Genome editing

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Genome editing, or genome engineering is a type of genetic engineering in which DNA is inserted, deleted, modified or replaced in the genome of a living organism. In 2018, the common methods for such editing use engineered nucleases, or "molecular scissors". These nucleases create site-specific double-strand breaks (DSBs) at desired locations in the genome. The induced double-strand breaks are repaired through nonhomologous end-joining (NHEJ) or homologous recombination (HR), resulting in targeted mutations ('edits').

As of 2015 four families of engineered nucleases were used: meganucleases, zinc finger nucleases (ZFNs), transcription activator-like effector-based nucleases (TALEN), and the clustered regularly interspaced short palindromic repeats (CRISPR/Cas9) system.[1][2][3][4] Nine genome editors were available as of 2017.[5]

Genome editing was selected by Nature Methods as the 2011 Method of the Year.[6] The CRISPR-Cas system was selected by Science as 2015 Breakthrough of the Year.[7]


A common approach in modern biological research is to modify the DNA sequence (genotype) of an organism (or a single cell) and observe the impact of this change on the organism (phenotype). This approach is called reverse genetics and its significance for modern biology lies in its relative simplicity. This method contrasts with that of forward genetics, where a new phenotype is first observed and then its genetic basis is studied. This course is more complex because phenotypic changes are often a result of multiple genetic interactions.

Among the key requirements of reverse genetic analysis is the ability to modify the DNA sequence of the target organism. This can be achieved by:

These approaches have several drawbacks:

  • PCR- and phage-mediated approaches are less successful in more complex organisms such as mammals, where delivery becomes more difficult.[citation needed]
  • They also require stringent selection steps and thus the addition of selection-specific sequences, along with those incorporated into the DNA.
  • Recombination-based method.g. in mouse embryonic stem cells treated with donor DNA, only 1 in a million DNA molecules was incorporated at the desired position.[10]

The use of other mutagenic techniques (such as P-element transgenesis in Drosophila) also has limitations, the major one being the randomness of incorporation and the possibility of affecting other genes and expression patterns.

Hence, genome editing with engineered nucleases (GEEN) is a promising new approach. This rapidly evolving technology overcomes these shortcomings and uses relatively simple concepts.

Double stranded breaks

Fundamental to the use of nucleases in genome editing is the concept of DNA double stranded break (DSB) repair mechanics. These DSBs occur due to the use of specific enzymes which break the weak hydrogen bonds holding the two strands of DNA together in a double helical structure. One of the known DSB repair pathways that are essentially functional in all organisms[citation needed] are the non-homologous end joining (NHEJ) and homology directed repair (HDR).

NHEJ uses a variety of enzymes to directly join the DNA ends in a double-strand break. In contrast, in HDR, a homologous sequence is utilized as a template for regeneration of missing DNA sequence at the break point. The natural properties of these pathways form the very basis of nuclease-based genome editing.

NHEJ is error-prone, and has been shown to cause mutations at the repair site in approximately 50% of DSB in mycobacteria,[11] and also its low fidelity has been linked to mutational accumulation in leukemias.[12] Thus if one is able to create a DSB at a desired gene in multiple samples, it is very likely that mutations will be generated at that site in some of the treatments because of errors created by the NHEJ infidelity.

On the other hand, the dependency of HDR on a homologous sequence to repair DSBs can be exploited by inserting a desired sequence within a sequence that is homologous to the flanking sequences of a DSB which, when used as a template by HDR system, would lead to the creation of the desired change within the genomic region of interest.

Despite the distinct mechanisms, the concept of the HDR based gene editing is in a way similar to that of homologous recombination based gene targeting. However, the rate of recombination is increased by at least three orders of magnitude when DSBs are created and HDR is at work thus making the HDR based recombination much more efficient and eliminating the need for stringent positive and negative selection steps.[13] So based on these principles if one is able to create a DSB at a specific location within the genome, then the cell’s own repair systems will help in creating the desired mutations.

Site-specific double stranded breaks

Creation of a DSB in DNA should not be a challenging task as the commonly used restriction enzymes are capable of doing so. However, if genomic DNA is treated with a particular restriction endonuclease many DSBs will be created. This is a result of the fact that most restriction enzymes recognize a few base pairs on the DNA as their target and very likely that particular base pair combination will be found in many locations across the genome. To overcome this challenge and create site-specific DSB, three distinct classes of nucleases have been discovered and bioengineered to date. These are the Zinc finger nucleases (ZFNs), transcription-activator like effector nucleases (TALEN) and meganucleases. Below is a brief overview and comparison of these enzymes and the concept behind their development.

