genome being targeted. Similar to  Defective Interfering Particles (DIPs) , the agent competes with the pathogen within an infected cell for critical viral replication resources, reducing the viral replication rate and resulting in reduced pathogenesis. [ 1 ] [ 2 ]  But, in contrast to DIPs, TIPs are engineered to have an in vivo  basic reproductive ratio  ( R 0 ) that is greater than 1 ( R 0 >1). [ 3 ]  The term "TIP" was first introduced in 2011 [ 4 ]  based on models of its mechanism-of-action from 2003. [ 3 ]  Given their unique  R 0 >1 mechanism of action, TIPs exhibit high barriers to the evolution of antiviral resistance [ 5 ]  and are predicted to be resistance proof. [ 4 ]  Intervention with therapeutic interfering particles can be prophylactic (to prevent or ameliorate the effects of a future infection), or a single-administration therapeutic (to fight a disease that has already occurred, such as HIV or COVID-19). [ 6 ] [ 4 ] [ 3 ] [ 7 ] [ 5 ]  Synthetic DIPs that rely on stimulating innate antiviral immune responses (i.e., interferon) were proposed for influenza in 2008 [ 8 ]  and shown to protect mice to differing extents  [ 9 ] [ 10 ] [ 11 ]  but are technically distinct from TIPs due to their alternate molecular mechanism of action which has not been predicted to have a similarly high barrier to resistance. [ 12 ]   Subsequent work tested the pre-clinical efficacy of TIPs against HIV, [ 6 ]  a synthetic DIP for SARS-CoV-2 (in vitro), [ 7 ]  and a TIP for SARS-CoV-2 (in vivo). [ 5 ] [ 13 ]


Therapeutic Interfering Particles, often referred to as TIPs, are typically synthetic, engineered versions of naturally occurring  defective interfering particles  (DIPs), in which critical portions of the virus genome are deleted rendering the TIP unable to replicate on its own. Often a TIP has the vast majority of the virus genome deleted. [ 5 ]  However, TIPs are engineered to retain specific elements of the genome that allow them to efficiently compete with the wild-type virus for critical replication resources inside an infected cell. TIPs thereby deprive wild-type virus of replication material through  competitive inhibition , [ 14 ]  and therapeutically reduce viral load. [ 6 ]  Competitive inhibition enables TIPs to conditionally replicate and efficiently mobilize between cells, essentially "piggybacking" on wild-type virus, to act as single-administration antivirals with a high genetic barrier to the evolution of resistance. [ 15 ]  TIPs have been engineered for HIV [ 6 ] [ 14 ]  and SARS-CoV-2, [ 7 ]  and do not induce innate immune responses such as  interferon [ 5 ]


Three mechanistic criteria define a TIP:


As a result of these mechanistic criteria, TIPs have been referred to as "piggyback" [ 17 ]  or alternatively as "virus hijackers". [ 18 ] [ 19 ]


TIPs do not stimulate or function through the induction of innate cellular immune responses (such as interferon). In fact, stimulation of innate cellular antiviral mechanisms has been shown to contravene criterion (#3) (i.e.,  R 0 >1), as innate immune mechanisms inhibit efficient mobilization of TIPs. [ 3 ]  As such, several VLP-based therapy proposals for influenza and other viruses [ 20 ]  that do not satisfy these criteria are DIPs, but not TIPs.


TIPs are built off the phenomenon of  defective interfering particles  (DIPs) discovered by  Preben Von Magnus  in the early 1950s, during his work on influenza viruses. [ 21 ] [ 22 ] [ 23 ] [ 2 ]  DIPs are spontaneously arising virus mutants, first described by von Magnus as "incomplete" viruses, in which a critical portion of the viral genome has been lost. Direct evidence for DIPs was only found in the 1960s by Hackett, who observed the presence of "stumpy" particles of vesicular stomatitis virus in electron micrographs, [ 24 ]  and the DIP terminology was formalized in 1970 by Huang and Baltimore. [ 25 ]  DIPs have been reported for many classes of DNA and RNA viruses in clinical and laboratory settings.


Whereas DIPs had been proposed as potential therapeutics that would act via stimulation of the immune system [ 20 ]  – a concept [ 8 ]   [ 26 ]   tested in influenza with mixed results [ 9 ]   [ 10 ]  – the TIP R 0 >1 mechanism of action was first proposed in 2003 [ 3 ]  with the term “TIP” and the unique benefits of the R 0 >1 mechanism shown in 2011. [ 4 ]


