Where is the most promising avenue to success in Pharmaceuticals with CRISPR-Cas9?

Author: Larry H. Bernstein, MD, FCAP

Where is the most promising avenue to success in Pharmaceuticals with CRISPR-Cas9?  Volume 2 (Volume Two: Latest in Genomics Methodologies for Therapeutics: Gene Editing, NGS and BioInformatics, Simulations and the Genome Ontology), Part 2: CRISPR for Gene Editing and DNA Repair

There has been a rapid development of methods for genetic engineering that is based on an initial work on bacterial resistance to viral invasion.  The engineering called RNA inhibition (RNA_i) has gone through several stages leading to a more rapid and more specific application with minimal error.

It is a different issue to consider this application with respect to bacterial, viral, fungal, or parasitic invasion than it would be for complex human metabolic conditions and human cancer. The difference is that humans and multi-organ species are well differentiated systems with organ specific genome translation to function.

I would expect to see the use of genomic alteration as most promising in the near term for the enormous battle against antimicrobial, antifungal, and antiparasitic drug resistance.  This could well be expected to be a long-term battle because of the invading organisms innate propensity to develop resistance.

A CRISPR/Cas system mediates bacterial innate immune evasion and virulence

Timothy R. Sampson, Sunil D. Saroj, Anna C. Llewellyn, Yih-Ling Tzeng & David S. Weiss

Affiliations, Contributions, Corresponding author

Nature 497, 254–257 (09 May 2013),  http://dx.doi.org:/10.1038/nature12048

CRISPR/Cas (clustered regularly interspaced palindromic repeats/CRISPR-associated) systems are a bacterial defence against invading foreign nucleic acids derived from bacteriophages or exogenous plasmids^{1, 2, 3, 4}. These systems use an array of small CRISPR RNAs (crRNAs) consisting of repetitive sequences flanking unique spacers to recognize their targets, and conserved Cas proteins to mediate target degradation^{5, 6, 7, 8}. Recent studies have suggested that these systems may have broader functions in bacterial physiology, and it is unknown if they regulate expression of endogenous genes^9, 10. Here we demonstrate that the Cas protein Cas9 of Francisella novicida uses a unique, small, CRISPR/Cas-associated RNA (scaRNA) to repress an endogenous transcript encoding a bacterial lipoprotein. As bacterial lipoproteins trigger a proinflammatory innate immune response aimed at combating pathogens^11, 12, CRISPR/Cas-mediated repression of bacterial lipoprotein expression is critical for F. novicida to dampen this host response and promote virulence. Because Cas9 proteins are highly enriched in pathogenic and commensal bacteria, our work indicates that CRISPR/Cas-mediated gene regulation may broadly contribute to the regulation of endogenous bacterial genes, particularly during the interaction of such bacteria with eukaryotic hosts.

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Zhang lab unlocks crystal structure of new CRISPR/Cas9 genome editing tool

Paul Goldsmith,  2015 Aug

In a paper published today in Cell researchers from the Broad Institute and University of Tokyo revealed the crystal structure of theStaphylococcus aureus Cas9 complex (SaCas9)—a highly efficient enzyme that overcomes one of the primary challenges to in vivo mammalian genome editing.

First identified as a potential genome-editing tool by Broad Institute core member Feng Zhang and his colleagues (and published by Zhang lab in April 2015), SaCas9 is expected to expand scientists’ ability to edit genomes in vivo. This new structural study will help researchers refine and further engineer this promising tool to accelerate genomic research and bring the technology closer to use in the treatment of human genetic disease.

“SaCas9 is the latest addition to our Cas9 toolbox, and the crystal shows us its blueprint,” said co-senior author Feng Zhang, who in addition to his Broad role, is also an investigator at the McGovern Institute for Brain Research, and an assistant professor at MIT.

The engineered CRISPR-Cas9 system adapts a naturally-occurring system that bacteria use as a defense mechanism against viral infection. The Zhang lab first harnessed this system as an effective genome-editing tool in mammalian cells using the Cas9 enzymes from Streptococcus thermophilus (StCas9) andStreptococcus pyogenes (SpCas9). Now, Zhang and colleagues have detailed the molecular structure of SaCas9, providing scientists with a high-resolution map of this enzyme. By comparing the crystal structure of SaCas9 to the crystal structure of the more commonly-used SpCas9 (published by the Zhang lab in February 2014), the team was able to focus on aspects important to Cas9 function— potentially paving the way to further develop the experimental and therapeutic potential of the CRISPR-Cas9 system.