Engineered nucleases

Groups of engineered nucleases used for GEEN. Matching colors signify DNA recognition patterns

Meganuclease-based engineering

Meganucleases, discovered in the late 1980s, are enzymes in the endonuclease family which are characterized by their capacity to recognize and cut large DNA sequences (from 12 to 40 base pairs).[14] The most widespread and best known meganucleases are the proteins in the LAGLIDADG family, which owe their name to a conserved amino acid sequence.

Meganucleases, found commonly in microbial species, have the unique property of having very long recognition sequences (>14bp) thus making them naturally very specific.[15][16] However, there is virtually no chance of finding the exact meganuclease required to act on a specific DNA sequence. To overcome this challenge, mutagenesis and high throughput screening methods have been used to create meganuclease variants that recognize unique sequences.[16] Others have been able to fuse various meganucleases and create hybrid enzymes that recognize a new sequence.[17] Yet others have attempted to alter the DNA interacting aminoacids of the meganuclease to design sequence specific meganucelases in a method named rationally designed meganuclease (US Patent 8,021,867 B2).

There are two methods, which can be combined, for creating custom meganucleases:

  • Mutagenesis involves generating collections of variants using a meganuclease with properties similar to the desired enzyme, then selecting these variants using high-throughput screening. This procedure can be optimized by adopting what are known as "semi-rational" methods, in which the structural data is electronically processed in order to focus the mutagenesis to the part of the enzyme that interacts with DNA and triggers the cleavage.[18]
  • Combinatorial assembly is a method whereby protein subunits from different enzymes can be associated or fused.[19]

A large bank containing several tens of thousands of protein units has been created. These units can be combined to obtain chimeric meganucleases that recognize the target site, thereby providing research and development tools that meet a wide range of needs (fundamental research, health, agriculture, industry, energy, etc.).

This technique has enabled the development of several meganucleases specific for sequences in the genomes of viruses, plants, etc., and the industrial-scale production of two meganucleases able to cleave the human XPC gene; mutations in this gene result in Xeroderma pigmentosum, a severe monogenic disorder that predisposes the patients to skin cancer and burns whenever their skin is exposed to UV rays.[20]

Another approach involves using computer models to try to predict as accurately as possible the activity of the modified meganucleases and the specificity of the recognized nucleic sequence.[21] The Northwest Genome Engineering Consortium, a US consortium funded by the National Institutes of Health, has adopted this approach with the aim of treating leukemia by modifying hematopoietic stem cells. The model’s prediction has been verified and guided by means of directed mutagenesis and in vitro biochemical analysis.

A third approach has been taken by the American biotechnology company Precision Biosciences, Inc. The company, funded by the National Institutes of Health and the National Institute of Standards and Technology, has developed a fully rational design process called the Directed Nuclease Editor (DNE) which is capable of creating highly specific engineered meganucleases that successfully target and modify a user-defined location in a genome.[22]

Meganucleases have the benefit of causing less toxicity in cells than methods such as Zinc finger nuclease (ZFN), likely because of more stringent DNA sequence recognition;[16] however, the construction of sequence-specific enzymes for all possible sequences is costly and time consuming, as one is not benefiting from combinatorial possibilities that methods such as ZFNs and TALEN-based fusions utilize.

Zinc finger nuclease-based engineering

As opposed to meganucleases, the concept behind ZFNs and TALEN technology is based on a non-specific DNA cutting enzyme, which can then be linked to specific DNA sequence recognizing peptides such as zinc fingers and transcription activator-like effectors (TALEs).[23] The key to this was to find an endonuclease whose DNA recognition site and cleaving site were separate from each other, a situation that is not common among restriction enzymes.[23] Once this enzyme was found, its cleaving portion could be separated which would be very non-specific as it would have no recognition ability. This portion could then be linked to sequence recognizing peptides that could lead to very high specificity

Zinc finger motifs occur in several transcription factors. The zinc ion, found in 8% of all human proteins, plays an important role in the organization of their three-dimensional structure. In transcription factors, it is most often located at the protein-DNA interaction sites, where it stabilizes the motif. The C-terminal part of each finger is responsible for the specific recognition of the DNA sequence.

