StemCell Therapy

$1 Million Awarded to Continue to Develop Genetically Engineered Stem Cell Products to Fight Gastroesophageal Cancer  PR Newswire

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$1 Million Awarded to Continue to Develop Genetically Engineered Stem Cell Products to Fight Gastroesophageal Cancer - PR Newswire

Engineered animals show new way to fight mercury pollution  EurekAlert

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Engineered animals show new way to fight mercury pollution - EurekAlert

Genetically modified foods: benefits and applications  Meer

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Genetically modified foods: benefits and applications - Meer

Genetically modified zebrafish and fruit flies munch on mercury to make it less toxic  Yahoo

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Genetically modified zebrafish and fruit flies munch on mercury to make it less toxic - Yahoo

1. Introduction

Since the identification of DNA as the unit of heredity and the basis for the central dogma of molecular biology [1] that DNA makes RNA and RNA makes proteins, scientists have pursued experiments and methods to understand how DNA controls heredity. With the discovery of molecular biology tools such as restriction enzymes, DNA sequencing, and DNA cloning, scientists quickly turned to experiments to change chromosomal DNA in cells and animals. In that regard, initial experiments that involved the co-incubation of viral DNA with cultured cell lines progressed to the use of selectable markers in plasmids. Delivery methods for random DNA integration have progressed from transfection by physical co-incubation of DNA with cultured cells, to electroporation and microinjection of cultured cells [2,3,4]. Moreover, the use of viruses to deliver DNA to cultured cells has progressed in tandem with physical methods of supplying DNA to cells [5,6,7]. Homologous recombination in animal cells [8] was rapidly exploited by the mouse genetics research community for the production of gene-modified mouse ES cells, and thus gene-modified whole animals [9,10].

This impetus to understand gene function in intact animals was ultimately manifested in the international knockout mouse project, the purpose of which was to knock out every gene in the mouse genome, such that researchers could choose to make knockout mouse models from a library of gene-targeted knockout ES cells [11,12,13]. Thousands of mouse models have resulted from that effort and have been used to better understand gene function and the bases of human genetic diseases [14]. This project required high-throughput pipelines for the construction of vectors, including bacterial artificial chromosome (BAC) recombineering technology [13,15,16,17]. BACs contain long segments of cloned genomic DNA. For example, the C57BL/6J mouse BAC library, RPCI-23, has an average insert size of 197 kb of genomic DNA per clone [18]. Because of their size, BACs often carry all of the genetic regulatory elements to faithfully recapitulate the expression of genes contained in them, and thus can be used to generate BAC transgenic mice [19,20]. Recombineering can be used to insert reporters in BACs that are then used to generate transgenic mice to accurately label cells and tissues according to the genes in the BACs [21,22,23,24,25,26]. A panoply of approaches to genetic engineering are available for researchers to manipulate the genome. ES cell and BAC transgene engineering have given way to directly editing genes in zygotes, consequently avoiding the need for ES cell or BAC intermediates on the way to an animal model.

Prior to the adaptation of Streptococcus pyogenes Cas9 protein to cause chromosome breaks, three other endonuclease systems were used: (1) rare-cutting meganucleases, (2) zinc finger nucleases (ZFNs), and (3) transcription activator-like effector (TALE) nucleases (TALENs) [27]. The I-CreI meganuclease recognizes a 22 bp DNA sequence [28,29]. Proof-of-concept experiments demonstrated that the engineered homing endonuclease I-CreI can be used to generate transgenic mice and transgenic rats [30]. I-CreI specificity can be adjusted to target specific sequences in DNA by protein engineering methodology, although this limits its widespread application to genetic engineering [31]. Subsequently, ZFN technology was developed to cause chromosome breaks [32]. A single zinc finger is made up of 30 amino acids that bind three base pairs. Thus, three zinc fingers can be combined to specifically recognize nine base pairs on one DNA strand and a triplet of zinc fingers is made to bind nine base pairs on the opposite strand. Each zinc finger is fused to the DNA-cutting domain of the FokI restriction endonuclease. Because FokI domains only cut DNA when they are present as dimers, a ZFN monomer binding to a chromosome cannot induce a DNA break [32], instead requiring ZFN heterodimers for sequence-specific chromosome breaks. It is estimated that 1 in every 500 genomic base pairs can be cleaved by ZNFs [33]. Compared with meganucleases, ZFNs are easier to construct because of publicly available resources [34]. Additionally, the value of ZFNs in mouse and rat genome engineering was demonstrated in several studies that produced knockout, knockin, and floxed (described below) animal models [35,36,37]. The development of transcription activator-like effector nucleases (TALENs) followed after ZFN technology [38]. TALENs are made up of tandem repeats of 34 amino acids. The central amino acids at positions 12 and 13, named repeat variable di-residues (NVDs), determine the base to which the repeat will bind [38]. To achieve a specific chromosomal break, 15 TALE repeats assembled and fused to the FokI endonuclease domain (TALEN monomer) are required. Thus, one TALEN monomer binds to 15 base pairs on one DNA strand, and a second TALEN monomer binds to bases on the opposite strand [38]. When the FokI endonuclease domains are brought together, a double-stranded DNA break occurs. In this way, a TALEN heterodimer can be used to cause a sequence-specific chromosome break. It has been estimated that, within the entire genome, TALENs have potential target cleavage sites every 35 bp [39]. Compared with ZFNs, TALENs are easier to construct with publicly available resources [40,41], and TALENs have been adopted for use in mouse and rat genome engineering in several laboratories that have produced knockout and knockin animal models [42,43,44,45,46].

