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How does one insert cas9 into animal cells?

How does one insert cas9 into animal cells?


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How could cas9 be inserted into cells by researchers looking to edit a genome? I imagine for engineering bacterial systems you could just put in the cas9 coding region in an expression vector, but is that how it is also done for eukaryotic cells?


If you are talking about the nuclease functionality of Cas9, then you aren't adding the Cas9 gene into the genome, you are transfecting it on a plasmid. Usually there is either some form of permeabilization done to the cells, or the construct is inserted into a viral vector in order to transfect the cells in tissue culture. In multicellular animals, you would need to use the viral vector route as would need to be done with any gene therapy technique. Addgene is a nonprofit plasmid repository that has many of the CRISPR/Cas9 constructs that have been developed, and they have an excellent resource on their Website.

You wouldn't really want Cas9 active in the cell after it has done its job, as you increase your likelihood of off-target hits. The target shouldn't be in the genome anyway if the targeting event occurred, because you have either substituted in another piece of DNA or you have cut the DNA and the DNA repair mechanisms have introduced random nucleotides in order to repair the double stranded break.

Eukaryotic cells, for the most part, cannot maintain plasmids. The Cas9 gene will be transcribed along with the guide RNA. The resulting protein/RNA complex targets the sequence of interest and makes a cut. When the cell replicates the plasmid will likely be lost, but the daughter cells will inherit the genomic edit made by the targeting event.

If you are looking to insert a different gene at the target site, say a reporter into the genome, then you co-transfect a linear piece of DNA with homology arms around the cut site so that that donor will be inserted via homologous break repair.

One of the Cas9 constructs that has bene developed has disabled the nuclease activity, so that the Cas9/RNA complex ends up acting as a binding protein to suppress the activity you are looking to study without actually deleting or replacing the gene. In this case you may look to insert the Cas9 cassette, in which case you would use a recombination event as you would with any other type of incorporation into the genome.


How does CRISPR Cas9 work?

It can be used to insert genetic sequences into a locus of choice. To understand how you really need to understand gene targeting by homologous recombination, and to understand that, you need to understand homologous recombination. You can find material to cover this in more depth online but I'll try explain at the most simple level.

DNA damage is pretty bad for cells, especially double-strand breaks (DSBs) where both strands of the double helix are split. So cells really need to repair these, and in fact there's two mechanisms they have of doing so:

a) Non-homologous end joining - where they just take the two broken bits and splice them together - this isn't precise and can often lead to insertions/deletions with a possible frameshift mutation.

b) Homologous recombinantion - this is when the cell uses the homologous chromosome as a template to fix the DSB. The details of how it works aren't important but just understand that the cell looks for homologous sequences upstream and downstream of the DSB (5' and 3' homology arms respectively) and matches that to the homology arms in the template DNA and copies over everything between them in to fill in the gap and repair the DSB.

What this means then, is that we can exploit this system to insert genes simply by putting in a fake template DNA for the broken DNA to use as a repair copy. All we need are 5' and 3' homology arms but with whatever gene we want between them. This known as gene targeting:

If you have a selection marker (e.g. Antibiotic resistance), we can select for cells that have taken up the gene in that spot. But really the system as it is, relies on waiting for DSBs to happen by chance.

This is where the CRISPR/Cas9 system comes in. As you say, it is basically a programmable pair molecular of molecular scissors that can make a DSB anywhere. This is simply done by making a guide RNA that is complementary to the locus where you want to make the cut. So overall, if you inject the guide RNA with the cas9 enzyme - you'll get a DSB where you want. You also need to inject a homologous recombination repair template with homology arms to insert in your gene when the DSB tries to repair itself.


Incidentally, you can also use the CRISPR/Cas9 system in a knockout mode instead of a gene insertion mode. Here you do the same steps but don't have a repair template. Instead you want repair to occur through non-homologous end joining to create indel, and thus frameshift, mutations. This will create a premature stop codon and the mRNA product will be degraded so you've knocked that gene out.


CRISPR-Cas9 gene editing: Check three times, cut once

Bayesian color map depicting Cas9 particle movement in different regions of a mammalian cell nucleus. Red/orange denotes faster movement, and blue denotes slower movement. Cas9 movement is generally more restricted in silenced/lowly expressed regions of the genome known as heterochromatin. Credit: UC Berkeley/HHMI

Two new studies from the University of California, Berkeley, should give scientists who use CRISPR-Cas9 for genome engineering greater confidence that they won't inadvertently edit the wrong DNA.

The gene editing technique, created by UC Berkeley biochemist Jennifer Doudna and her European colleague Emmanuelle Charpentier, has taken the research and clinical communities by storm as an easy and cheap way to make precise changes in DNA in order to disable genes, correct genetic disorders or insert mutated genes into animals to create models of human disease.

The two new reports from Doudna's lab and that of UC Berkeley colleague Robert Tjian show in much greater detail how the Cas9 protein searches through billions of base pairs in a cell to find the right DNA sequence, and how Cas9 determines whether to bind, or bind and cut, thereby initiating gene editing. Based on these experiments, Cas9 appears to have at least three ways of checking to make sure it finds the right target DNA before it takes the irrevocable step of making a cut.

"CRISPR-Cas9 has evolved for accurate DNA targeting, and we now understand the molecular basis for its seek-and-cleave activity, which helps limit off-target DNA editing," said Doudna, a Howard Hughes Medical Institute investigator at UC Berkeley and professor of molecular and cell biology and of chemistry. Tjian is president of the Howard Hughes Medical Institute and a UC Berkeley professor of molecular and cell biology.

The studies also illustrate how well CRISPR/Cas9 works in human and animal cells - eukaryotes - even though "the technique was invented by bacteria to protect themselves from getting the flu," Doudna said.

CRISPR-Cas9 is a hybrid of protein and RNA - the cousin to DNA - that functions as an efficient search-and-snip system in bacteria. It arose as a way to recognize and kill viruses, but Doudna and Charpentier realized that it could also work well in other cells, including humans, to facilitate genome editing. The Cas9 protein, obtained from the bacteria Streptococcus pyogenes, functions together with a "guide" RNA that targets a complementary 20-nucleotide stretch of DNA. Once the RNA identifies a sequence matching these nucleotides, Cas9 cuts the double-stranded DNA helix.

One study, published in the Nov. 13 issue of Science, tracked Cas9-RNA molecules though the nucleus of mammalian cells as they rapidly searched through the entire genome to find and bind just the region targeted and no other.

The Cas9 enzyme must flex and bend in order to bind to the guide RNA (orange). Once the Cas9-RNA complex finds its target DNA (red), the cutting region of Cas9 (yellow) will swing into place relative to its mate (blue) only when the RNA and DNA correctly match. Only then does the enzyme cut the double-stranded DNA. These detailed movements were revealed by fluorescence experiments reported in a Nature paper from the Doudna lab. Credit: Sam Sternberg/UC Berkeley

"It's crazy that the Cas9 complex manages to scan the vast space of eukaryotic genomes," said graduate student Spencer Knight, first author of the Science paper.

Previous studies had suggested that there are many similar-looking DNA regions that Cas9 could bind and cut, which could limit its usefulness if precision were important. These off-target regions might share as few as four or five nucleotides with the 20-nucleotide primer, just enough for Cas9 to recognize.