Paper cited: Nishimasu H et al. “Crystal Structure of Staphylococcus aureus Cas9.” Cell, http://dx.doi.org:/10.1016/j.cell.2015.08.007

Advances in CRISPR-Cas9 genome engineering: lessons learned from RNA interference

Rodolphe Barrangou1,†, Amanda Birmingham2,†, Stefan Wiemann3, Roderick L. Beijersbergen4, Veit Hornung5 and Anja van Brabant Smith2
Nucleic Acids Research, 2015 Mar 23.  http:dx.doi.org:/10.1093/nar/gkv226

RNAi and CRISPR-Cas9 have many clear similarities. Indeed, the mechanisms of both use small RNAs with an on-target specificity of ∼18–20 nt. Both methods have been extensively reviewed recently (3–5) so we only highlight their main features here. RNAi operates by piggybacking on the endogenous eukaryotic pathway for microRNA-based gene regulation (Figure 1A). microRNAs (miRNAs) are small, ∼22-nt-long molecules that cause cleavage, degradation and/or translational repression of RNAs with adequate complementarity to them(6).RNAi reagentsfor research aim to exploit the cleavage pathway using perfect complementarity to their targets to produce robust downregulation of only the intended target gene. The CRISPRCas9 system, on the other hand, originates from the bacterial CRISPR-Cas system, which provides adaptive immunity against invading genetic elements (7). Generally, CRISPR-Cas systems provide DNA-encoded (7), RNAmediated (8), DNA- (9) or RNA-targeting(10) sequencespecific targeting. Cas9 is the signature protein for Type II CRISPR-Cas systems (11

Figure 1. (not shown) The RNAi and CRISPR-Cas9 pathways in mammalian cells. (A) miRNA genes code for primary miRNAs that are processed by the Drosha/DGCR8 complex to generate pre-miRNAs with a hairpin structure. These molecules are exported from the nucleus to the cytoplasm, where they are further processed by Dicer to generate ∼22-nt-long double-stranded mature miRNAs. The RNA duplex associates with an Argonaute (Ago) protein and is then unwound; the strand with a more unstable 5 end (known as the guide strand) is loaded into Ago to create the RNA-induced silencing complex (RISC) while the unloaded strand is discarded. Depending on the degree of complementarity to their targets, miRNAs cause either transcript cleavage and/or translational repression and mRNA degradation. siRNAs directly mimic mature miRNA duplexes, while shRNAs enter the miRNA pathway at the pre-miRNA hairpin stage and are processed into such duplexes. (B) CRISPR-Cas9-mediated genome engineering in mammalian cells requires crRNA, tracrRNA and Cas9. crRNA and tracrRNA can be provided exogenously through a plasmid for expression of a sgRNA, or chemically synthesized crRNA and tracrRNA molecules can be transfected along with a Cas9 expression plasmid. The crRNA and tracrRNA are loaded into Cas9 to form an RNP complex which targets complementary DNA adjacent to the PAM. Using the RuvC and HNH nickases, Cas9 generates a double-stranded break (DSB) that can be either repaired precisely (resulting in no genetic change) or imperfectly repaired to create a mutation (indel) in the targeted gene. There are a myriad of mutations that can be generated; some mutations will have no effect on protein function while others will result in truncations or loss of protein function. Shown are mutations that will induce a frame shift in the coding region of the mRNA (indicated by red X’s), resulting in either a truncated, non-functional protein or loss of protein expression due to nonsense-mediated decay of the mRNA.

Both RNAi and CRISPR-Cas9 have experienced significant milestones in their technological development, as highlighted in Figure 2 (7–14,16–22,24–51) (highlighted topics have been detailed in recent reviews (2,4,52–58)). The CRISPR-Cas9 milestones to date have mimicked a compressed version of those for RNAi, underlining the practical benefit of leveraging similarities to this well-trodden research path. While RNAi has already influenced many advances in the CRISPR-Cas9 field, other applications of CRISPR-Cas9 have not yet been attained but will likely continue to be inspired by the corresponding advances in the RNAi field (Table 1). Of particular interest are the potential parallels in efficiency, specificity, screening and in vivo/therapeutic applications, which we discuss further below.