The recognized sequences are short, made up of around 3 base pairs, but by combining 6 to 8 zinc fingers whose recognition sites have been characterized, it is possible to obtain specific proteins for sequences of around 20 base pairs. It is therefore possible to control the expression of a specific gene. It has been demonstrated that this strategy can be used to promote a process of angiogenesis in animals.[24] It is also possible to fuse a protein constructed in this way with the catalytic domain of an endonuclease in order to induce a targeted DNA break, and therefore to use these proteins as genome engineering tools.[25]

The method generally adopted for this involves associating two proteins – each containing 3 to 6 specifically chosen zinc fingers – with the catalytic domain of the FokI endonuclease. The two proteins recognize two DNA sequences that are a few nucleotides apart. Linking the two zinc finger proteins to their respective sequences brings the two endonucleases associated with them closer together. FokI requires dimerization to have nuclease activity and this means the specificity increases dramatically as each nuclease partner would recognize a unique DNA sequence. To enhance this effect, FokI nucleases have been engineered that can only function as heterodimers and have increased catalytic activity.[26]

Several approaches are used to design specific zinc finger nucleases for the chosen sequences. The most widespread involves combining zinc-finger units with known specificities (modular assembly). Various selection techniques, using bacteria, yeast or mammal cells have been developed to identify the combinations that offer the best specificity and the best cell tolerance. Although the direct genome-wide characterization of zinc finger nuclease activity has not been reported, an assay that measures the total number of double-strand DNA breaks in cells found that only one to two such breaks occur above background in cells treated with zinc finger nucleases with a 24 bp composite recognition site and obligate heterodimer FokI nuclease domains.[27]

The heterodimer functioning nucleases would avoid the possibility of unwanted homodimer activity and thus increase specificity of the DSB. Although the nuclease portions of both ZFNs and TALEN constructs have similar properties, the difference between these engineered nucleases is in their DNA recognition peptide. ZFNs rely on Cys2-His2 zinc fingers and TALEN constructs on TALEs. Both of these DNA recognizing peptide domains have the characteristic that they are naturally found in combinations in their proteins. Cys2-His2 Zinc fingers typically happen in repeats that are 3 bp apart and are found in diverse combinations in a variety of nucleic acid interacting proteins such as transcription factors. TALEs on the other hand are found in repeats with a one-to-one recognition ratio between the amino acids and the recognized nucleotide pairs. Because both zinc fingers and TALEs happen in repeated patterns, different combinations can be tried to create a wide variety of sequence specificities.[15] Zinc fingers have been more established in these terms and approaches such as modular assembly (where Zinc fingers correlated with a triplet sequence are attached in a row to cover the required sequence), OPEN (low-stringency selection of peptide domains vs. triplet nucleotides followed by high-stringency selections of peptide combination vs. the final target in bacterial systems), and bacterial one-hybrid screening of zinc finger libraries among other methods have been used to make site specific nucleases.

Zinc finger nucleases are research and development tools that have already been used to modify a range of genomes, in particular by the laboratories in the Zinc Finger Consortium. The US company Sangamo BioSciences uses zinc finger nucleases to carry out research into the genetic engineering of stem cells and the modification of immune cells for therapeutic purposes.[28][29] Modified T lymphocytes are currently undergoing phase I clinical trials to treat a type of brain tumor (glioblastoma) and in the fight against AIDS.[27]


General overview of the TALEN process

Transcription activator-like effector nucleases (TALENs) are artificial restriction enzymes generated by fusing a specific DNA-binding domain to a non-specific DNA cleaving domain. The DNA binding domains, which can be designed to bind any desired DNA sequence, comes from TAL effectors, DNA-binding proteins excreted by plant pathogenic Xanthomanos app. Tal effectors consists of repeated domains, each which contains a highly considered sequence of 34 amino acids, and recognize a single DNA nucleotide. The nuclease can create double strand breaks at the target site that can be repaired by error-prone non-homologous end-joining (NHEJ), resulting in gene disruptions through the introduction of small insertions or deletions. TALEN constructs are used in a similar way to designed zinc finger nucleases, and have three advantages in targeted mutagenesis: (1) DNA binding specificity is higher, (2) off-target effects are lower, and (3) construction of DNA-binding domains is easier.