"There is a lot of off-target binding by Cas9, but we found that these interactions are very brief - from milliseconds to seconds - before Cas9 moves on," he said.

Because these exploratory bindings - perhaps as many as 300,000 of them - are often very short-lived, a few thousand CRISPR-Cas9 complexes can scour the entire genome to find one targeted stretch of DNA. Cas9 must also recognize a short three-base-pair DNA sequence immediately following the primer sequence, dubbed PAM, which occurs about 300 million times within the human genome.

"If Cas9 bound for tens of seconds or minutes at each off-target site, it would never, ever be able to find a target and cut in a timely manner," Knight said.

Cas9's final checkpoint

The other study, published online Oct. 28 in Nature, showed that once Cas9 binds to a region of DNA, it performs another check before two distant sections of the Cas9 protein complex come together, like the blades of a scissors, to precisely align the active sites that cut double-stranded DNA.

"We found that RNA-guided Cas9 can bind some off-target DNA sequences, which differ from the correct target by just a few mutations, very tightly. Surprisingly, though, the region of Cas9 that does the cutting is inhibited because of the imperfect match. But when the correctly matching DNA is located, Cas9 undergoes a large structural change that releases this inhibition and triggers DNA cutting," said first author Samuel Sternberg, who recently received his Ph.D. at UC Berkeley. He was able to observe these changes using a fluorescently labeled version of the Cas9 complex.

"We think that this structural change is the last checkpoint, or proofreading stage, of the DNA targeting reaction," he said. "First, Cas9 recognizes a short DNA segment next to the target - the PAM - then the target DNA is matched up with the guide RNA via Watson-Crick base-pairing. Finally, when a perfect match is identified, the last part of the protein swings into place to enable cutting and initiate genome editing."

A smaller Cas9 protein from a different species of bacteria, Staphylococcus aureus, likely exploits the same strategy to improve the precision of DNA targeting, suggesting that "this important feature has been preserved throughout evolutionary time," he added.

"This is good news, in that it suggests that you have more than one checkpoint to ensure correct Cas9 binding," Knight said. "There's not just sequence regulation, there is also temporal regulation: it has to engage with the DNA and park long enough that it can actually rearrange and cut."

The discoveries from Doudna, Tjian and their teams shed light on the molecular basis of off-target effects during genome editing applications, and may guide the future design of more accurate Cas9 variants.


CRISPR/Cas9: BIOLOGY, MECHANISM OF ACTION AND CHALLENGES

CRISPR/Cas9 technique has not only shown tremendous growth in scientific research but it has also drawn much attention as a promising gene editing tool for cancer, genetic disorders, and disease-causing bacteria. Out of 13469 articles that show the word “CRISPR” as Best Match, around 5665 articles have been published in PubMed from 2018 to till now.

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The burgeoning field of CRISPR/Cas technology has surpassed other gene editing tools due to its ease to handle and low cost.

CRISPR is the short form of the name “Clustered Regularly Interspaced Short Palindromic Repeats”. The CRISPR Associated protein is shortened as a Cas protein. Therefore the technique is named as CRISPR/Cas technology in short.

The CRISPR/Cas systems naturally occur in bacteriaand archaea (Prokaryotes) to acquire immunity against viral infections and plasmids. Later, scientists started to adopt this system to edit genes in prokaryotes as well as eukaryotes.

The first sign of the CRISPR/Cas system was discovered by a Japanese research group in 1987. They identified a pattern of short repeat sequences interspersed with short, non-repetitive “spacers” in Escherichia coli genome. In 2012, two scientists (Doudna and Charpentier) programmed the CRISPR/Cas9 system to cleave specific DNA sequences. This lead to emerge CRISPR/Cas9 as a promising gene editing tool.

The CRISPR/Cas system can be classified into two classes as Class1 and Class2. Class 1 CRISPR/Cas systems employ multi Cas protein complex while Class2 CRISPR/Cas systems accomplish single Cas protein. Further two Classes are divided into six types based on the presence of specific signature genes.

The widely used CRISPR/Cas9 technique belongs to the Class 2 type II CRISPR/Cas system. The CRISPR/Cas9 system naturally occurs in Streptococcus pyogenes bacterium as an adaptive immune system to disrupt virus and plasmid which invade bacteria. In this system, a short sequence of foreign DNA (virus or plasmid) is integrated into the bacterium genome to create “identity” to recognize similar invasions prior to their infections in the future. Once a similar virus invades, the bacteria encode complementary ribonucleic acid strand (RNA) of “identity” which can bind to the complementary DNA sequence of the virus and then cleave it.

The CRISPR/Cas9 system consists of three components: crRNA (CRISPR RNA), tracrRNA (Transactivating CRISPR RNA) and Cas9 protein. This Cas9 protein shows helicase activity (unwind DNA double-strand) as well as nuclease activity (cleave DNA strand).In the bacterial CRISPR/Cas9 system, guide RNA (gRNAà crRNA + tracrRNA) directs Cas9 protein which functions as a DNA endonuclease enzyme to cleave viral DNA strands.

Figure 1: Classification of CRISPR/Cas System

Instead of two separate RNA molecules, researchers have synthesized one RNA molecule (sgRNA- single guide RNA) which can be used as a similar molecule to crRNA+tracrRNA.It has simplified three component CRISPR/Cas9 system to the two-component system (sgRNA+Cas9).

MECHANISM OF ACTION

The defense mechanism of the CRISPR/Cas9 system in bacteria can be divided into three phases:

Each phase is described below with a diagram.

After the viral infection, cas operon produces (transcription followed by translation) cas1-cas2 protein complex which can identify protospacer/spacer sequence (“identity”) of viral DNA and integrates it to the CRISPR array flanked by repeat sequences.

Later, the tracr gene and CRISPR array transcribe (convert gene into respective RNA molecule) tracrRNA and pre-crRNA respectively whereas the cas9 gene transcribes and translates (convert RNA molecule into respective protein) into cas9 protein.

Figure 2: Spacer acquisition of CRISPR/Cas9 system

The tracrRNA anneals to the pre-crRNA repeat. Then tracrRNA: pre-crRNA duplex binds to cas9 protein. Further, the complex recruits RNaseIII enzyme that cleaves the pre-crRNA at the repeat. Finally, an unknown nuclease trims 5’ repeat-derived portion of the crRNA by leaving 20 nucleotide long spacer sequence for the final phase. This forms the final CRISPR/Cas9 complex for interference.

Figure 3: crRNA processing of CRISPR/Cas9 System

The Cas9 of the effector complex identifies target DNA sequence through PAM (Protospacer Adjacent Motif) recognition that locates in the non-target DNA strand (The PAM sequence which recognized by Streptococcus pyogenes is 5’NGG 3′). The 20 nucleotides long, spacer sequence of mature crRNA binds to the target DNA strand in viral DNA. It allows the cas9 protein to activate its two nuclease domains, HNH and RuvC. These two domains HNH and RuvC cleave target DNA strand and non-target DNA strand of virus respectively, by generating a blunt, double-strand break, 3 base pairs front of the PAM. This leads to the destruction of the viral genome.