Figure2. Timeline of milestones for RNAi and CRISPR-Cas9. Milestones in the RNAi field are noted above the line and milestones in the CRISPR-Cas9 field are noted below the line. These milestones have been covered in depth in recent reviews (2,4,52–29).
Table 1. Summary of improvements in the CRISPR-Cas9 field that can be anticipated by corresponding RNAi advances

Work performed during the first few years of intensive RNAi investigations demonstrated that, when taking 70– 75% reduction in RNA levels as a heuristic threshold for efficiency (59), only a small majority of siRNAs and shRNAs function efficiently (24,60) when guide strand sequences are chosen randomly. This observation led to the development in 2004 of rational design algorithms for siRNA molecules (Figure2), followed later by similar algorithms for shRNAs. These methods have been able to achieve∼75% correlation and >80% positive predictive power in identifying functional siRNAs (61) but have been somewhat less effective for shRNAs (62) (perhaps because in most cases, shRNAs produce less knockdown than do siRNAs, likely due to a smaller number of active molecules in each cell). crRNAs also vary widely in efficiency: reports have demonstrated indel (insertion and deletion) creation rates between 5 and 65% (20,25), though the average appears to be between 10 and 40% in unenriched cell populations. Indeed, a growing amount of evidence suggests a wide range of crRNA efficiency between genes and even between exons of the same gene, yielding some ‘super’ crRNAs that are more functional(26,27).

Perhaps in no other area are the lessons of RNAi as obvious as in that of specificity. While RNAi was originally hailed as exquisitely specific (64), subsequent research has shown that in some circumstances it can trigger non-specific effects and/or sequence-specific off-target effects (65). Many non-specific effects seen with this approach are mediated by the inadvertent activation of pattern recognition receptors (PRRs) of the innate immune system that have evolved to sense the presence of nucleic acids in certain sub-cellular compartments. siRNA length, certain sequence motifs, the absence of 2-nt 3 overhangs and cell type are important factors for induction of the mammalian interferon response (66–68). Additionally, the general perturbation of cellular or tissue homeostasis by the delivery process itself can also trigger unwanted responses (most likely secondary to innate immune damage-sensing pathways) such as the wide-spread alteration of gene expression caused by cationic lipids, especially when used at high concentrations (69). Such nonspecific effects associated with delivery will still exist for CRISPR-Cas9 but can likely be overcome by minimizing lipid concentration as is now routinely done in RNAi studies. Similarly, the introduction of chemical modifications into the backbone of an siRNA duplex (e.g. 2-O-methyl ribosyl) can block the recognition of RNA molecules by PRRs (66,70–71),

RNAi can also produce sequence-specific off-target effects, which were initially described in early 2003 (31), but whose potential impact was not fully appreciated until well after the method had become a widely used research and screening technique (e.g. (74)). Cleavage-based off-targeting, which occurs when RISC encounters an unintended transcript target with perfect or near-perfect complementarity to its guide strand, can induce knockdownequivalenttothatofintendedtargetdown-regulation and was originally hypothesized to be the main cause of sequence-specific off-target effects. It took several years to determine that these effects were in fact primarily caused byRNAireagentsactingina‘miRNA-like’fashion,downregulating unintended targets by small (usually <2-fold) amounts primarily through seed-based interactions with the 3 UTR of those unintended targets. Because miRNAlike off-targeting is generally seed-based and all transcripts contain matches to a variety of 6–8-base motifs, such off targeting can affect tens to hundreds of transcripts. Furthermore, if the RNAi reagent contains a seed mimicking that of an endogenous miRNA, the off-targeting may affect the pathway or family of targets evolutionarily selected for regulation by that miRNA. It is not possible to design RNAi reagents that do not contain seed regions found in the transcriptome’s 3 UTRs and the non-seed factors that conclusively determine whether or not a seed-matched transcript is in fact off-targeted have not yet been identified. Both rational design and chemical modifications such as 2 O-methyl ribosyl substitutions can mitigate seed-based off-target effects (32), but without a full solution, specificity remains a well-known pain point for RNAi users.