In 2016 the US government launched a major funding initiative ( DARPA  INTERCEPT, [ 26 ] [ 27 ] [ 28 ]  ) to discover and engineer antiviral TIPs for diverse viruses, based on prior investments from the US  National Institutes of Health . [ 29 ]  This program led to renewed interest in the concept of interfering particles as therapies with the development of technologies to isolate DIPs for influenza [ 30 ] [ 31 ] [ 32 ]  and engineer TIPs for HIV and Zika virus. [ 14 ]  The first successful experimental demonstration of the TIP concept was reported in 2019 [ 6 ]  for HIV, and the discovery of a TIP for SARS-CoV-2 was reported in 2020 [ 7 ]  and results on the effect on hamsters in 2021. [ 33 ]  In 2020, the US government funded first-in-human clinical trials of TIPs. [ 34 ] [ 35 ]


Toehold mediated strand displacement (TMSD)  is an  enzyme -free  molecular tool  to exchange one strand of  DNA  or  RNA  (output) with another strand (input). It is based on the hybridization of two complementary strands of DNA or RNA via  Watson-Crick base pairing  (A-T/U and C-G) and makes use of a process called  branch migration . [ 1 ]  Although  branch migration  has been known to the scientific community since the 1970s, TMSD has not been introduced to the field of DNA nanotechnology until 2000 when Yurke et al. was the first who took advantage of TMSD. [ 1 ] [ 2 ]  He used the technique to open and close a set of DNA tweezers made of two  DNA helices  using an auxiliary strand of DNA as fuel. [ 1 ] [ 3 ]  Since its first use, the technique has been modified for the construction of autonomous molecular motors, catalytic amplifiers, reprogrammable DNA nanostructures and  molecular logic gates . [ 3 ] [ 4 ]  It has also been used in conjunction with RNA for the production of kinetically-controlled ribosensors. [ 5 ]  TMSD starts with a double-stranded DNA complex composed of the original strand and the protector strand. [ 2 ]  The original strand has an overhanging region the so-called “toehold” which is complementary to a third strand of DNA referred to as the “invading strand”. The invading strand is a sequence of  single-stranded DNA  (ssDNA) which is complementary to the original strand. [ 3 ] [ 2 ]  The toehold regions initiate the process of TMSD by allowing the complementary invading strand to hybridize with the original strand, creating a DNA complex composed of three strands of DNA. [ 3 ] [ 6 ]  This initial endothermic step is rate limiting [ 1 ]  and can be tuned by varying the strength (length and sequence composition e.g. G-C or A-T rich strands) of the toehold region. [ 3 ]  The ability to tune the rate of strand displacement over a range of 6 orders of magnitude generates the backbone of this technique and allows the kinetic control of DNA or RNA devices. [ 4 ]
After the binding of the invading strand and the original strand occurred,  branch migration  of the invading domain then allows the displacement of the initial hybridized strand (protector strand). [ 1 ]  The protector strand can possess its own unique toehold and can, therefore, turn into an invading strand itself, starting a  strand-displacement cascade . [ 2 ] [ 4 ] [ 7 ]  The whole process is energetically favored and although a reverse reaction can occur its rate is up to 6 orders of magnitude slower. [ 4 ]
Additional control over the system of toehold mediated strand displacement can be introduced by toehold sequestering. [ 4 ] [ 8 ] [ 9 ]


A slightly  different variant of strand displacement  has also been introduced using a strand displacing polymerase enzyme. [ 10 ] [ 11 ]  Unlike TMSD, it used the  polymerase enzyme  as a source of energy and it referred to as polymerase-based strand displacement. [ 11 ]


Toehold sequestering is a technique to “mask” the toehold region, rendering its accessibility. [ 4 ] [ 3 ]  There are several ways to do so but the most common approaches are hybridizing the toehold with a complementary strand [ 7 ]  or by designing the toehold region to form a  hairpin loop . [ 12 ]  Masking and unmasking of the toehold domains together with the ability to precisely control the kinetics of the reaction makes toehold mediated strand displacement a valuable tool in the field of  DNA nanotechnology [ 4 ]  Moreover, biosensors based on toehold mediated strand displacement reaction are useful in single molecule detection of DNA targets and SNP discrimination. [ 13 ]


Transcription activator-like effector nucleases  ( TALEN ) are  restriction enzymes  that can be engineered to cut specific sequences of DNA. They are made by fusing a  TAL effector   DNA-binding domain  to a DNA cleavage domain (a  nuclease  which cuts DNA strands). Transcription activator-like effectors (TALEs) can be engineered to bind to practically any desired DNA sequence, so when combined with a nuclease, DNA can be cut at specific locations. [ 1 ]  The restriction enzymes can be introduced into cells, for use in  gene editing  or for  genome  editing  in situ , a technique known as  genome editing with engineered nucleases . Alongside  zinc finger nucleases  and  CRISPR/Cas9 , TALEN is a prominent tool in the field of  genome editing .