Figure 4: Interference of CRISPR/Cas9 System

Bacteria employ the CRISPR/Cas9 system to protect them from invasions. Nevertheless, the same system can be harnessed to edit mammalian genome including humans. Unlike bacteria, it gives rise to double-strand breaks in DNA which are later repaired either by non-homologous end joining or homology-directed repair mechanism in mammalian cells. This garnered much attention from the scientific community to be employed in this state of the art technique to eradicate human diseases such as genetic blood disorders, neurodegenerative diseases, and cancers. Up to date, several clinical trials have been recruited to employ the CRISPR/Cas9 technique to treat cancer and beta thalassemia according to ClinicalTrials.gov database.

CHALLENGES

Even though the CRISPR/Cas9 system holds immense promise to treat human diseases, it faces technical challenges.

The CRISPR/Cas9 system naturally occurs in Streptococcus pyogene bacterium which is harmful to human health. Once it delivers to human cells, the human body can create immunogenicity (provoking an immune response in the human body by substance) which makes the CRISPR/Cas9 system fails inside the body.

Researchers often employ Adeno associated virus vectors (AAV) to deliver the CRISPR/Cas system in vivo. As Cas9 protein is large in size, packaging into the AAV vector is a real challenge. Therefore researchers should either discover alternative vectors which have low immunogenicity or cas9 variants which are smaller in size for packaging.

One more issue is that the CRISPR/Cas9 system causes off-target effects by cutting the wrong piece of DNA. Therefore, it is practical to discover cas9 variants that have broad PAM compatibility and high DNA specificity to avoid this obstacle.

Further, this powerful technique should be employed responsibly and carefully not only to benefit all humankind but also to avoid misconduct which can lead to severe ethical issues in human society.


CRISPR-Cas9 gene editing: check three times, cut once

Two new studies from UC Berkeley should give scientists who use CRISPR-Cas9 for genome engineering greater confidence that they won’t inadvertently edit the wrong DNA.

The gene editing technique, created by UC Berkeley biochemist Jennifer Doudna and her colleague, Emmanuelle Charpentier, director of the Max Planck Institute of Infection Biology in Berlin, has taken the research and clinical communities by storm as an easy and cheap way to make precise changes in DNA in order to disable genes, correct genetic disorders or insert mutated genes into animals to create models of human disease.

The two new reports from Doudna’s lab and that of UC Berkeley colleague Robert Tjian show in much greater detail how the Cas9 protein searches through billions of base pairs in a cell to find the right DNA sequence, and how Cas9 determines whether to bind, or bind and cut, thereby initiating gene editing. Based on these experiments, Cas9 appears to have at least three ways of checking to make sure it finds the right target DNA before it takes the irrevocable step of making a cut.

“CRISPR-Cas9 has evolved for accurate DNA targeting, and we now understand the molecular basis for its seek-and-cleave activity, which helps limit off-target DNA editing,” said Doudna, a Howard Hughes Medical Institute investigator at UC Berkeley and professor of molecular and cell biology and of chemistry. Tjian is president of the Howard Hughes Medical Institute and a UC Berkeley professor of molecular and cell biology.

The studies also illustrate how well CRISPR/Cas9 works in human and animal cells – eukaryotes – even though “the technique was invented by bacteria to protect themselves from getting the flu,” Doudna said.

CRISPR-Cas9 is a hybrid of protein and RNA – the cousin to DNA – that functions as an efficient search-and-snip system in bacteria. It arose as a way to recognize and kill viruses, but Doudna and Charpentier realized that it could also work well in other cells, including humans, to facilitate genome editing. The Cas9 protein, obtained from the bacteria Streptococcus pyogenes, functions together with a “guide” RNA that targets a complementary 20-nucleotide stretch of DNA. Once the RNA identifies a sequence matching these nucleotides, Cas9 cuts the double-stranded DNA helix.

One study, published in the Nov. 13 issue of Science, tracked Cas9-RNA molecules though the nucleus of mammalian cells as they rapidly searched through the entire genome to find and bind just the region targeted and no other.

“It’s crazy that the Cas9 complex manages to scan the vast space of eukaryotic genomes,” said graduate student Spencer Knight, first author of the Science paper.

Previous studies had suggested that there are many similar-looking DNA regions that Cas9 could bind and cut, which could limit its usefulness if precision were important. These off-target regions might share as few as four or five nucleotides with the 20-nucleotide primer, just enough for Cas9 to recognize.

“There is a lot of off-target binding by Cas9, but we found that these interactions are very brief – from milliseconds to seconds – before Cas9 moves on,” he said.

Because these exploratory bindings – perhaps as many as 300,000 of them – are often very short-lived, a few thousand CRISPR-Cas9 complexes can scour the entire genome to find one targeted stretch of DNA. Cas9 must also recognize a short three-base-pair DNA sequence immediately following the primer sequence, dubbed PAM, which occurs about 300 million times within the human genome.

“If Cas9 bound for tens of seconds or minutes at each off-target site, it would never, ever be able to find a target and cut in a timely manner,” Knight said.

Cas9’s final checkpoint

The other study, published online Oct. 28 in Nature, showed that once Cas9 binds to a region of DNA, it performs another check before two distant sections of the Cas9 protein complex come together, like the blades of a scissors, to precisely align the active sites that cut double-stranded DNA.

“We found that RNA-guided Cas9 can bind some off-target DNA sequences, which differ from the correct target by just a few mutations, very tightly. Surprisingly, though, the region of Cas9 that does the cutting is inhibited because of the imperfect match. But when the correctly matching DNA is located, Cas9 undergoes a large structural change that releases this inhibition and triggers DNA cutting,” said first author Samuel Sternberg, who recently received his Ph.D. at UC Berkeley. He was able to observe these changes using a fluorescently labeled version of the Cas9 complex.

“We think that this structural change is the last checkpoint, or proofreading stage, of the DNA targeting reaction,” he said. “First, Cas9 recognizes a short DNA segment next to the target – the PAM – then the target DNA is matched up with the guide RNA via Watson-Crick base-pairing. Finally, when a perfect match is identified, the last part of the protein swings into place to enable cutting and initiate genome editing.”

A smaller Cas9 protein from a different species of bacteria, Staphylococcus aureus, likely exploits the same strategy to improve the precision of DNA targeting, suggesting that “this important feature has been preserved throughout evolutionary time,” he added.

“This is good news, in that it suggests that you have more than one checkpoint to ensure correct Cas9 binding,” Knight said. “There’s not just sequence regulation, there is also temporal regulation: it has to engage with the DNA and park long enough that it can actually rearrange and cut.”

The discoveries from Doudna, Tjian and their teams shed light on the molecular basis of off-target effects during genome editing applications, and may guide the future design of more accurate Cas9 variants.

The studies were funded by the National Science Foundation (MCB-1244557) and the California Institute for Regenerative Medicine (CIRM, RB4-06016).

Co-authors with Knight, Doudna and Tjian on the Science paper are Liangqi Xie, Benjamin Guglielmi, Lea Witkowsky, Lana Bosanac and Elisa Zhang of UC Berkeley Wulan Deng, Jean-Baptiste Masson and Zhe Liu of Janelia Research Campus, located in Ashburn, Virginia, of the Howard Hughes Medical Institute and Mohamed El Beheiry and Maxime Dahan of the Laboratoire Physico-Chimie Curie in the Institut Curie in Paris, France. Both Doudna and Tjian are members of the Li Ka Shing Biomedical and Health Sciences Center at UC Berkeley.