Of particular importance is evaluating whether the lower efficiencies seen using CRISPR-Cas9 are sufficient to generate a desired phenotype in the screening assay––that is, determining whether the phenotype is detectable in the targeted cell population. In this regard, two factors are of special concern: the ploidy of the gene locus of interest (as tumor cell lines are often aneuploid) and the likelihood of disrupting the reading frame by the induced mutation (since +3 or−3 indels would not serve this purpose). Taking these factors into account, the chance of obtaining a high percentage of cells that have a functional knockout in a bulk cell culture is relatively low under typical screening conditions. Consequently, it is unlikely that traditional arrayed loss-of-signal screens such as those common in RNAi will be widely feasible in bulk-transfected cells using CRISPR-Cas9.

RNAi has demonstrated tremendous value as a functional genomics tool, especially with the technological advances described above that enhance efficiency and decrease offtarget effects (118). Likewise, CRISPR-Cas9 has already proven to be a valuable tool for functional genomics studies. Although we have highlighted many points on which the RNAi field can offer pertinent guidance for the effective development and exploitation of CRISPR-Cas9, it is important to remember the fundamental differences that underlie these techniques (Table 3). These contrasts must be considered when selecting the most appropriate method for studying a particular gene or genome.
Molecular consequences. One such fundamental difference between the two is the molecular consequences of their actions. RNAi results in knockdown at the RNA level while CRISPR-Cas9 causes a change in the DNA of the genome; as a corollary, RNAi happens predominantly in the cytoplasm, while CRISPRCas9 acts in the nucleus. These contrasts highlight the differing applicability of the techniques: for example, circRNAs (119,120) that differ from their linear counterparts by splice order in the final transcript can be interrogated by RNAi but not CRISPR-Cas9, while intron functionality can be investigated by CRISPR-Cas9 but not RNAi. For more prosaic targets of interest, in some cases the resulting phenotype associated with either knockdown or knockout may be similar but in others there may be significant differences that result from repression of gene expression compared to a complete null genotype.AlthoughCRISPRCas9-based approaches for drug target identification have been developed (121), repression of gene expression may better model a potential drug’s means of activity and thus be more relevant for drug discovery efforts.

Duration of effect. Because of differences in their mode of action, CRISPRCas9 and RNAi also differ in their duration of effect. siRNA knockdown is typically transient (lasting 2–7 days), while genome engineering with CRISPR-Cas9 induces a permanent effect that, if all alleles are affected, sustainably removes gene function and activity. shRNA knockdown can be either short- or long-term depending on whether the shRNA is continuously expressed, providing some middleground; shRNA activity can also be turned on and off with inducible vectors (122,123) although some leakage can occur even in the off state, depending on the inducible system. Inducible or transient systems will also likely be necessary for studying essential genes viaCRISPR-Cas9

Modulation of non-coding genes Most protein-coding genes will be easily down-modulated by either RNAi or CRISPR-Cas9. For permanent disruption of protein-coding genes using CRISPR-Cas9, frameshift mutations in a critical coding exon (i.e. an early protein-coding exon that is used by all relevant transcript variants) must occur, while RNAi reagents can be targeted essentially anywhere within the transcript.However,knockdown or knockout of non-coding RNAs is more nuanced. The study of small non-coding genes, particularly, is complicated for both RNAi and CRISPR-Cas9 by the limited design space for targeting the non-coding gene without affecting nearby genes.

The fact that CRISPR-Cas9 is not an endogenous mammalian system provides the opportunity for innovative protein evolution studies that are not possible with RNAi. Given this, we anticipate that the CRISPR-Cas9 field will expand beyond the canonical S. pyogenes SpyCas9 in combination with the NGG PAM that has been the focus of virtually all mammalian applications to date. Indeed, other Cas9 proteins are being increasingly characterized (145) with their respective PAMs (of various sizes and sequences) in order to expand targeting specificity.

The new frontier of genome engineering with CRISPR-Cas9
GENOME EDITING
Jennifer A. Doudna* and Emmanuelle Charpentier
Science 346, 1258096 (2014). http://dx.doi .org/10.1126/ science.125809

Fig. 1.Timeline of CRISPR-Cas and genome engineering research fields. Key developments in both fields are shown. These two fields merged in 2012 with the discovery that Cas9 is an RNA-programmable DNA endonuclease, leading to the explosion of papers beginning in 2013 in which Cas9 has been used to modify genes in human cells as well as many other cell types and organisms.