The efficiencies of producing specific double-strand chromosome breaks, using prior technologies such as meganucleases, ZFNs, and TALENs [28,32,38], were surpassed when CRISPR/Cas9 technology was shown to be effective in mammalian cells [47,48,49]. The essential feature that all of these technologies have in common is the production of a chromosome break at a specific location to facilitate genetic modifications [50]. In particular, the discovery of bacterial CRISPR-mediated adaptive immunity, and its application to genetic modification of human and mouse cells in 2013 [47,48,49], was a watershed event to modern science. Moreover, the introduction of CRISPR/Cas9 methodology has revolutionized transgenic mouse generation. This paradigm shift can be seen by changes in demand for nucleic acid microinjections into zygotes, and ES cell microinjections into blastocysts at the University of Michigan Transgenic Core (Figure 1). While previously established principles of genetic engineering using mouse ES cell technology [51,52,53] remain applicable, CRISPR/Cas9 methodologies have made it much easier to produce genetically engineered model organisms in mice, rats, and other species [54,55]. Herein, we discuss principles in genetic engineering for the design and characterization of targeted alleles in mouse and rat zygotes, or in cultured cell lines, for the production of animal and cell culture models for biomedical research.

Recent trends in nucleic acid microinjection in zygotes, and embryonic stem (ES) cell microinjections into blastocysts, for the production of genetically engineered mice at the University of Michigan Transgenic Core. As shown, prior to the introduction of CRISPR/Cas9, the majority of injections were of ES cells, to produce gene-targeted mice, and DNA transgenes, to produce transgenic mice. After CRISPR/Cas9 became available, adoption was slow until 2014, when it was enthusiastically embraced, and the new technology corresponded to a reduced demand for ES cell and DNA microinjections.

There are many types of genetic modifications that can be made to the genome. The ability to specifically target locations in the genome has expanded our ability to make changes that include knockouts (DNA sequence deletions), knockins (DNA sequence insertions), and replacements (replacement of DNA sequences with exogenous sequences). Deletions in the genome can be used to knockout gene expression [56,57]. Short deletions in the genome can be used to remove regulatory elements that knockout gene expression [58], activate gene expression [59], or change protein structure/function by changing coding sequences [60].

Insertion of new genomic information can be used to knock in a variety of genetic elements. Knockins are also powerful approaches for modifying genes. Just as genomic deletions can be used to change gene function, knockins can be used to block gene function by inserting fluorescent reporter genes such as eGFP or mCherry, in such a way as to knock out the gene at the insertion point [61,62]. It is also possible to knock in fluorescent protein reporter genes, without knocking out the targeted gene [63,64]. Just as fluorescent proteins can be used to label proteins and cells, short knockins of epitope tags in proteins can be used to label proteins for detection with antibodies [64,65].

Replacement of DNA sequences in the genome can be used to achieve two purposes at the same time, such as blocking gene function, while activating the function of a new gene such as the lacZ reporter [66]. Large-scale sequence replacements are possible with mouse ES cell technology, such as the replacement of the mouse immunoglobulin locus with the human immunoglobulin locus to produce a humanized mouse [67]. Furthermore, very small replacements of single nucleotides can be used to model point mutations that are suspected of causing human disease [68,69,70].

A special type of DNA sequence replacement is the conditional allele. Conditional alleles permit normal gene expression until the site-specific Cre recombinase removes a loxP-flanked critical exon to produce a floxed (flanked by loxP) exon. Cre recombinase recognizes 34 bp loxP (locus of recombination) elements, and catalyzes recombination between the two loxP sites [71,72]. Therefore, deletion of the critical exon causes a premature termination codon to occur in the mRNA transcript, triggering its nonsense-mediated decay and failure to make a protein [13,73]. Engineering conditional alleles was the approach used by the international knockout mouse project [13]. Mice with cell- and tissue-specific Cre recombinase expression are an important resource for the research community [74].

Other site-specific recombinases, such as FLP, Dre, and Vika, that work on the same principle have also been applied to mouse models [75,76,77,78,79,80]. Recombinase knockins can be designed to knock out the endogenous gene or preserve its function [81,82]. A variation in the conditional allele is the inducible allele, which is silent until its expression is activated by Cre recombinase [79]. For example, reporter models can activate the expression of a fluorescent protein [83], change fluorescent reporter protein colors from red to green [84], or use a combinatorial approach to produce up to 90 fluorescent colors [85]. Another type of inducible allele is the FLEX allele. FLEX genes are Cre-dependent gene switches based on the use of heterotypic loxP sites [86]. In one application that combined Cre and FLP recombinases, it was demonstrated that a gene inactivated in ES cells by a gene trap could be switched back on and then switched off again [87]. In another application of heterotypic loxP sites in mouse ES cells, it was demonstrated that genes could be made conditional by inversion (COIN) [88]. This application has been used to produce mice with conditional genes for point mutations [89] and has been applied to produce conditional single exon genes that lack critical exons by definition [90].

The central principle of gene targeting with CRISPR/Cas9, or other directed DNA endonucleases, is that a double-strand DNA break is generated in the cell of interest. Following a chromosomal break, the principal outcomes of interest are nonhomologous end joining (NHEJ) repair [91] or homology-directed repair (HDR) [92]. When the break is directed to a coding exon in a gene, the outcome of NHEJ is usually a small insertion or deletion of DNA sequence at the break (indel), causing frame shifts in mRNA transcripts that lead to premature termination codons, causing nonsense-mediated mRNA decay and loss of protein expression [73]. The HDR pathway copies a template during DNA repair, and thus the insertion of modified genetic sequences in the form of a DNA donor. This DNA donor can introduce new information into the genome flanked by homology arms on either side of the chromosome break. Typical applications of HDR include the use of genetic engineering to abrogate gene expression (gene knockouts), to modify amino acid codons (i.e.; point mutations), to replace genes with new genes (e.g.; knockins of fluorescent reporters, Cre recombinase, cDNA coding sequences), to produce conditional genes (floxed genes that are normally expressed until they are inactivated by Cre recombinase), to produce Cre-inducible genes (genes that are only expressed after Cre recombinase activates them), and to delete DNA from chromosomes (e.g.; delete regulatory elements that control gene expression, delete entire genes, or delete up to a megabase of chromosome segments). The simplest of these modifications is abrogation of gene expression. Multifunctional alleles, such as FLEX alleles, require the cloning or synthesis of multi-element plasmid DNA donors for HDR.