Co-authors with Sternberg and Doudna of the Nature paper are Benjamin LaFrance and Matias Kaplan of UC Berkeley.

Doudna is director of UC Berkeley’s Innovative Genomics Initiative and a faculty scientist at Lawrence Berkeley National Laboratory.

RELATED INFORMATION


Figure 1: CRISPR-Cas9 (www.stockadobe.com)

While the role of biologics in treating human diseases has evolved dramatically over the past decade, so has genetic engineering. Rational genetic engineering to enhance biotherapeutic proteins has become a reality catalyzed by publication of the genome sequences of multiple Chinese hamster ovary (CHO) cell lines. Novel “designer” CHO cells modulate posttranslational modifications (PTMs) of recombinant proteins by genome editing, and it is now possible to knock-in or knock-out genes of yeast and mammalian cells precisely (within one DNA base pair), quickly, and efficiently. Eventually, such techniques will be used in the development of cost-effective recombinant therapeutic proteins.

The gene-editing tool based on bacterial clustered regularly interspaced short palindromic repeats (CRISPR)–CRISPR-associated protein 9 (Cas9) has improved genome editing dramatically, making it faster, easier, less expensive, and more efficient than before. This system needs only one protein and one RNA molecule to achieve RNA-programed DNA cleavage. That capability has enabled CHO researchers to elucidate the mechanistic basis behind the high-level protein production and product quality attributes (PQAs) of interest. Currently, the emphasis in CHO gene editing is directed toward expanding product diversity as well as controlling and improving product quality and yields. Although routine PTM optimization across a cell’s glycosylation “machinery” is not yet a reality, researchers hope to eventually be able to combine the expression from nonmammalian hosts with human-like antibody glycosylation performed in engineered yeast or even plants.

But the use of promoter engineering to achieve precision transcriptional control for CHO-cell synthetic biotechnology could be achieved in the near future. Similarly, multiplexed genome editing (simultaneous targeting of multiple related or unrelated targets) could enable the examination and manipulation of whole genomes or protein networks.Thus, it has revolutionized knockout and knockin events and has played a crucial role in the understanding of the genomes of mammalian cells and the use of yeasts for metabolic engineering. Such editing capabilities also have enabled high, stable, and consistent expression of transgenes encoding complex therapeutic proteins such as monoclonal antibodies (MAbs).

Posttranslational Modifications
Advanced gene-editing technologies such as CRISPR-Cas9, transcription activator-like effector (TALE) nucleases, and RNA interference (RNAi) tools are complemented by the combination of next-generation sequencing with systems biotechnology to facilitate further enhancements in cell glycosylation processing. Such tools will enable cell engineers to make highly refined and targeted modifications to fast-track the processing capability of cells. That will bring consistent improvements in yield and cell quality as well as increased process efficiency, thus leading to increased affordability for future healthcare needs.

Cell engineers are witnessing a proliferation of enabling tools as molecular and cell biologists continue to develop sophisticated techniques to decipher attributes that are critical to drug-product quality. Improving the glycosylation profiles of biologics enhances product bioactivity and quality, and N-linked glycosylation plays a crucial role in the efficacy of therapeutic proteins. Proven therapeutic efficacy of many glycoengineered proteins has stimulated the development of novel optimized expression systems (e.g., mammalian cells) for producing nonfucosylated antibodies.

It is now becoming feasible to produce material rapidly for pharmacology, formulation, and toxicology studies without having to establish a stable cell line. Such capability can shorten timelines for stable cell line development from over six months to about three weeks, thus enabling a generation of stable clones at the research stage. At the same time, different production systems for glyco-optimized proteins (e.g., yeast) already have been engineered to produce the main steps of the human N-glycosylation pathway and thus have enabled biobetter versions of therapeutic MAbs.

Application of CRISPR-Cas9 tools has led to multiplexed-knockout phenotypes. Similarly, upcoming achievements with (small) noncoding RNAs in CHO cells also could support functional genomics efforts to enhance producer cells. That would expand the list of engineering targets significantly (e.g., as achieved with obinutuzumab, a humanized and glycoengineered anti-CD20 MAb, with bisected afucosylated Fc-region carbohydrates and GlycoMAb technology, from GlycArt-Roche).

Not long ago, CHO cells were considered “black boxes” because researchers lacked the genomic information of those cells. That hindered efforts to understand the molecular basis of high-quality production of recombinant proteins in CHO cells. Notably, impressive progress in CHO cell culture technology has been achieved by empirical approaches such as screening and process optimization, thus enabling high yields (10 g/L and more). However, such yields often are achieved only by certain types of cell lines, and a high degree of variability in recombinant CHO cells requires laborious and expensive processes to select the best clone for the production of each new therapeutic candidate.

Traditional recombinant cells are based on random plasmid integration of an expression cassette in the host genome. Transgenes are likely to be inserted in heterochromatin regions, which results in very weak gene-expression levels. So a substantial number of clones (generally multiple hundreds to a few thousand) must be screened to find those rare cells with a stably integrated plasmid in highly transcriptionally active chromatin regions (“hotspots”).

Sequencing efforts have resulted in availability of additional genomic sequence data for different CHO cell lines. Moreover, an active effort is underway in the CHO cell-line community to refine the genome assembly and annotation for the Chinese hamster. In the past 20 years, several molecular and cellular biology tools have been developed for targeted-site gene integration for eventually minimizing the randomness of gene insertion and increasing the predictability of high transgene expression.

Genome Editing
First-Generation Genome-Editing Tools: Several recombinases have been used for targeted site approaches, with Cre–lox and Flp–FRT being the most commonly used. Recombination-mediated cassette exchange (RMCE) technology also is attracting interest for targeted gene insertion. That technology has increased success rates and reduced timelines for the generation of stable industrial-grade CHO cell lines expressing MAbs reliably.

Second-Generation Genome-Editing Tools: This group includes endonucleases such as zinc finger nucleases (ZFN), meganucleases, transcription activator-like effectors nucleases (TALEN), and CRISPR-Cas9. Both ZFN and TALEN technologies rely on the ability to customize a DNA-binding domain for a specific sequence (the targeted sequence for cleavage) combined to a nuclease effector domain. CRISPR-Cas9 technology already has been validated for use with CHO cells and has been implemented to reduce production variability among clones. However, ZFN, TALEN and CRISPR-Cas9 technologies are used mostly for gene-specific knockout.
One critical challenge is to identify hotspots in a host genome that enable good expression levels and stability. Such specific integration might not be a one-size-fits-all approach because some therapeutic proteins could require a particular level of expression to fold correctly or to present adequate quality attributes (e.g., glycosylation and proteolytic processing). The discovery of additional naturally occurring RNA-guided nucleases offers increased targeting flexibility.