Functionality of CRISPR-Cas9 Bioinformatic analyses first identified Cas9 (formerly COG3513, Csx12, Cas5, or Csn1) as a large multifunctional protein (36) with two putative nuclease domains, HNH (38, 43, 44) and RuvC-like (44). Genetic studies showed that S. thermophilus Cas9 is essential for defense against viral invasion (45, 66), might be responsible for introducing DSBs into invading plasmids and phages (67), enables in vivo targeting of temperate phages and plasmids in bacteria (66, 68), and requires the HNH and RuvC domains to interfere with plasmid transformation efficiency (68). In 2011 (66), trans-activating crRNA (tracrRNA) —a small RNA that is trans-encoded upstream of the type II CRISPR-Cas locus in Streptococcus pyogenes—was reported to be essential for crRNA maturation by ribonuclease III and Cas9, and tracrRNA-mediated activation of crRNA maturation was found to confer sequence-specific immunity against parasite genomes. In 2012 (64), the S.pyogenes CRISPR-Cas9proteinwasshown tobeadual-RNA–guidedDNAendonucleasethat uses the tracrRNA:crRNA duplex (66) to direct DNA cleavage (64) (Fig. 2). Cas9 uses its HNH domain to cleave the DNA strand that is complementary to the 20-nucleotide sequence of the crRNA; the RuvC-like domain of Cas9 cleaves the DNA strand opposite the complementary strand (64, 65) (Fig. 2). Mutating either the HNH or the RuvC-like domain in Cas9 generates a variant protein with single-stranded DNA cleavage (nickase) activity, whereas mutating both domains (dCas9; Asp10 → Ala, His840 → Ala) results in an RNA guided DNA binding protein(64,65). DNA target recognition requires both base pairing to the crRNA sequence and the presence of a short sequence (PAM) adjacent to the targeted sequence in the DNA (64, 65) (Fig. 2). The dual tracrRNA:crRNA was then engineered as a single guide RNA (sgRNA) that retains two critical features: the 20-nucleotide sequence at the 5′end of the sgRNA that determines the DNA target site by Watson-Crick base pairing,and the double-stranded structure at the 3′ side of the guide sequence that binds to Cas9 (64) (Fig. 2). This created a simple two-component system in which changes to the guide sequence (20 nucleotides in the native RNA) of the sgRNA can be used to program CRISPR-Cas9 to target any DNA sequence of interest as long as it is adjacent to a PAM (64).

Fig. 2. Biology of the type II-A CRISPR-Cas system.The type II-A system from S. pyogenes is shown as an example. (A) The cas gene operon with tracrRNA and the CRISPR array. (B) The natural pathway of antiviral defense involves association of Cas9 with the antirepeat-repeat RNA (tracrRNA: crRNA) duplexes, RNA co-processing by ribonuclease III, further trimming, R-loop formation, and target DNA cleavage. (C) Details of the natural DNA cleavage with the duplex tracrRNA:crRNA

Mechanism of CRISPR-Cas9–mediated genome targeting. Structural analysis of S. pyogenes Cas9 has revealed additional insights into the mechanism of CRISPR-Cas9 (Fig. 3). Molecular structures of Cas9 determined by electron microscopy and x-ray crystallography show that the protein undergoes large conformational rearrangement upon binding to the guide RNA, with a further change upon association with a target doublestranded DNA (dsDNA). This change creates a channel, running between the two structural lobes of the protein, that binds to the RNA-DNA hybrid as well as to the coaxially stacked dualRNA structure of the guide corresponding to the crRNA repeat–tracrRNA antirepeat interaction (77, 78). An arginine-rich a helix (77–79) bridges the two structural lobes of Cas9 and appears to be the hinge between them.