CRISPRs (Clustered Regularly Interspaced Short Palindromic Repeats) are genetic elements that bacteria use as a kind of acquired immunity to protect against viruses. They consist of short sequences that originate from viral genomes and have been incorporated into the bacterial genome. Cas (CRISPR associated proteins) process these sequences and cut matching viral DNA sequences. By introducing plasmids containing Cas genes and specifically constructed CRISPRs into eukaryotic cells, the eukaryotic genome can be cut at any desired position.[30] Several companies, including Editas, have been working to monetize the CRISPR method while developing gene-specific therapies.[31][32]

Precision and efficiency of engineered nucleases

Meganucleases method of gene editing is the least efficient of the methods mentioned above. Due to the nature of its DNA-binding element and the cleaving element, it is limited to recognizing one potential target every 1,000 nucleotides.[33] ZFN was developed to overcome the limitations of meganuclease. The number of possible targets ZFN can recognized was increased to one in every 140 nucleotides.[33] However, both methods are unpredictable due to the ability of their DNA-binding elements affecting each other. As a result, high degrees of expertise and lengthy and costly validations processes are required.

TALE nucleases being the most precise and specific method yields a higher efficiency than the previous two methods. It achieves such efficiency because the DNA-binding element consists of an array of TALE subunits, each of them having the capability of recognizing a specific DNA nucleotide chain independent from others, resulting in a higher number of target sites with high precision. New TALE nucleases take about one week and a few hundred dollars to create, with specific expertise in molecular biology and protein engineering.[33]

CRISPR nucleases have a slightly lower precision when compared to the TALE nucleases. This is caused by the need of having a specific nucleotide at one end in order to produce the guide RNA that CRISPR uses to repair the double-strand break it induces. It has been shown to be the quickest and cheapest method, only costing less than two hundred dollars and a few days of time.[33] CRISPR also requires the least amount of expertise in molecular biology as the design lays in the guide RNA instead of the proteins. One major advantage that CRISPR has over the ZFN and TALEN methods is that it can directed to target different DNA sequences using its ~80nt CRISPR sgRNAs, while both ZFN and TALEN methods required construction and testing of the proteins created for targeting each DNA sequence.[34]

Because off-target activity of an active nuclease would have potentially dangerous consequences at the genetic and organismal levels, the precision of meganucleases, ZFNs, CRISPR, and TALEN-based fusions has been an active area of research. While variable figures have been reported, ZFNs tend to have more cytotoxicity than TALEN methods or RNA-guided nucleases, while TALEN and RNA-guided approaches tend to have the greatest efficiency and fewer off-target effects.[35] Based on the maximum theoretical distance between DNA binding and nuclease activity, TALEN approaches result in the greatest precision.[4]

Multiplex Automated Genomic Engineering (MAGE)

Synthetic DNA is repeatedly introduced at multiple targeted areas of the chromosome and/or loci and then is replicated producing cells with/without mutations.

The methods for scientists and researchers wanting to study genomic diversity and all possible associated phenotypes were very slow, expensive, and inefficient. Prior to this new revolution, researchers would have to do single-gene manipulations and tweak the genome one little section at a time, observe the phenotype, and start the process over with a different single-gene manipulation.[36] Therefore, researchers at the Wyss Institute at Harvard University designed the MAGE, a powerful technology that improves the process of in vivo genome editing. It allows for quick and efficient manipulations of a genome, all happening in a machine small enough to put on top of a small kitchen table. Those mutations combine with the variation that naturally occurs during cell mitosis creating billions of cellular mutations.

Chemically combined, synthetic single-stranded DNA (ssDNA) and a pool of oligionucleotides are introduced at targeted areas of the cell thereby creating genetic modifications. The cyclical process involves transformation of ssDNA (by electroporation) followed by outgrowth, during which bacteriophage homologous recombination proteins mediate annealing of ssDNAs to their genomic targets. Experiments targeting selective phenotypic markers are screened and identified by plating the cells on differential medias. Each cycle ultimately takes 2.5 hours to process, with additional time required to grow isogenic cultures and characterize mutations. By iteratively introducing libraries of mutagenic ssDNAs targeting multiple sites, MAGE can generate combinatorial genetic diversity in a cell population. There can be up to 50 genome edits, from single nucleotide base pairs to whole genome or gene networks simultaneously with results in a matter of days.[36]