Some researchers also have engineered Cas9 enzymes to exhibit relaxed protospacer-adjacent motif (PAM) specificities. Such crucial advances expand the number of target loci that are amenable to RNA-guided genome editing. Other scientists have engineered Cas9 nuclease and managed to increase its DNA specificity dramatically. Thanks to the fusion of dead Cas9 (dCas9) to a cytidine deaminase enzyme that operates on ssDNA, “base editing” now can be used to generate point mutations in genomes.

Remarkably, CHO cells showed increased resistance to apoptosis after successful simultaneous (multiplex) disruption of fucosyltransferase 8 (FUT8), Bcl-2 associated X-protein (BAX), and Bcl-2 homologous antagonist killer (BAK) genes by CRISPR-Cas9 (1). The ZFN knockout approach achieved specific deletion of the glutamine synthetase (GS) and the dihydrofolate reductase (DHFR) genes in CHO cells, thus improving the selection stringency of the generated cell lines. Finally, both ZFN and a CRISPR-Cas9 approaches also were used for FUT8 gene-specific knockout. The resulting cell lines completely abolished fucosylation on the Fc domain of immunoglobulin G (IgG).

A less commonly used tool is mammalian artificial chromosome expression (ACE). This minigenome serves as an autonomous genetic element that replicates with cells. Its DNA sequence is customizable with different regulating elements that could help its expression.

The PiggyBac transposon system (System Biosciences) uses an efficient transposase purified from the cabbage looper moth (Trichoplusia ni) to integrate the gene of interest into a host genome. It has shown to improve yields for stable production of antibodies in CHO cell lines (2).

Genome-Editing Achievements: Introduction of additional N-glycan target sites into desired positions on the protein backbone by genetic mutation has been used to create glycoproteins with enhanced levels of glycosylation (overexpression of sialyltransferases and other glycosyltransferases, inhibition of sialidases) and sialylation. That has extended serum halflives and improved in vivo activity (3). (N-glycans also can be crucial for protein folding.) Thanks to a comprehensive ZFN knockout screen of glycosyltransferase genes and the identification of key genes that control decisive steps in N-glycosylation in CHO cells, it is now possible to provide homogeneous glycoforms.

Overexpression of B-cell lymphoma 2 (Bcl-2) or B-cell lymphoma-extra large (Bcl-xL) has shown precise and efficient inhibition of apoptosis in recombinant CHO cell cultures by enhancing the culture’s longevity, cell viability, and endurance under environmental stresses. Consequently, that renders greater yields of therapeutic protein, which is a definite economic advantage in the biopharmaceutical industry. Similarly, down-regulation of caspases (e.g., caspase-8 and -9) and knockouts of proapoptotic genes (e.g., BAX and BAK) enhance the viability of both batch and fed-batch cultures. (Down-regulation of such genes can be achieved with various genome-editing techniques such as ZFNs, TALENs, and Cas systems).

One group that helped bridge those two technologies is the research team at the Memorial Sloan Kettering Institute Center for Cell Engineering. Directed by Michel Sadelain, the group has been an integral part of CAR-T therapy research, particularly targeting the CD19 molecule on white blood cells. CD19 is expressed on cells found in B-cell leukemia and resides on the cell surface, thereby making it available for antibody access. In 2017, Sadelain’s group published a study showing that directing a CD19-specific CAR to the T-cell receptor alpha constant (TRAC) locus increased T-cell potency. The modified cells “vastly outperformed conventionally generated CAR T-cells in mouse models of acute lymphoma leukemia,” with human trials planned (1).

CAR T-cells typically are made using retroviruses, but they insert CAR genes into random loci in genomes. CRISPR can be used to deliver CAR genes to specified targets in genomes. And current research is exploring the use of CRISPR–Cas9 editing to create next-generation CAR T-cell products, including “universal” CAR T-cells, making these cells more potent and controllable (2).

References
1 Eyquem J, et al. Targeting a CAR to the TRAC Locus with CRISPR/Cas9 Enhances Tumour Rejection. Nature 543(7643) 2017: 113–117 https://doi.org/10.1038/nature21405.

Therapeutic Protein Expression Regulation By Smart Promoters and Epigenomic Reprogramming
In the near future, research laboratories will be able to design transcriptional control systems and synthetic promoters, control the timing of gene expression at will by trigger-inducible transcription factors (TF), and protect against chromatin silencing. Researchers already have begun to edit the epigenome to alter regulation of a target gene. Moreover, effector fusions can be used to expand the repertoire of genome engineering modalities achievable using Cas9. And nonediting CRISPR tools are helping researchers to screen engineered cells for phenotypes of interest quickly and precisely.

CRISPR Intellectual Property Landscape and the Gene-Editing Innovation Roadmap
There is some risk that patent filings by academic institutions claiming critical components of the CRISPR-Cas9 technology could deter or slow down the development and use of the technology. However, research organizations and universities can establish a workable balance between access and control for essential research tools through sound management of intellectual property (IP). (For example, when the Cohen–Boyer patents were granted, Stanford University created a pioneering licensing program that provided a predictable legal framework for using its inventions. Nonexclusive licenses were made available to both companies and academic institutions. In fact, patent protection has played a significant role in the development of the technology to produce recombinant DNA in bacteria).

It has been widely publicized that several organizations have been filing patents over fundamental parts of the CRISPR-Cas9 system. Commercial assignees Dow AgroSciences and DuPont Nutrition Science together hold 33 inventions, all of which are related to crop and animal agriculture (genome-editing crop and weeds) and food (dairy industry) applications of the technology. In addition, the French company Cellectis holds rights on a broad patent for gene editing of cells in vitro. Academic institutions, through their licensing and spinoffs, are primarily in control of medical applications of CRISPR-Cas (e.g., The Broad Institute of Harvard University and the Massachusetts Institute of Technology (MIT) to Editas) with commercial partners (University of California at Berkeley to Caribou Biosciences and sublicensed to Intellia and Novartis).

It’s interesting that most patent holders appear to be pursuing a strategy of keeping an international option open for their portfolios. Fortunately, signs indicate that the pace of discovery and development of CRISPR is likely to continue, with a high probability of further improvements. The Broad Institute and MIT are building a portfolio on those principles and diversifying it.

Other gene-editing technologies might emerge to compete with or possibly even displace CRISPR-Cas (see “Alternative Gene-Editing Methods” box). Follow-on breakthroughs are likely to take center stage. Current patent holders will have difficulty pursuing restricted access to CRISPR-Cas9 for research use because it already is widely used in academic laboratories. The Broad Institute, MIT, and UC Berkeley already offer free use of the technologies they control for academic research purposes through Addgene, a nonprofit organization. Addgene uses the general terms of the Uniform Biological Material Transfer Agreement (UBMTA), which indicates that discoveries are owned by the recipient of the biological material and can be licensed for commercial use.

Inevitably, some of those new commercial applications could fall within the broadly drafted claims of the original patent and therefore require a second, commercial license. Inventors of follow-on applications using a CRISPR-Cas technology are likely to need a commercial sublicense from the respective exclusive commercial licensee that controls that technology (e.g., Editas, Caribou, Intellia, CRISPR Therapeutics, and Cellectis/Calyxt) rather than from the originating university.