TAL effectors  are proteins that are secreted by  Xanthomonas  bacteria via their  type III secretion system  when they  infect plants . [ 2 ]  The DNA binding domain contains a repeated highly conserved 33–34  amino acid  sequence with divergent 12th and 13th amino acids. These two positions, referred to as the Repeat Variable Diresidue (RVD), are highly variable and show a strong correlation with specific  nucleotide  recognition. [ 3 ] [ 4 ]   This straightforward relationship between amino acid sequence and DNA recognition has allowed for the engineering of specific DNA-binding domains by selecting a combination of repeat segments containing the appropriate RVDs. [ 1 ]  Notably, slight changes in the RVD and the incorporation of "nonconventional" RVD sequences can improve targeting specificity. [ 5 ]


The non-specific DNA cleavage domain from the end of the  FokI  endonuclease can be used to construct hybrid  nucleases  that are active in a yeast assay. [ 6 ] [ 7 ]   These reagents are also active in plant cells [ 8 ] [ 9 ]   and in animal cells. [ 9 ] [ 10 ] [ 11 ] [ 12 ]  Initial TALEN studies used the wild-type FokI cleavage domain, but some subsequent TALEN studies [ 11 ] [ 13 ] [ 14 ]  also used FokI cleavage domain variants with mutations designed to improve cleavage specificity [ 15 ] [ 16 ]  and cleavage activity. [ 17 ]  The FokI domain functions as a dimer, requiring two constructs with unique DNA binding domains for sites in the target genome with proper orientation and spacing.  Both the number of amino acid residues between the TALE DNA binding domain and the FokI cleavage domain and the number of bases between the two individual TALEN binding sites appear to be important parameters for achieving high levels of activity. [ 10 ] [ 18 ]


The simple relationship between amino acid sequence and DNA recognition of the TALE binding domain allows for the efficient engineering of proteins. In this case,  artificial gene synthesis  is problematic because of improper  annealing  of the repetitive sequence found in the TALE binding domain. [ 19 ]  One solution to this is to use a publicly available software program (DNAWorks [ 20 ] ) to calculate  oligonucleotides  suitable for assembly in a two step  PCR  oligonucleotide assembly followed by whole gene amplification. A number of modular assembly schemes for generating engineered TALE constructs have also been reported. [ 9 ] [ 19 ] [ 21 ] [ 22 ] [ 23 ] [ 24 ]  Both methods offer a systematic approach to engineering DNA binding domains that is conceptually similar to the modular assembly method for generating  zinc finger  DNA recognition domains.


Once the TALEN constructs have been assembled, they are inserted into  plasmids ; the target cells are then  transfected  with the plasmids, and the gene products are  expressed  and enter the nucleus to access the genome. Alternatively, TALEN constructs can be delivered to the cells as mRNAs, which removes the possibility of genomic integration of the TALEN-expressing protein. Using an mRNA vector can also dramatically increase the level of homology directed repair (HDR) and the success of introgression during gene editing.


TALEN can be used to edit genomes by inducing double-strand breaks (DSB), which cells respond to with repair mechanisms.


Non-homologous end joining  (NHEJ) directly ligates DNA from either side of a double-strand break where there is very little or no sequence overlap for annealing. This repair mechanism induces errors in the genome via  indels  (insertion or deletion), or chromosomal rearrangement; any such errors may render the gene products coded at that location non-functional. [ 10 ]   Because this activity can vary depending on the species, cell type, target gene, and nuclease used, it should be monitored when designing new systems.  A simple heteroduplex cleavage assay can be run which detects any difference between two alleles amplified by PCR. Cleavage products can be visualized on simple agarose gels or slab gel systems.


Alternatively, DNA can be introduced into a genome through NHEJ in the presence of exogenous double-stranded DNA fragments. [ 10 ]


Homology directed repair  can also introduce foreign DNA at the DSB as the transfected double-stranded sequences are used as templates for the repair enzymes. [ 10 ]


TALEN has been used to efficiently modify plant genomes, [ 25 ]  creating economically important food crops with favorable nutritional qualities. [ 26 ]   They have also been harnessed to develop tools for the production of  biofuels . [ 27 ]   In addition, it has been used to engineer stably modified human  embryonic stem cell  and  induced pluripotent stem cell  (IPSCs) clones and human erythroid cell lines, [ 11 ] [ 28 ]  to generate knockout  C. elegans , [ 12 ]   knockout rats , [ 13 ]  knockout mice, [ 29 ]  and knockout  zebrafish . [ 14 ] [ 30 ]  Moreover, the method can be used to generate knockin organisms. Wu et al.obtained a Sp110 knockin cattle using Talen nickases to induce increased resistance of tuberculosis. [ 31 ]  This approach has also been used to generate knockin rats by TALEN mRNA microinjection in one-cell embryos. [ 32 ]