MAGE experiments can be divided into three classes, characterized by varying degrees of scale and complexity: (i) many target sites, single genetic mutations; (ii) single target site, many genetic mutations; and (iii) many target sites, many genetic mutations.[36] An example of class three was reflected in 2009, where Church and colleagues were able to program Escherichia coli to produce five times the normal amount of lycopene, an antioxidant normally found in tomato seeds and linked to anti-cancer properties. They applied MAGE to optimize the 1-deoxy-d-xylulose-5-phosphate (DXP) metabolic pathway in Escherichia coli to overproduce isoprenoid lycopene. It took them about 3 days and just over $1,000 in materials. The ease, speed, and cost efficiency in which MAGE can alter genomes can transform how industries approach the manufacturing and production of important compounds in the bioengineering, bioenergy, biomedical engineering, synthetic biology, pharmaceutical, agricultural, and chemical industries.


Plants, animals and human genes that are successfully targeted using ZFN, which demonstrates the generality of this approach

As of 2012 efficient genome editing had been developed for a wide range of experimental systems ranging from plants to animals, often beyond clinical interest, and was becoming a standard experimental strategy in research labs.[37] The recent generation of rat, zebrafish, maize and tobacco ZFN-mediated mutants and the improvements in TALEN-based approaches testify to the significance of the methods, and the list is expanding rapidly. Genome editing with engineered nucleases will likely contribute to many fields of life sciences from studying gene functions in plants and animals to gene therapy in humans. For instance, the field of synthetic biology which aims to engineer cells and organisms to perform novel functions, is likely to benefit from the ability of engineered nuclease to add or remove genomic elements and therefore create complex systems.[37] In addition, gene functions can be studied using stem cells with engineered nucleases.

Listed below are some specific tasks this method can carry out:

Targeted gene modification in plants

Overview of GEEN workflow and editing possibilities

Genome editing using meganucleases,[38] ZFNs, and TALEN provides a new strategy for genetic manipulation in plants and are likely to assist in the engineering of desired plant traits by modifying endogenous genes. For instance, site-specific gene addition in major crop species can be used for 'trait stacking' whereby several desired traits are physically linked to ensure their co-segregation during the breeding processes.[26] Progress in such cases have been recently reported in Arabidopsis thaliana[39][40][41] and Zea mays. In Arabidopsis thaliana, using ZFN-assisted gene targeting, two herbicide-resistant genes (tobacco acetolactate synthase SuRA and SuRB) were introduced to SuR loci with as high as 2% transformed cells with mutations.[42] In Zea mays, disruption of the target locus was achieved by ZFN-induced DSBs and the resulting NHEJ. ZFN was also used to drive herbicide-tolerance gene expression cassette (PAT) into the targeted endogenous locus IPK1 in this case.[43] Such genome modification observed in the regenerated plants has been shown to be inheritable and was transmitted to the next generation.[43]

In addition, TALEN-based genome engineering has been extensively tested and optimized for use in plants.[44] TALEN fusions have also been used to improve the quality of soybean oil products[45] and to increase the storage potential of potatoes[46]

Several optimizations need to be made in order to improve editing plant genomes using ZFN-mediated targeting.[47] These include the reliable design and subsequent test of the nucleases, the absence of toxicity of the nucleases, the appropriate choice of the plant tissue for targeting, the routes of introduction or induction of enzyme activity, the lack of off-target mutagenesis, and a reliable detection of mutated cases.[47]

Gene therapy

The ideal gene therapy practice is that which replaces the defective gene with a normal allele at its natural location. This is advantageous over a virally delivered gene as there is no need to include the full coding sequences and regulatory sequences when only a small proportions of the gene needs to be altered as is often the case.[48] The expression of the partially replaced genes is also more consistent with normal cell biology than full genes that are carried by viral vectors.

Gene targeting through ZFNs or TALEN-based approaches can also be used to modify defective genes at their endogenous chromosomal locations. Examples include the treatment of X-linked severe combined immunodeficiency (X-SCID) by ex vivo gene correction with DNA carrying the interleukin-2 receptor common gamma chain (IL-2Rγ)[49] and the correction of Xeroderma pigmentosum mutations in vitro using TALEN.[50] Insertional mutagenesis by the retroviral vector genome induced leukemia in some patients, a problem that is predicted to be avoided by these technologies. However, ZFNs may also cause off-target mutations, in a different way from viral transductions. Currently many measures are taken to improve off-target detection and ensure safety before treatment.