The processes of CRISPR/Cas9-mediated modifications of genes (gene editing) to produce a new cell line or animal model have in common a series of steps to achieve the final product. First, a gene of interest is identified and the final desired allele is specified. The next step is to identify single guide RNA(s) (gRNAs) that will be used to target a chromosomal break in one or more places. There are numerous online websites that can be used for this purpose [93]. One of the most up-to-date and versatile sites is CRISPOR (http://crispor.tefor.net) [94]. Interestingly, the authors provide evidence that the predictive powers of algorithms vary depending on whether they were based on the analysis of gRNAs delivered as RNA molecules, versus gRNAs delivered as U6-transcribed DNA molecules [94]. In any event, the selection of a gRNA target (20 nucleotides), adjacent to a protospacer-adjacent motif (PAM; NGG motif), should not be done without the aid of a computer algorithm that minimizes the possibility of off-target hits. After a gRNA target is identified, a decision is made to obtain gRNAs. While it is possible to produce in vitro-transcribed gRNAs, this may be inadvisable in so much as in vitro-transcribed RNAs can trigger innate immune responses and cause cytotoxicity in cells [95]. Chemically synthesized gRNAs using phosphorothioate modifications that improve gRNA stability may be preferable alternatives to in vitro-transcribed molecules [96,97]. With a gRNA in hand, a Cas9 protein is then selected. There are numerous forms of Cas9 that can be used for different purposes [98]. For practical purposes, we limit our discussion to Cas9 varieties that are on the market. A number of commercial entities sell wild-type Cas9 protein. When wild type Cas9 is used to target the genome with nonspecific guides, the frequency of off-target genomic hits, besides the desired Cas9 target, is very likely to increase [94,99]. Alternatives to the wild-type protein include enhanced specificity Cas9 from Sigma-Aldrich [100], and high-fidelity Cas9 from Integrated DNA Technologies [101]. In addition, there are other versions such as HF1 Cas9 [102], hyperaccurate Cas9 [103], and evolved Cas9 [104], all available in plasmid format from Addgene.org. As may be inferred from the names of these engineered Cas9 versions, they are designed to be more specific than wild type Cas9. Once the gRNAs and Cas9 protein are on hand, then it is a simple matter to combine them and deliver them to the target cell to produce a chromosome break and achieve a gene knockout by introducing premature termination codons or DNA sequence deletion of regulatory regions or entire genes.

The most challenging type of genetic engineering is the insertion (i.e.; knockin) of a long coding sequence to express a fluorescent reporter protein, Cre recombinase, or conditional allele (floxed gene). In addition to these genetic modifications, numerous other types of specialized reporters can be introduced, each designed to achieve a different purpose. There is great interest in achieving rapid and efficient gene insertions of reporters in animal models with CRISPR/Cas9 technology. It is generally recognized that, the longer the insertion, the less efficient it is to produce a knockin animal. Additional challenges are allele-specific differences that affect efficiency. For example, it is fairly efficient to produce knockins into the genomic ROSA26 locus in mice, while other loci are targeted less efficiently, and thus refractory to knockins. This accessibility to CRISPR/Cas9 complexes mirrors observations in mouse ES cell gene targeting technology, in which it was reported that some genes are not as efficiently targeted as others [105].

When the purpose of the experiment is to specifically modify the DNA sequence by changing amino acid codons, or introducing new genetic information, then a DNA donor must be delivered to the cells with Cas9 reagents. After the selected gRNAs and Cas9 proteins are demonstrated to produce the desired chromosome break, the DNA donor is designed and procured. The donor should be designed to insert into the genome such that it will not be cleaved by Cas9, usually by mutating the PAM site. The DNA donor may take the form of short oligonucleotides (<200 nt) [106,107], long single-stranded DNA molecules (>200 nt) [108], or double-stranded linear or circular DNA molecules of varying lengths [109,110].

DNA donor design principles should include the following: (1) nucleotide changes that prevent CRISPR/Cas9 cleavage of the chromosome, after introduction of the DNA donor; (2) insertion of restriction enzyme sites unique to the donor, to simplify downstream genotyping; (3) insertions of reporters or coding sequences, at least 1.5 kb in length, that can be introduced as long single-stranded DNA templates with short 100 base pair arms of homology [111], or as circular double-stranded DNA plasmids with longer (1.5 or 2 kb) arms of homology [63,110]; and (4) insertions of longer coding sequences, such as Cas9, that use circular double-stranded DNA donors with longer arms of homology [63,112]. It is also possible to use linear DNA fragments as donors [63,110,113], although random integration of linear DNA molecules is much higher than those of circular donors, thus requiring careful quality control.