One group that helped bridge those two technologies is the research team at the Memorial Sloan Kettering Institute Center for Cell Engineering. Directed by Michel Sadelain, the group has been an integral part of CAR-T therapy research, particularly targeting the CD19 molecule on white blood cells. CD19 is expressed on cells found in B-cell leukemia and resides on the cell surface, thereby making it available for antibody access. In 2017, Sadelain’s group published a study showing that directing a CD19-specific CAR to the T-cell receptor alpha constant (TRAC) locus increased T-cell potency. The modified cells “vastly outperformed conventionally generated CAR T-cells in mouse models of acute lymphoma leukemia,” with human trials planned (1).

CAR T-cells typically are made using retroviruses, but they insert CAR genes into random loci in genomes. CRISPR can be used to deliver CAR genes to specified targets in genomes. And current research is exploring the use of CRISPR–Cas9 editing to create next-generation CAR T-cell products, including “universal” CAR T-cells, making these cells more potent and controllable (2).

Epigenetic Control of Therapeutic Proteins
Conferring an open chromatin state in targeted chromosome loci (DNA stretches composed of nucleosome-depleted regions) can benefit transgene expression. Indeed, cis-acting epigenetic regulatory elements can help to remodel the chromatin environment, maintaining an active transcriptional state around a transgene. One type of epigenetic regulatory element (ERE) is the scaffold/matrix attachment region (S/MAR), thanks to which recombinant proteins could improve their expression levels significantly. Another class of EREs is chromatin opening elements (UCOEs), which confer an unmethylated, open chromatin state for transgene expression. UCOEs also have helped to increase the productivity of cell line producers of recombinant proteins.

Nonetheless, S/MAR and UCOE decrease the variability of expression between the different clones. Some epigenetic elements not only can help to increase the expression level of biotherapeutics, but also can increase the number of clones that have integrated a transgene with a more defined copy number of transgenes per cell, thus accelerating the selection process.

Protein Folding and Secretion of Recombinant Cells
Secretory bottlenecks of CHO cells can be relieved by overexpressing soluble N-ethylmaleimide–sensitive factor attachment protein receptors (SNAREs). New targets for cell-engineering approaches also can be identified based on metabolomics profiling. A bottleneck at the malate dehydrogenase II (MDHII) level was characterized for the tricarboxylic acid (TCA) cycle in CHO cells, and pyruvate metabolism was shown to vary between high-producing and low-producing anti-CD20 CHO clones (2). Over the years, many cell engineering strategies have been used in attempts to increase such titers by optimizing selection markers, gene expression, cell growth, proliferation, protein folding, and secretion. Among those engineering tools, CRISPR-Cas9 and RMCE technologies will contribute significantly to the advance of glycoprotein production shortly.

With remarkable advances in genome editing, technologies such as the CRISPR-Cas9 tool as well as the combination of next-generation sequencing with systems biotechnology exhibit extraordinary potential for discovery and modulation of novel cell engineering targets. Such efforts might finally pave the way for development of rationally designed host cells. Efforts now are ongoing to engineer PTMs in microbial, insect, and plant cell systems to make those systems more suitable for therapeutic protein production.

References
1 Baek E, Noh SM, Lee GM. Antiapoptosis Engineering for Improved Protein Production from CHO Cells. Heterologous Protein Production in CHO Cells. Humana Press: New York, NY, 2017: 71–85.

2 Lalonde ME, Durocher Y. Therapeutic Glycoprotein Production in Mammalian Cells. J. Biotechnol. 251, 2017: 128–140 https://doi.org/10.1016/j.jbiotec.2017.04.028.

3 Wang Q, et al. Glycoengineering of CHO Cells to Improve Product Quality. Heterologous Protein Production in CHO Cells. Humana Press: New York, NY, 2017, 25–44.


The future of CRISPR/Cas9

The rapid progress in developing Cas9 into a set of tools for cell and molecular biology research has been remarkable, likely due to the simplicity, high efficiency and versatility of the system. Of the designer nuclease systems currently available for precision genome engineering, the CRISPR/Cas system is by far the most user friendly. It is now also clear that Cas9&rsquos potential reaches beyond DNA cleavage, and its usefulness for genome locus-specific recruitment of proteins will likely only be limited by our imagination.


Intron targeting-mediated and endogenous gene integrity-maintaining knockin in zebrafish using the CRISPR/Cas9 system

Animals carrying exogenous genes integrated at specific genomic loci are versatile tools for biological research 1 . Zebrafish (Danio rerio), an emerging vertebrate animal model, is widely used in studies on genetics, developmental biology and neurobiology. Although loss-of-function genomic editing for zebrafish has been well developed 2,3,4 , lack of feasible methods for inserting a large exogenous DNA sequence into the zebrafish genome is becoming a bottleneck for zebrafish-relevant research. It was reported that the coding sequence of enhanced green fluorescent protein (EGFP) can be integrated at the zebrafish tyrosine hydroxylase (th) locus through TALEN-mediated double-stranded breaks and homologous recombination (HR) with a low efficiency 6 . However, the targeted gene was destroyed and EGFP failed to express 6 . Recently, by using the type II bacterial clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system (CRISPR/Cas9), two non-HR-based knockin approaches were developed to insert Gal4 (a transcriptional transactivator) and EGFP into zebrafish genomic loci with a relatively high efficiency 7,8 . However, the coding sequence of targeted endogenous genes was disrupted or the expression pattern of inserted exogenous genes could not well recapitulate the endogenous ones as insertion occurred within either the exon 7 or cis-regulatory elements of targeted genes 8 . These disadvantages will limit their application in neuroscience research. Here, using the CRISPR/Cas9 system, we developed an intron targeting-mediated and HR-independent efficient knockin approach for zebrafish, with which the intactness of the coding sequence and regulatory elements of targeted endogenous genes are maintained.

The Th protein is a rate-limiting enzyme for synthesizing two important neuromodulators, dopamine and noradrenline. As dopamine and noradrenline are synthesized and released by dopaminergic and noradrenergic neurons, respectively, Th is a specific marker for these cells. We designed a short guide RNA (sgRNA) targeting the last intron of the zebrafish th and performed co-injection of sgRNA and mRNA of zebrafish codon-optimized Cas9 (zCas9) into one-cell-stage zebrafish embryo, which yielded a cleavage efficiency of ∼ 83% (Supplementary information, Table S1A and S1B). Next, we designed a donor plasmid th-P2A-EGFP consisting of three parts: a left arm, a P2A-EGFP coding sequence, and a right arm (Figure 1A). To retain the full coding sequence of th, the left arm begins from the upstream of the 5′ side of the sgRNA target site in the last intron, spans the whole last exon E13, and ends at the last base just before the stop codon of th. To keep the normal control of Th expression, the right arm includes the stop codon and 3′ regulatory elements of th. The P2A peptide is a linker for multicistronic expression 9 .