In 2011, Sangamo BioSciences (SGMO) introduced the Delta 32 mutation (a suppressor of CCR5 gene which is a co-receptor for HIV-1 entry into T cells therefore enabling HIV infection) using Zinc Finger Nuclease (ZFN). Their results were presented at the 51st Interscience Conference on Antimicrobial Agents and Chemotherapy (ICAAC) held in Chicago from September 17–20, 2011.[51] Researchers at SGMO mutated CCR5 in CD4+ T cells and subsequently produced an HIV-resistant T-cell population.[52]

Gene editing is used to generate modified custom immune cells. For example, a recent report indicated that T cells could be modified to inactivate the glucocorticoid receptor; the resulting immune cells are fully functional but resistant to the effects of commonly used corticosteroids.[53] Similarly, scientists at Cellectis recently generated custom T-cells expressing chimeric antigen receptors using TALEN technology.[54] These T-cells can be engineered to be resistant to anti-cancer drugs and to invoke immune responses against targets of interest.[55]

The first clinical use of TALEN-based genome editing was in the treatment of CD19+ acute lymphoblastic leukemia in an 11-month old child.[56] Modified donor T cells were engineered to attack the leukemia cells, to be resistant to Alemtuzumab, and to evade detection by the host immune system after introduction. A few weeks after therapy, the patient's condition improved. Though physicians were cautious, the patient was still in remission more than one year after treatment.[57][58][59]

Eradicating diseases

CRISPR-Cas9 is being used in conjunction with gene drives as a potential to alter the traditional Mendelian laws of inheritance. Researchers have successfully used CRISPR-Cas9 gene drives to modify genes associated with sterility in A. gambiae, the vector for malaria.[60] If used widespread, this technique could suppress A. gambiae populations and with it malaria. Alternate genes are being researched that affect the vector's ability to carry the disease instead of causing intentional sterility and possible extinction. This technique has further implications in eradicating other vector borne diseases such as yellow fever, dengue, Zika, West Nile, Schistomiasis, Leishmaniasis and Lymes disease.

Given that there is currently no approved Lyme vaccine for humans, but there is for dogs, Kevin Esvelt, an associate professor of biological engineering at MIT has proposed using the vaccine on white-footed mice. If effective, CRISPR-Cas9 gene drives could be used in conjunction with genes associated with antibody forming in white-footed mice and then planted in the cells of mouse gametes. Esvelt is currently seeking approval of the residence of Nantucket, MA to utilize the island as a test environment for his experiment.[61]

The CRISPR-Cas9 system can be programmed to modulate the population of any bacterial species by targeting clinical genotypes or epidemiological isolates. It can selectively enable the beneficial bacterial species over the harmful ones by eliminating pathogen, which gives it an advantage over broad-spectrum antibiotics.[36]

Antiviral applications for therapies targeting human viruses such as HIV, herpes, and hepatitis B virus are under research. CRISPR can be used to target the virus or the host to disrupt genes encoding the virus cell-surface receptor proteins.[62]

Extensive research is being done on CRISPR-Cas9 in correcting genetic mutations which cause genetic diseases such as Down syndrome, spina bifida, anencephaly, and Tuner and Klinefelter syndromes using targeted gene therapy, if these genetic mutations are identified early enough in the embryo stages, which we currently already have the capability to do so consistently. With many of the genetic diseases caused from one overexpressed gene, CRISPR-Cas9 can be used to silence an entire chromosome, or delete the overexpressed gene with Cas9 endonuclease cutting.[63]

Other studies are being done on using CRISPR to target specific genes in cancer cells in hopes of using genome editing to inhibit cell proliferation and tumorigenicity of cancer cells.[64]

Prospects and limitations

In the future, an important goal of research into genome editing with engineered nucleases must be the improvement of the safety and specificity of the nucleases. For example, improving the ability to detect off-target events can improve our ability to learn about ways of preventing them. In addition, zinc-fingers used in ZFNs are seldom completely specific, and some may cause a toxic reaction. However, the toxicity has been reported to be reduced by modifications done on the cleavage domain of the ZFN.[48]

In addition, research by Dana Carroll into modifying the genome with engineered nucleases has shown the need for better understanding of the basic recombination and repair machinery of DNA. In the future, a possible method to identify secondary targets would be to capture broken ends from cells expressing the ZFNs and to sequence the flanking DNA using high-throughput sequencing.[48]

Because of the ease of use and cost-efficiency of CRISPR, extensive research is currently being done on it. There are now more publications on CRISPR than ZFN and TALEN despite how recent the discovery of CRISPR is.[62] Both CRISPR and TALEN are favored to be the choices to be implemented in large-scale productions due to their precision and efficiency.