The establishment of genetically modified mouse and rat models can be divided into three phases, after potential founder animals are born from CRISPR/Cas9-treated zygotes. In the first phase, animals with genetic modifications are identified. The first phase requires a sensitive and specific genotyping assay to identify cells or animals harboring the desired knockin. Genotyping potential founder mice for knockins typically begins with a PCR assay using a primer that recognizes the exogenous DNA sequence and a primer in genomic DNA outside of the homology arm in the targeting vector. Accordingly, PCR assays are designed to specifically detect the upstream and downstream junctions of the inserted DNA in genomic DNA. Subsequent assays may be used to confirm that the entire exogenous sequence is intact. Conditional genes represent a special case of insertion, as PCR assays designed to detect correct insertion of loxP-flanked exons will also detect genomic DNA [108]. In the second phase, founders are mated and G1 pups are identified that inherited the desired mutation [114]. In the third phase, it is essential to sequence additional genomic regions upstream and downstream of the inserted targeting vector DNA, because Cas9 is very efficient at inducing chromosomal breaks, but has no repair function. Thus, it is not unusual to identify deletions/insertions that flank the immediate vicinity of the Cas9 cut site or inserted targeting vector DNA sequences [115,116]. If such deletions affect nearby exons, gene expression can be disrupted, and confounding phenotypes may arise.

For gene knockouts, PCR amplicons from primers that span the chromosome break site are analyzed by DNA sequencing. Any animals that are wild-type at the allele are not further characterized or used, so as to prevent any off-target hits from entering the animal colony or confounding phenotypes. Animals that show disrupted DNA sequences at the Cas9 cut site are mated with wild-type animals for the transmission of mutant alleles that produce premature termination codons, for gene knockout models [57,73]. As founders from Cas9-treated zygotes are genetic mosaics [55,115], it is essential to mate them to wild-type breeding partners, such that obligate heterozygotes are produced. In the heterozygotes, the wild-type sequence and the mutant sequence can be precisely identified by techniques such as TOPO TA cloning (Invitrogen, CA, USA) or next-generation sequencing (NGS) methods [117,118,119,120]. Animals carrying a defined indel, with the desired properties, are then used to establish lines for phenotyping. The identical approach is used when short DNA sequences are deleted by two guide RNAs [58]. Intercrossing mosaic founders will produce offspring carrying two different mutations with different effects on gene expression. These animals are not suitable for line establishment.

CRISPR/Cas9 gene editing in immortalized cell lines presents a set of challenges unique from those used in the generation of transgenic animals. Cell lines encompass a wide range of characteristics, resulting in each line being handled differently. Some of these characteristics include phenotype heterogeneity, aberrant chromosome ploidy, varying growth rates, DNA damage response efficiency, transfection efficiency, and clonability. While the principles of CRISPR/Cas9 experimental design, as stated above, remain the same, three major considerations must be taken into account when using cell lines: (1) copy number variation, or the number of alleles of the gene of interest; (2) transfection efficiency of the cell line; and (3) clonal isolation of the modified cell line. In cell lines, all alleles need to be modified in the generation of a null phenotype, or in the creation of a homozygous genotype. Unlike transgenic animals, where single allele gene edits can be bred to homozygosity, CRISPR/Cas9-edited cells must be screened for homozygous gene edits. Copy number variations within the cell line can decrease the efficiency and add labor and time (i.e.; editing 3 or 4 copies versus editing 1 or 2). Furthermore, an aberrant number of chromosomes, deletions, duplications, pseudogenes, and repetitive regions complicate genetic backgrounds for PCR analysis of the CRISPR edits. To help with some of these issues, one common approach is to use NGS on all the clonal isolates for a complete understanding of copy number variations for each clonal cell line generated, and the exact sequence for each allele.

As all cell types are not the same, different CRISPR/Cas9 delivery techniques may need to be tested to identify which method works best. One approach is to use viruses or transposons to deliver CRISPR/Cas9 reagents (detailed below). However, the viruses and transposons themselves will integrate into the genome, as well as allowing long-term expression of CRISPR/Cas9 in the cell. This prolonged expression of gRNAs and Cas9 protein may lead to off-target effects. Moreover, transfection and electroporation can have varying efficiencies, depending on the cell lines and the form of CRISPR/Cas9 reagents (e.g.; DNA plasmids or ribonucleoprotein particles (RNPs)).

Following delivery, clonal isolation is required to identify the edited cell line, and at times, can result in the isolation of a cell phenotype different than that expected, arising from events apart from the desired gene edit. While flow cytometry can aid in isolating individual cells, specific flow conditions, such as pressure, may require adjustment to ensure cell viability. Furthermore, one clonal isolate from a cell line may possess a different number of alleles for the targeted gene than another clonal isolate. Additionally, not all cell lines will grow from a single cell, thus complicating isolation. Growth conditions and cell viability can also change when isolating single cells.

Despite these challenges, new advances in CRISPR technology can likely alleviate some of these difficulties when editing cell lines. For example, fluorescently tagged Cas9 and RNAs help to isolate only transfected cells, which helps to eliminate time wasted on screening untransfected cells. Cas9-variants that harbor mutations that only create single-strand nicks (Cas9-nickases) complexed with two different, but proximal gRNAs can increase HDR-mediated knockin [48,121]. Similarly, fusing Cas9 with base-editing enzymes can also increase the efficiency of editing, without causing double-strand breaks [121].

Viral and transposon vectors have been engineered to be safe, efficient delivery systems of exogenous genetic material into cells. The natural lifecycle of some viruses and transposons includes the stable integration into the host genome. In the field of genome engineering, these vectors can be used to modify the genome in a non-directed fashion, by inserting cassettes expressing any cDNA, shRNA, miRNA, or any non-coding RNA. The most widely used vectors capable of integrating ectopic genetic material into cells are retroviruses, lentiviruses, and adeno-associated virus (AAV). These viruses are flanked by terminal repeats that mark the boundaries of the integration. In engineering these viruses into recombinant vector systems, all the viral genes are removed from the flanking terminal repeats and supplied in trans for the recombinant virus to be packaged. These gutted, nonreplicable viral vectors allow for the packaging, delivery, integration, and expression of cDNAs of interest, shRNAs, and CRISPR/Cas9, without viral replication in various biological targets.