Intron targeting-mediated EGFP knockin at the zebrafish th locus. (A) Schematic of the intron targeting-mediated strategy for generating EGFP knockin at the zebrafish th locus by using the CRISPR/Cas9 system. The sgRNA target sequence is shown in red and the protospacer adjacent motif (PAM) sequence in green. The left and right arm sequences of the donor plasmid are indicated by the brown lines with double arrows. The left arm is 1 298 bp, and the right arm is 671 bp. The th-P2A-EGFP cassette was integrated into the th locus after co-injection of the donor with the sgRNA and zCas9 mRNA. The zebrafish th has 13 exons, and E12 and E13 represent the 12th and 13th exons, respectively. Right: the schematic of the mRNA and protein of the targeted th gene. (B) Representative projected in vivo confocal images (dorsal view) of th-P2A-EGFP knockin F1 larvae at 3 dpf, showing specific EGFP expression in various dopaminergic (OB, Pre, PT, and HI) and noradrenergic neurons (LC and MO). The white arrowheads mark non-specific signaling on the skin. A, anterior D, dorsal R, right. Scale bar, 50 μm. (C) PCR analysis of the 5′ and 3′ junctions of F1 progenies from the 7# founder. The F1, R1, F2 and R2 primers are shown in A. (D) 5′ and 3′ junction sequences of F1 progenies of three th-P2A-EGFP knockin F0 founders. The indel mutations are highlighted in yellow, and the PAM and sgRNA target sequences are shown in green and red, respectively. (E) Whole-mount in situ double immunostaining of th-P2A-EGFP knockin F1 larvae, showing that EGFP signaling (green) co-localizes with Th signals (red). The white arrowheads mark non-specific signaling on the skin. Scale bar, 20 μm. (F) Western blot of the Th expression in WT and heterozygous th-P2A-EGFP knockin F1 embryos. (G) Dopamine (DA) immunostaining of th-P2A-EGFP knockin F1 (top) and WT larvae (bottom left). Bottom right: DA intensity of DA-positive neurons in WT and knockin F1 larvae. The numbers on the bars represent the numbers of cells examined. Scale bar, 10 μm. (H) Bright-field image showing in vivo whole-cell recording of EGFP-expressing LC neurons in a homozygous th-P2A-EGFP knockin F2 larvae. The black arrow indicates the recording microelectrode. Scale bar, 10 μm. (I) Whole-cell currents of an EGFP-expressing LC neuron. Under voltage-clamp mode, the neuron was held at −60 mV and voltage pulses from −100 to 30 mV with an interval of 10 mV were applied. Action potential currents (arrow) appear near −30 mV.

We co-injected the donor plasmid, sgRNA and the zCas9 mRNA into one-cell-stage fertilized zebrafish egg. As both the donor plasmid DNA and the last intron of th contain sgRNA target site, concurrent cleavage by sgRNA/Cas9 would result in efficient and specific integration of the donor DNA into the th locus via non-HR. Indeed, we observed EGFP expression in the brain of injected larvae 3 days post fertilization (dpf) (33/139 Supplementary information, Figure S1A1 and Table S1B). Based on their location and morphology 10 , EGFP-expressing cells included dopaminergic neurons in the posterior tubercular (PT), intermediate hypothalamus (HI) and pretectum (Pre), and noradrenergic neurons in the locus coeruleus (LC) and medulla oblongata (MO) (Supplementary information, Figure S1A1). Successful non-HR-mediated insertion of the th-P2A-EGFP donor was then verified by PCR using target site- and donor-specific primers and junction sequencing analysis (Supplementary information, Figures S1A2 and S1A3). The specificity of knockin events was further confirmed by in situ immunohistochemistry staining, which revealed that EGFP was co-localized with Th in 46 out of 48 Th-positive cells in three larvae examined (Supplementary information, Figures S1A4 and S1A5).

To examine the germline transmission of knockin events, 25 embryos showing mosaic expression of EGFP were raised to adulthood. Each of them was then outcrossed to wild-type (WT) zebrafish, and their F1 progenies were screened for EGFP signal. Three F0 founders were identified, and EGFP-positive F1 progenies were produced at rates ranging from 15.5% to 21.1% (Supplementary information, Table S1C). As expected, in comparison with F0 (Supplementary information, Figure S1A1), more EGFP-expressing dopaminergic and noradrenergic neurons were observed in F1 progenies (Figure 1B), including neurons in the olfactory bulb (OB), Pre, PT, HI, LC and MO. PCR and junction sequencing analysis of F1 progenies confirmed the inheritance of the genomic integration of their corresponding F0 founders (Figure 1C and 1D). Immunostaining was also performed in the F1 embryos of th-P2A-EGFP knockin fish, and EGFP signal was found to be well co-localized with the Th protein (98% ± 1%, mean ± SEM, in 5 larvae Figure 1E), suggesting the high specificity of EGFP expression in the stable knockin lines.

As the full reading frame and regulatory elements of th were maintained by using this knockin strategy, both the integrity and expression pattern of the gene product should be normal. To examine these points, we extracted the total protein from F1 embryos carrying EGFP knockin at th and performed western blot analysis. F1 embryos were heterozygous because they were generated by crossing knockin F0 founders with WT fish. By using a Th antibody, two bands for knockin embryos were detected (Figure 1F). The lower band at around 56 kDa represents the WT Th protein derived from a WT th allele. The P2A peptide is about 2 kDa and cleaved between the last two amino acids. If knockin events did not affect the integrity of the Th protein, the cleavage of the Th-P2A-EGFP protein will result in two products: Th-P2A fusion protein (58 kDa) and EGFP protein (Figure 1A). Therefore, the upper band at around 58 kDa indicates the integrity of the Th protein produced from a knockin th allele. Furthermore, the expression levels of the WT Th protein and the knockin Th-P2A fusion protein were almost equal (Figure 1F), further suggesting that our knockin strategy does not impair the expression level of the targeted endogenous gene. To examine whether knockin events affect Th functions, we then performed immunostaining of dopamine, the level of which can reflect the activity of Th. The intensities of dopamine signals in dopaminergic neurons were not significantly different between knockin F1 and WT embryos (P = 0.4 Figure 1G), suggesting that Th function is not affected by knockin events.

To examine the physiological normality of neurons carrying targeted integration, in vivo whole-cell recording was subsequently performed in homozygous th-P2A-EGFP knockin F2 larvae (Figure 1H). EGFP-expressing neurons exhibited a normal intrinsic membrane property, as reflected by outwardly rectifying whole-cell currents (Figure 1I).

To extend the application of our knockin strategy to other exogenous genes, we generated knockin fish carrying the transactivator protein Gal4 at the th locus by using the same strategy, in which only the EGFP coding sequence was replaced with the Gal4 sequence (th-P2A-Gal4 Supplementary information, Figure S1B1). After injection of Gal4 knockin-relevant elements into fertilized eggs of Tg(UAS:GCaMP5) transgenic zebrafish, integration events were visualized by the expression of GCaMP5 in dopaminergic or noradrenergic neurons (Supplementary information, Figure S1B2). As GCaMP5 is a genetically encoded calcium indicator, we could observe mechanical stimulus-induced calcium responses in neurons by puffing water to the fish tail through a micropipette. We also injected the Gal4 knockin-relevant elements into WT fish to screen for th-P2A-Gal4 knockin founders. As the Gal4 protein has no fluorescence, we raised the injected knockin embryos to adulthood without prior selection and crossed these adults with Tg(UAS:Kaede) transgenic fish. Two founders were identified among the total of 28 injected fish (Supplementary information, Table S1D), as evidenced by the fact that dopaminergic neurons were labeled by Kaede in their progenies, which were produced at a mean rate of ∼ 7% (Supplementary information, Figure S1B4 and Table S1D). Successful insertion of the th-P2A-Gal4 donor was then verified by PCR and junction sequencing analysis in F1 progenies (Supplementary information, Figures S1B5 and S1B6).