TALEN has also been utilized experimentally to correct the genetic errors that underlie disease. [ 33 ]  For example, it has been used  in vitro  to correct the genetic defects that cause disorders such as  sickle cell disease , [ 28 ] [ 34 ]   xeroderma pigmentosum , [ 35 ]  and  epidermolysis bullosa . [ 36 ]  Recently, it was shown that TALEN can be used as tools to harness the immune system to fight cancers; TALEN-mediated targeting can generate T cells that are resistant to chemotherapeutic drugs and show anti-tumor activity. [ 37 ] [ 38 ]


In theory, the genome-wide specificity of engineered TALEN fusions allows for correction of errors at individual genetic loci via homology-directed repair from a correct exogenous template. [ 33 ]   In reality, however, the  in situ  application of TALEN is currently limited by the lack of an efficient delivery mechanism, unknown immunogenic factors, and uncertainty in the specificity of TALEN binding. [ 33 ]


Another emerging application of TALEN is its ability to combine with other genome engineering tools, such as  meganucleases . The DNA binding region of a TAL effector can be combined with the cleavage domain of a meganuclease to create a hybrid architecture combining the ease of engineering and highly specific DNA binding activity of a TAL effector with the low site frequency and specificity of a meganuclease. [ 39 ]


In comparison to other genome editing techniques, TALEN falls in the middle in terms of difficulty and cost. Unlike  ZFNs , TALEN recognizes single nucleotides. It's far more straightforward to engineer interactions between TALEN DNA binding domains and their target nucleotides than it is to create interactions with ZFNs and their target nucleotide triplets. [ 40 ]  On the other hand,  CRISPR  relies on ribonucleotide complex formation instead of protein/DNA recognition. gRNAs [ definition needed ]  have occasionally limitations regarding feasibility due to lack of PAM sites [ definition needed ]  in the target sequence and even though they can be cheaply produced, the current development lead to a remarkable decrease of cost for TALENs, so that they are in a similar price and time range like CRISPR based genome editing [ clarification needed ] .


The off-target activity of an active nuclease may lead to unwanted double-strand breaks and may consequently yield chromosomal rearrangements and/or cell death.  Studies have been carried out to compare the relative nuclease-associated toxicity of available technologies.  Based on these studies  [ 18 ]  and the maximal theoretical distance between DNA binding and nuclease activity, TALEN constructs are believed to have the greatest precision of the currently available technologies. [ 41 ]


A  transgene  is a  gene  that has been transferred naturally, or by any of a number of  genetic engineering  techniques, from one organism to another. The introduction of a transgene, in a process known as  transgenesis , has the potential to change the  phenotype  of an organism.  Transgene  describes a segment of  DNA  containing a gene sequence that has been isolated from one organism and is introduced into a different organism. This non-native segment of DNA may either retain the ability to produce  RNA  or  protein  in the transgenic organism or alter the normal function of the transgenic organism's genetic code. In general, the DNA is incorporated into the organism's  germ line . For example, in  higher vertebrates  this can be accomplished by injecting the foreign DNA into the  nucleus  of a fertilized  ovum . This technique is routinely used to introduce human disease genes or other genes of interest into strains of  laboratory mice  to study the function or  pathology  involved with that particular gene.


The construction of a transgene requires the assembly of a few main parts. The transgene must contain a  promoter , which is a regulatory sequence that will determine where and when the transgene is active, an  exon , a protein coding sequence (usually derived from the  cDNA  for the protein of interest), and a stop sequence. These are typically combined in a bacterial  plasmid  and the coding sequences are typically chosen from transgenes with previously known functions. [ 1 ]


Transgenic or  genetically modified organisms , be they bacteria, viruses or fungi, serve many research purposes.  Transgenic plants , insects, fish and mammals (including humans) have been bred. Transgenic plants such as corn and soybean have replaced wild strains in agriculture in some countries (e.g. the United States). Transgene escape has been documented for GMO crops since 2001 with persistence and invasiveness. Transgenetic organisms pose ethical questions and may cause  biosafety  problems.


The idea of shaping an organism to fit a specific need is not a new science. However, until the late 1900s farmers and scientists could breed new strains of a plant or organism only from closely related species because the DNA had to be compatible for offspring to be able to reproduce. [ citation needed ]


In the 1970 and 1980s, scientists passed this hurdle by inventing procedures for combining the DNA of two vastly different species with  genetic engineering . The organisms produced by these procedures were termed transgenic. Transgenesis is the same as  gene therapy  in the sense that they both transform cells for a specific purpose. However, they are completely different in their purposes, as gene therapy aims to cure a defect in cells, and transgenesis seeks to produce a genetically modified organism by incorporating the specific transgene into every cell and changing the  genome . Transgenesis will therefore change the germ cells, not only the somatic cells, in order to ensure that the transgenes are passed down to the offspring when the organisms reproduce. Transgenes alter the genome by blocking the function of a host gene; they can either replace the host gene with one that codes for a different protein, or introduce an additional gene. [ 2 ]