Similar to recombinant viruses, transposon vectors are also gutted, separating the transposase from the terminal repeat-flanked genetic material to be inserted into the genome. DNA transposons are mobile elements (jumping genes) that integrate into the host genome through a cut-and-paste mechanism [122]. Transposons, much like viral vectors, are flanked by repeats that mark the region to be transposed [123]. The enzyme transposase binds the flanking DNA repeats and mediates the excision and integration into the genome. Unlike viral vectors, transposons are not packaged into viral particles, but form a DNA-protein complex that stays in the host cell. Thus, the transgene to be integrated can be much larger than the packaging limits of some viruses.

Two transposons, Sleeping Beauty (SB) and piggybac (PB), have been engineered and optimized for high activity for generating transgenic mammalian cell lines [124,125,126]. Sleeping Beauty is a transposable element resurrected from fish genomes. The SB system has been used to generate transgenic HeLa cell lines, T-cells expressing chimeric antigen receptors that recognize tumor-specific antigens, and transgenic primary human stem cells [127,128,129]. The insect-derived PB system also has been used to generate transgenic cell lines [126,130,131]. The PB system was used to generate induced pluripotent stem cells (iPSCs) from mouse embryonic fibroblasts, by linking four or five cDNAs of the reprogramming (Yamanaka) factors [132] with intervening peptide self-cleavage (P2A) sites, thus delivering all of the factors in one vector [130]. Furthermore, once reprogrammed, the transgene may be removed by another round of PB transposase activity, leaving no genetic trace of integration or excision (i.e.; transgene-free iPSCs). Following PB transposase activity, epigenetic differences remaining at the endogenous promoters of the reprogramming factor genes result in sustained expression and pluripotency, despite transgene removal.

Aside from transgene insertion, Sleeping Beauty (SB) and piggyback (PB) have both been engineered to deliver CRISPR/Cas9 reagents into cells [133,134,135]. Similar to lentivirus, the stable integration of CRISPR/Cas9 by transposons could increase the efficacy of targeting and modifying multiple alleles. SB and PB have been used to deliver multiple gRNAs to target multiple genes (instead of just one), aiding in high-throughput screening. Furthermore, owing to the nature of PB excision stated above, the integrated CRISPR/Cas9 can be removed once a clonal cell line is established, to limit off-target effects. However, engineered transposons must be transfected into cells. As stated above, efficiencies vary between different cell lines and transfection methods. One potential solution to overcome this challenge is to merge technologies. For example, instead of transfecting cells with a plasmid harboring a gRNA flanked by SB terminal repeats (SB-CRISPR), the SB-CRISPR may be flanked by recombinant AAV (rAAV) terminal repeats (AAV-SB-CRISPR), allowing for packaging into rAAV. To that end, rAAV-SB-CRISPR has been used to infect primary murine T-cells, and deliver the SB-CRISPR construct [136].

Retroviruses are RNA viruses that replicate through a DNA intermediate [137]. They belong to a large family of viruses including both onco-retroviruses, such as the Moloney murine leukemia virus (MMLV) (simply referred to as retrovirus), and lentiviruses, including human immunodeficiency virus (HIV). In all retroviruses, the RNA genome is flanked on both sides by long terminal repeats (LTRs); packaged with viral reverse transcriptase, integrase, and protease, surrounded by a protein capsid; and then enveloped into a lipid-based particle [138]. Envelope proteins interact with specific host cell surface receptors to mediate entry into host cells through membrane fusion. Then, the RNA genome is reverse-transcribed by the associated viral reverse transcriptase. The proviral DNA is then transported into the nucleus, along with viral integrase, resulting in integration into the host cell genome [139]. By contrast, the retroviral MMLV pre-integration complex is incapable of crossing the nuclear membrane, thus requiring the cell to undergo mitosis to gain access to chromatin [139], while lentiviral pre-integration complexes can cross nuclear membrane pores, allowing genome integration in both dividing and non-dividing cells.

Large-scale assessments of genomic material composition have uncovered features associated with retroviral insertion into mammalian genomes [140]. Although determination of integration target sites remains ill-defined, it does depend on both cellular and viral factors. For retroviruses such as MMLV, integration is preferentially targeted to promoter and regulatory regions [140,141,142]. Such preferences can be genotoxic owing to insertional activation of proto-oncogenes in patients undergoing gene therapy treatments for X-linked severe combined immunodeficiency [143,144], WiskottAldrich syndrome [143], and chronic granulomatous disease [145]. Likewise, retroviral integration can generate chimeric and read-through transcripts driven by strong retroviral LTR promoters, post-transcriptional deregulation of endogenous gene expression by introducing retroviral splice sites (leading to aberrant splicing), and retroviral polyadenylation signals that lead to premature termination of endogenous transcripts [142,146,147].

Unlike retroviruses, lentiviruses prefer to integrate into transcribed portions of expressed genes in gene-rich regions, distanced from promoters and regulatory elements [140,142,148]. The cellular protein LEDGF/p75 aids in the target site selection by binding directly to both the active gene and the viral integrase within the HIV pre-integration complex [149]. Although the propensity of lentivirus to integrate into the body of expressed genes should increase the incidence of post-transcriptional deregulation, deletion of promoter elements from the lentiviral LTR (self-inactivating (SIN) vectors) has been reported to decrease transcriptional termination, but increase the generation of chimeric transcripts [149]. Overall, it appears that lentiviral SIN vectors are less likely to cause tumors than retroviral vectors with an active LTR promoter [148,150,151,152].