It was reported that the CRISPR/Cas9 system shows a high frequency of off-target (OT) cleavage in human cell lines, and the specificity of Cas9 targeting can tolerate up to three base pair (bp) mismatches between a sgRNA and its target DNA 11 . We therefore searched all zebrafish genomic loci containing up to 3-bp mismatches in comparison with the coding sequence of the th sgRNA, and found three potential OT sites. PCR and sequencing analysis of those potential OT sites in the genome of injected WT embryos or th-P2A-EGFP knockin F1 embryos did not reveal indels (Supplementary information, Table S1E), suggesting a low OT rate associated with our knockin strategy.

The applicability of our knockin strategy was further validated by targeting other endogenous genes specifically expressed in different types of cells, as exemplified by the integration of EGFP into the zebrafish tryptophan hydroxylase 2 (tph2), glial fibrillary acidic protein (gfap), and flk1 loci. These EGFP insertions resulted in the specific labeling of serotoninergic neurons, glia and vascular endothelial cells, respectively (Supplementary information, Figures S1C-S1E, and Table S1A and S1B). It is worth noticing that, in the case of the tph2 knockin, the second last intron was selected for targeting, indicating that the last intron is the first but not the only choice for targeting. Furthermore, by replacing the P2A in the gfap-P2A-EGFP plasmid with a flexible serine-serine linker sequence, we succeeded in fusing an EGFP tag to endogenous Gfap (Supplementary information, Figure S1F), demonstrating that our knockin strategy can also be used to tag endogenous proteins.

Taking advantage of both the HR for donor design and the non-HR for donor integration, we developed a novel CRISPR/Cas9-mediated intron-targeting knockin strategy, by which knockin zebrafish can be efficiently generated without disruption of targeted endogenous genes. Compared with HR, error-prone non-homologous end joining (NHEJ)-involved non-HR knockin for zebrafish has two advantages. First, NHEJ is at least 10-fold more active than HR during early zebrafish development 12 . Second, unlike HR, NHEJ does not need the precise homology between the parent zebrafish and the targeting donor, avoiding time-consuming screening and genotyping of parent animals. More importantly, to maintain the integrity of targeted endogenous genes, we designed sgRNAs targeting introns, so that NHEJ-mediated indel mutations do not change the reading frame of targeted genes. In addition, intron targeting also theoretically increases the rate of in-frame insertion up to 3-fold in comparison with exon-based targeting. Furthermore, we artificially added the endogenous genome sequence spanning from the sgRNA target site to the 3′ intergenic region into donor plasmids. Therefore, the predicted forward ligation of the donor into the targeted locus retains the original reading frame and both 5′ and 3′ regulatory elements of targeted genes. Taken together, this strategy has two advantages: (1) inserted exogenous genes can faithfully recapitulate the expression pattern of targeted endogenous genes (2) the expression and function of targeted endogenous genes are maintained. Thus, the readiness, high efficiency and targeted gene integrity maintenance make our strategy an applicable knockin approach for zebrafish and even other organisms.


1. Introduction

For many years, molecular biologists have sought ways to use cellular repair mechanisms to manipulate DNA through genome editing. In this way, they would have the power to change the genome by correcting a mutation or introducing a new function (Rodriguez, 2016). For this purpose, genome editing technologies were developed (Memi et al., 2018). In recent years, clustered regularly interspaced short palindromic repeats technology (CRISPR-Cas9) has become the most preferred method of gene editing. This technology has advantages such as high accuracy, easy handling, and relatively low cost compared to previous technologies, such as zinc-finger nuclease (ZFN) and transcription activator-like effector nuclease (TALEN). Thanks to these benefits, CRISPR-Cas9 technology can be easily applied in any molecular biology laboratory.

Genome editing technologies are used in the formation of human disease models in experimental animals and for the understanding of basic gene functions. They also have great therapeutic potential for future treatment of untreated diseases such as certain cancers, genetic disorders, and HIV/AIDS. Today, genome editing in somatic cells is one of the promising areas of therapeutic development (Otieno, 2015). However, various bioethical issues have arisen due to the potential impact of these technologies on the safety of food stocks and clinical applications (Hundleby and Harwood, 2018 Hirch et al., 2019). This review discusses the challenges, possible consequences, and bioethical issues of CRISPR-Cas9 in detail.


Chickens and pigs with integrated genetic scissors

Researchers at the TUM have demonstrated a way to efficiently study molecular mechanisms of disease resistance or biomedical issues in farm animals. Researchers are now able to introduce specific gene mutations into a desired organ or even correct existing genes without creating new animal models for each target gene. This reduces the number of animals required for research..

CRISPR/Cas9 enables desired gene manipulations

CRISPR/Cas9 is a tool to rewrite DNA information. Genes can be inactivated or specifically modified using this method. The CRISPR/Cas9 system consists of two components.

The gRNA (guide RNA) is a short sequence that binds specifically to the DNA segment of the gene that is to be modified. The Cas9 nuclease, the actual "gene scissors," binds to the gRNA and cuts the respective section of the target DNA. This cut activates repair mechanisms that can inactivate gene functions or incorporate specific mutations.

Healthy chickens and pigs with integrated gene scissors

"The generated animals provide the gene scissors, the Cas9 protein, right along with them. So all we have to do is to introduce the guide RNAs to get animals which have specific genetic characteristics," explains Benjamin Schusser, Professor of Reproductive Biotechnology at the TUM. "The initial generation of these animals took about three years. Cas9 can now be used at all stages of animal development, since every cell in the body permanently possesses the Cas9 protein. We have been successfully able to utilize this technique in chicken embryos as well as in living pigs."

The healthy chickens and pigs produced by the researchers thus possess the Cas9 nuclease in all organs studied. This is particularly useful in biomedical and agricultural research.

Analytical tool to fight viral or cancer diseases

Pigs are used as disease models for humans because their anatomy and physiology are much more similar to humans in comparison to mice (currently a common disease model). Thus, a modified pig may help to better understand the mechanism of carcinogenesis in humans. Potential new treatments for humans can also be tested in animal models.

"Due to the presence of Cas9 in the cells the processes are significantly accelerated and simplified," says Angelika Schnieke, Professor of Livestock Biotechnology at the TUM. "Cas9-equipped animals make it possible, for example, to specifically inactivate tumor-relevant genes and to simulate cancer development."

Cas9 pigs and chickens enable researchers to test which genes might be involved in the formation of traits, such as disease resistance, directly in the animal. "The mechanism of the CRISPR/Cas9 system may also be useful for combating infections using DNA viruses. Initial cell culture experiments showed that this already works for the avian herpes virus," says Prof. Schusser.

Important resource for biomedical and agricultural research

Prof. Schnieke notes, "Our Cas9-expressing chickens and pigs represent an innovative resource for genome editing in the biomedical and agricultural sciences, but beyond that, these animals are also available to other research groups. Hence, efficient genome editing in living animals has the potential to significantly advance biomedical and agricultural research."



Comments:

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