Genome editing occurs also as a natural process without artificial genetic engineering. The agents that are competent to edit genetic codes are viruses or subviral RNA-agents.[65]

Although GEEN has higher efficiency than many other methods in reverse genetics, it is still not highly efficient; in many cases less than half of the treated populations obtain the desired changes.[42] For example, when one is planning to use the cell's NHEJ to create a mutation, the cell's HDR systems will also be at work correcting the DSB with lower mutational rates.

Traditionally, mice have been the most common choice for researchers as a host of a disease model. CRISPR can help bridge the gap between this model and human clinical trials by creating transgenic disease models in larger animals such as pigs, dogs, and non-human primates.[66][67] Using the CRISPR-Cas9 system, the programmed Cas9 protein and the sgRNA can be directly introduced into fertilized zygotes to achieve the desired gene modifications when creating transgenic models in rodents. This allows bypassing of the usual cell targeting stage in generating transgenic lines, and as a result, it reduces generation time by 90%.[67]

One potential that CRISPR brings with its effectiveness is the application of xenotransplantation. In previous research trials, CRISPR demonstrated the ability to target and eliminate endogenous retroviruses, which reduces the risk of transmitting diseases and reduces immune barriers.[62] Eliminating these problems improves donor organ function, which brings this application closer to a reality.

Human enhancement

Many transhumanists see genome editing as a potential tool for human enhancement.[68][69][70] Australian biologist and Professor of Genetics David Andrew Sinclair notes that "the new technologies with genome editing will allow it to be used on individuals (...) to have (...) healthier children" – designer babies.[71] According to a September 2016 report by the Nuffield Council on Bioethics in the future it may be possible to enhance people with genes from other organisms or wholly synthetic genes to for example improve night vision and sense of smell.[72][73]

The American National Academy of Sciences and National Academy of Medicine issued a report in February 2017 giving qualified support to human genome editing.[74] They recommended that clinical trials for genome editing might one day be permitted once answers have been found to safety and efficiency problems "but only for serious conditions under stringent oversight."[75]


In the 2016 Worldwide Threat Assessment of the US Intelligence Community statement United States Director of National Intelligence, James R. Clapper, named genome editing as a potential weapon of mass destruction, stating that genome editing conducted by countries with regulatory or ethical standards "different from Western countries" probably increases the risk of the creation of harmful biological agents or products. According to the statement the broad distribution, low cost, and accelerated pace of development of this technology, its deliberate or unintentional misuse might lead to far-reaching economic and national security implications.[76][77][78] For instance technologies such as CRISPR could be used to make "killer mosquitoes" that cause plagues that wipe out staple crops.[78]

According to a September 2016 report by the Nuffield Council on Bioethics, the simplicity and low cost of tools to edit the genetic code will allow amateurs – or "biohackers" – to perform their own experiments, posing a potential risk from the release of genetically modified bugs. The review also found that the risks and benefits of modifying a person's genome – and having those changes pass on to future generations – are so complex that they demand urgent ethical scrutiny. Such modifications might have unintended consequences which could harm not only the child, but also their future children, as the altered gene would be in their sperm or eggs.[72][73] In 2001 Australian researchers Ronald Jackson and Ian Ramshaw were criticized for publishing a paper in the Journal of Virology that explored the potential control of mice, a major pest in Australia, by infecting them with an altered mousepox virus that would cause infertility as the provided sensitive information could lead to the manufacture of biological weapons by potential bioterrorists who might use the knowledge to create vaccine resistant strains of other pox viruses, such as smallpox, that could affect humans.[73] Furthermore, there are additional concerns about the ecological risks of releasing gene drives into wild populations.[73][79][80]

See also


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Further reading

  • Customized Human Genes Scientific American articles
  • Connor, Steve (25 April 2014). "Scientific split - the human genome breakthrough dividing former colleagues". The Independent. Retrieved 2016-02-11. 
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