The 7.510 kb packaging limit of lentiviruses can accommodate the packaging, delivery, and stable integration of Cas9 cDNA, gRNAs, or Cas9 and gRNAs (all-in-one) to cells [153,154]. Often, a selectable marker, such as drug resistance, can also be included to isolate transduced cells. The high transduction efficiency of lentivirus can result in an abundance of CRISPR/Cas9-expressing cells to screen, compared with more traditional transfection methods. Stable and prolonged expression of CRISPR/Cas9 can facilitate targeting of multiple alleles of the gene of interest, resulting in more cells harboring homozygous gene modifications. Conversely, stable integration of CRISPR/Cas9 increases potential off-target effects. Moreover, lentiviral integration itself is a factor that may confound cellular phenotypes and should be considered when characterizing CRISPR-edited cell lines.

Adeno-associated virus (AAV) is a human parvovirus with a single-stranded DNA genome of 4.7 kb, which was originally identified as a contaminant of adenoviral preparations [155]. The genome is flanked on both sides by inverted terminal repeats (ITR) and contains two genes, rep and cap [156,157]. Different capsid proteins confer serotype and tissue-specific targeting of distinct AAVs, in vivo. AAV cannot replicate on its own, and requires a helper virus, such as adenovirus or herpes simplex virus (HSV), to provide essential proteins in trans. AAV is the only known virus to integrate into the human genome in a site-specific manner at the AAVS1 site on chromosome 19q13.3-qter [158,159,160]. Although the precise mechanism is not well understood, the Rep protein functions to tether the virus to the host genome through direct binding of the AAV ITR and the AAVS1 site [158,160,161]. In the recombinant AAV (rAAV) vector system, the rep and cap genes are removed from the packaged virus, resulting in the loss of site-specific integration into the AAVS1 site. Despite removal of Rep, it has been shown that rAAV can still integrate, albeit randomly, into the host genome, via nonhomologous recombination, at low frequencies [162,163,164]. Furthermore, numerous clinical trials, to date, have shown that rAAV integration is safe and has no genotoxicity [165,166,167]. However, this safety is controversial, owing to preclinical studies suggesting genotoxicity in mouse models [168,169,170,171]. More studies are needed to understand the cellular impact of rAAV integration.

rAAVs have been used to deliver one or two CRISPR guide RNAs (gRNAs), in cells and model animals, by taking advantage of different rAAV serotypes to target specific cells or tissue types. Owing to the packaging capacity of rAAV, SpCas9 must be delivered as a separate virus, unlike lentivirus, which can be delivered as an all-in-one CRISPR/Cas9 vector. However, alternate, smaller Cas9s can be packaged into rAAVs [172]. Furthermore, rAAVs can be used to deliver repair templates or single-stranded donor oligonucleotides (ssODNs) for homology-directed repair (HDR), relying on the single-stranded nature of the AAV genome [173,174]. It has also been observed that rAAVs can integrate into the genome at CRISPR/Cas9-induced breaks in various cultured mouse tissue types, including neurons and muscle [175]. This observation goes against the notion of rAAVs integrating only at the AAVS1 locus, and should be considered when analyzing and characterizing rAAV-mediated CRISPR-edited cells.

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Principles of Genetic Engineering - PubMed Central (PMC)

The next 'big thing' in genetically modified crops: Drought-tolerant and herbicide resistant wheat. Here's what you need to know  Genetic Literacy Project

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The next 'big thing' in genetically modified crops: Drought-tolerant and herbicide resistant wheat. Here's what you need to know - Genetic Literacy...

Genetic engineering and biotechnology: The future of food is here  Yourweather.co.uk

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Genetic engineering and biotechnology: The future of food is here - Yourweather.co.uk

Scientists Just Achieved a Major Milestone in Creating Synthetic Life  Yahoo! Voices

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Scientists Just Achieved a Major Milestone in Creating Synthetic Life - Yahoo! Voices

Two males give birth to child in incredible science experiment; the baby is now an adult | Mint  Mint

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Two males give birth to child in incredible science experiment; the baby is now an adult | Mint - Mint

Definition

Genetic engineering or genetic modification is a field of genetics that alters the DNA of an organism by changing or replacing specific genes. Used in the agricultural, industrial, chemical, pharmaceutical, and medical sectors, genetic engineering can be applied to the production of brewing yeasts, cancer therapies, and genetically-modified crops and livestock, among countless other options. The only criterion is that the modified product is or once was a living organism that contains DNA.

Examples of genetic engineering are listed according to sector in this article, where each sector applies DNA modification with a different goal. As the human genome contains between 20,000 and 25,000 genes and as these genes can extend from just a few hundred base pairs to over 2 million, the scope of genetic engineering is huge. However, there are lots of ethical questions that concern how far this kind of research should go and what applications are acceptable.

The chemical industry uses genetic engineering when it produces modified live microorganisms for chemical production. It is not possible to genetically engineer a chemical or material like an acid or a steel bar they do not contain DNA; however, bacteria that produce acid, for example, can be genetically modified.

Natural chemical compounds are essential for the existence of life. These have been mimicked over the years by man-made (synthetic) copies. One example of genetic engineering in todays chemical industry is an enzyme called protease. Protease engineering is the foundation of genetic modification in laundry detergent manufacturing.

Proteases are enzymes found in every living organism; their function is to catalyze (speed up) the breakdown of ester and peptide bonds that are found in many types of laundry stains. Protease genes give cells the manufacturing instructions for protease production inside the cell (protein synthesis). By manipulating these genes, we can change the ultimate form of the protease and some of its characteristics.

Earlier detergents did not have access to genetic-engineering technology but even then, researchers were able to modify proteases by selecting and producing the best strains. With genetic engineering, these enzymes can be further improved for even whiter whites. Once the gene for protease production was decoded it was possible to extract and modify it. Many modifications have been made that improve stain-removing results in varying pH and water temperature levels, for example.

Other genetic engineering examples in the chemical industry include less environmentally-damaging wastewater management. This involves modifying the genes of the many types of bacteria that digest waste without their leaving behind similarly harmful byproducts. Another example is manufacturing biodegradable plastics using genetically-modified strains of cyanobacteria.

Genetic engineering examples relating to crop production are often used to tell us why not to buy or eat them; however, a growing population without the time, space, or often the knowledge to produce crops at home means we need to use our agricultural land more efficiently. At the same time, it is important not to reduce natural habitats around the world. Genetically-modified (GM) crops are an answer in the form of increased crop yield on a smaller plot. Genetically modifying a crop concentrates on increased resistance to disease, increased fiber and nutrient content, or increased yield preferably a combination of all three. If we can obtain all the minerals and vitamins we need from a super-tomato that grows very quickly without needing pesticides or fertilizers, and will even grow in drought conditions, then the topic of GM crops suddenly looks very attractive indeed.

A lot of negative public comments have caused genetically modified crops to be unpopular; many GM crops even when legally grown cant find a big enough market. This means that farmers rarely want to take the financial risk to grow them.

There is no scientific evidence that a GM crop is dangerous to eat in comparison with a non-GM crop, but genetic engineering is quite new and we cant say for sure if the long-term effects are harmful to humans or the animals that eat them (that we might then eat in our burgers). The only GM crop grown legally in the European Union (EU) is MON 810 maize. Production of this maize in the EU might also be banned in the future. Federal law in the US is strict concerning GM testing but the production, sale, and consumption of GM crops are legal.

Genetic engineering examples in livestock rearing should always mention one Food and Drug Administration restriction that has recently been lifted. The import, sale, and raising of GM salmon eggs used to be banned in the US, although this wasnt due to fears that eating these fish could be dangerous to our health the ban was due to labeling laws. This ban has now been lifted.

In AquaAdvantage salmon, scientists combined the genes of Chinook salmon and the rather ugly ocean pout (below) to produce a continuously-growing salmon (salmon usually grows seasonally) that uses and requires fewer calories than wild or farmed alternatives. The company has spent twenty years testing this new food source; arguments against GM salmons use are usually based on the fact that twenty years is not very long in the average human lifespan.

While genetically modified beef is hard to find, it is still possible that your pot roast once ate GM feed. It might also have when alive been injected with genetically-engineered recombinant bovine growth hormone (rBGH). This hormone is also injected into dairy cows. It has been reported that milk from rBGH-treated cows contains higher levels of IGF-1, a hormone that seems to increase breast, prostate, colon, and lung cancer risk in humans. This is just one of the reasons why GM products are so controversial. But studies have also shown that the use of GM feeds increases health levels in animals and often means that farmers do not need to inject antibiotics and hormones into their livestock as these chemicals can pass into the bloodstreams of the people that eat the livestock or drink their milk, this can be a doubly positive result. The jury is still out.

GM chicken is not available in your local supermarket (yet) but chickens fed with GM feeds are often labeled as such. So it is the digested residues of different genetically modified crops and not a genetically modified bird that is roasting in the oven.

Genetically modified chicken eggs are being studied as a future source of natural chemical compounds. Female chickens can be genetically engineered to produce eggs that contain larger amounts of certain proteins. These proteins are commonly used in the manufacturing processes of pharmaceutical drugs. Future drug prices could be much more affordable thanks to genetic modification technology.

Genetic engineering examples in cancer therapy are already starting to show very positive results. The chicken egg makes an appearance here, too. In this field of genetic engineering, bacterial genes that produce particular proteins are modified. These proteins you might have heard of the very heavily studied Cas9 protein form antibodies that help to destroy viruses. This type of protein also supports a mechanism that alerts the immune response in humans. As this response is often suppressed by cancer cells, Cas9 might be able to help the body to recognize and then fight cancer. Cas9 is already being studied and trialed for genetic disorders such as sickle cell disease and cystic fibrosis.

Hereditary diseases and disorders might become a thing of the past thanks to genetic engineering there is just one problem, the ethical use of human embryos for research purposes.

Embryological genetic engineering is legal in some countries and these countries are given a lot of criticism. But when He Jiankui edited the genes of twin embryos and then had them implanted in a female who gave birth to these genetically-modified children, the world went crazy and Jiankui was subsequently jailed. Not only are the long-term effects of genetic engineering unknown, but any changes might carry through to subsequent generations or continue to change without the natural control that is evolution. For people who believe that life begins at conception or consider an embryo a living, conscious person, there are even more ethical arguments.

Many parents who undergo the process of in vitro fertilization (IVF) are offered the option of pre-implantation genetic diagnosis (PGD). This checks the DNA of the fertilized egg before it is inserted into the womb. The aim is to source possible genetic mutations. The parents are allowed to discard faulty eggs. Many believe that this is very wrong as we have not agreed on what is considered an undesired mutation. A genetic fault that causes miscarriage would be acceptable, perhaps. But what about gender, hereditary mental illness, eye color? In the past years, several fertility clinics in India have been called out for promising male offspring to couples, for example. This is not an example of genetic engineering, but many groups fear that certain physiological choices may edge their way into genetic engineering without being controlled. Today, genetic modification in humans follows practically the same ethical arguments as abortion.

The pros and cons of genetic engineering are not at all clear-cut. In the field of human genetic modification, our personal beliefs affect how this technology will develop and move forward. In countries where the law states that human life begins at week 24, the genetic engineering of embryos not carried to term is more likely to be accepted. This ethical question is part of what is known as the fetal personhood argument and is the main reason why genetic engineering in humans is meeting so much resistance.

In an agricultural setting, the publics fears concern the long-term effects of eating GM foods. These fears stop farmers from producing modified crops as they might not be able to sell them and, in many countries, it is unlawful to grow them. Personal issues are often opinions; the actual pros and cons concern the results of long-term scientific research. Unfortunately, genome editing is a new technology and we do not have any data that covers more than a few years certainly nothing that covers the lifetimes of one or more generations.

Genetic engineering pros should start with the fact that this topic has allowed us to learn so much more about our genes and the genes of other organisms. It is thanks to genetic engineering that we are learning how the entire range of DNA-containing organisms from bacteria to humans works.

Genetic engineering has given us fresh and unexpected knowledge that tells us how certain illnesses develop. The field has also provided targeted therapies that can cure or at least relieve these diseases. Not only the action of pharmaceuticals but also their cheaper production as in the case of GM chicken eggs can be made more efficient through this technology.

The combination of a growing global population and the need to maintain a very unstable ratio of agricultural land to natural habitats has led to the development of genetically-engineered crops. These crops are designed to have a greater yield, use fewer nutrients to grow, and require less acreage or fewer chemicals (herbicides and pesticides). Scientists can even improve taste, nutritional values, colors, and shapes.

Genetically-modified bacteria help to produce bio-fuels from genetically-modified crops. Bio-fuels reduce the effects of fossil fuel pollution. Cyanobacteria help us to produce biodegradable plastics and other GM microorganisms break down our waste. Genetic modification is strongly linked to our ecology and future.

And we use less of the earths resources when our livestock grows more quickly. When beef cattle grow to full size in one year instead of two or three, that is two years off of every animals carbon footprint. When bovine genes are modified to fight disease, our milk and meat have less antibiotic and hormone residue. Genetic engineering means less pressure to turn important, disappearing natural ecosystems into food-production factories.

The cons are mainly based on the lack of long-term studies into the effects of genetic engineering, both on an organism and on the organisms that eat it. Maybe even those that live alongside it. As with all new but potentially damaging technology, we just dont have enough data.

Another factor is that, although we have decoded the human genome, we do not know everything we need to about every function in the human body. For example, the gut microbiome is a quite recent hot topic. Scientists now accept that bacteria in the gut directly affect the brain which was rarely the case ten years ago. But exactly how the neurotransmitters of the brain interact with chemicals in the digestive tract is still a mystery. Examples like this mean that many people argue we should not try to fix something if we dont know exactly how it works, know what the long-term effects will be, or know if it is actually broken in the first place.

There are other hurdles, of course. Before knowing whether genetic engineering can safely eliminate a fatal disorder forever, we have to figure out if it is right to change the DNA of embryos, let them grow and be born, and then research their lives from birth to old age (and maybe their children and grandchildren, too) so that we can ensure the new cure is safe.

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Genetic Engineering - The Definitive Guide | Biology Dictionary

Constitutive expression of Cas9 and rapamycin-inducible Cre recombinase facilitates conditional genome editing in Plasmodium berghei  Nature.com

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Constitutive expression of Cas9 and rapamycin-inducible Cre recombinase facilitates conditional genome editing in Plasmodium berghei - Nature.com

Genetic engineering refers to scientific techniques that alter the DNA of an organism. Scientists might remove entire segments of DNA, insert additional genes from other organisms, or change one base pair. Genetic engineering can enhance, modify, or take away specific abilities of an organism to do different things. For example, scientists have engineered E. coli bacteria to mass-produce insulin, providing a vital treatment for people with diabetes.

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Funding was provided by grants from the National Science Foundation (#2223678) and Rita Allen Civic Science Fellows. This infographic was produced by the Science and Technology Policy Program and the Center for Health and Biosciences at Rice Universitys Baker Institute for Public Policy by Alicia L. Johnson. Some elements of this infographic have been created with BioRender.com.

This material may be quoted or reproduced without prior permission, provided appropriate credit is given to the author and Rice Universitys Baker Institute for Public Policy. The views expressed herein are those of the individual author(s), and do not necessarily represent the views of Rice Universitys Baker Institute for Public Policy.

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What is Genetic Engineering? - Baker Institute

Your cells are dying. All the time.  Genetic Literacy Project

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Your cells are dying. All the time. - Genetic Literacy Project

ARCUS breakthrough: An advanced gene editing tool appears to have cured an infant of an early onset metabolic disorder  Genetic Literacy Project

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ARCUS breakthrough: An advanced gene editing tool appears to have cured an infant of an early onset metabolic disorder - Genetic Literacy Project

How Genetic Modification is Changing the Future of Conservation  MSN

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How Genetic Modification is Changing the Future of Conservation - MSN

Researchers genetically engineer yeast to produce healthy fatty acid  University of Alberta

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Researchers genetically engineer yeast to produce healthy fatty acid - University of Alberta

The great gene editing debate: can it be safe and ethical?  BBC.com

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The great gene editing debate: can it be safe and ethical? - BBC.com

recombinant DNA Summary

Recombinant DNA, a segment of DNA that is generated by combining genetic material from at least two different species. Such new genetic combinations are of value to science, medicine, agriculture, and industry. A fundamental goal of genetics is to isolate, characterize, and manipulate genes.

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genetic engineering summary | Britannica

Anti-biotechnology campaigners embrace classic crops, are suspicious of hybrid varieties and claim genetic modification violates nature. Heres a primer on the differences  Genetic Literacy Project

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Anti-biotechnology campaigners embrace classic crops, are suspicious of hybrid varieties and claim genetic modification violates nature. Heres a...

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