Fig. 4. CRISPR-Cas9 as a genome engineering tool. (A) Different strategies for introducing blunt double-stranded DNA breaks into genomic loci, which become substrates for endogenous cellular DNA repair machinery that catalyze nonhomologous end joining (NHEJ) or homology-directed repair (HDR). (B) Cas9 can function as a nickase (nCas9) when engineered to contain an inactivating mutation in either the HNH domain or RuvC domain active sites. When nCas9 is used with two sgRNAs that recognize offset target sites in DNA, a staggered double-strand break is created. (C) Cas9 functions as an RNA-guided DNA binding protein when engineered to contain inactivating mutations in both of its active sites.This catalytically inactive or dead Cas9 (dCas9) can mediate transcriptional down-regulation or activation, particularly when fused to activator or repressor domains. In addition, dCas9 can be fused to fluorescent domains, such as green fluorescent protein (GFP), for live-cell imaging of chromosomal loci. Other dCas9 fusions, such as those including chromatin or DNA modification domains, may enable targeted epigenetic changes to genomic DNA.

The programmable binding capability of dCas9 can also be used for imaging of specific loci in live cells. An enhanced green fluorescent protein– tagged dCas9 protein and a structurally optimized sgRNA were shown to produce robust imaging of repetitiveand nonrepetitiveelementsin telomeres and coding genes in living cells (131). This CRISPR imaging tool has the potential to improve the current technologies for studying conformational dynamics of native chromosomes in living cells, particularlyifmulticolorimagingcanbedeveloped using multiple distinct Cas9 proteins. It may also be possible to couple fluorescent proteins or small molecules to the guide RNA, providing an orthogonal strategy for multicolor imaging using Cas9. Novel technologies aiming to disrupt proviruses may be an attractive approach to eliminating viral genomes from infected individuals and thus curing viral infections. An appeal of this strategy is that it takes advantage of the primary native functions of CRISPR-Cas systems as antiviral adaptive immune systems in bacteria. The targeted CRISPR-Cas9 technique was shown to efficiently cleave and mutate the long terminal repeat sites of HIV-1 and also to remove internal viral genes from the chromosome of infected cells (132, 133). CRISPR-Cas9 is also a promising technology in the field of engineering and synthetic biology. A multiplex CRISPR approach referred to as CRISPRm was developed to facilitate directed evolution of biomolecules (134). CRISPRm consists of the optimization of CRISPR-Cas9 to generate quantitative gene assembly and DNA library insertion into the fungal genomes, providing a strategy to improve the activity of biomolecules. In addition, it has been possible to induce Cas9 to bind single stranded RNA in a programmable fashion by using short DNA oligonucleotides containing PAM sequences (PAMmers) to activate the enzyme, suggesting new ways to target transcripts without prior affinity tagging (135).  Several groups have developed algorithmic tools that predict the sequence of an optimal sgRNA with minimized off-target effects (for example, http://tools.genome-engineering.org, http://zifit.partners.org, and www.e-crisp.org) (141–145).

Our understanding of how genomes direct development, normal physiology, and disease in higher organisms has been hindered by a lack of suitable tools for precise and efficient gene engineering. The simple two-component CRISPRCas9system,usingWatson-Crickbasepairing by aguideRNAtoidentifytargetDNAsequences,is a versatile technology that has already stimulated innovative applications in biology. Understanding the CRISPR-Cas9 system at the biochemical and structural level allows the engineering of tailored Cas9 variants with smaller size and increased specificity. A crystal structure of the smaller Cas9 protein from Actinomyces, for example, showed how natural variation created a streamlined enzyme, setting the stage for future engineered Cas9 variants (77). A deeper analysis of the large panel of naturally evolving bacterial Cas9 enzymes may also reveal orthologs with distinct DNA binding specificity, will broaden the choice of PAMs, and will certainly reveal shorter variants more amenable for delivery in human cells.

Furthermore, specific methods for delivering Cas9 and its guide RNA to cells and tissues should benefit the field of human gene therapy. For example, recent experiments confirmed that the Cas9 protein-RNA complex can be introduced directly into cells using nucleofection or cell-penetrating peptides to enable rapid and timed editing (89,152), and transgenic organisms
that express Cas9 from inducible promoters are being tested. An exciting harbinger of future research in this area is the recent demonstration that Cas9–guide RNA complexes, when injected into adult mice, provided sufficient editing in the liver to alleviate a genetic disorder (153). Understanding the rates of homology-directed repair afterCas9-mediatedDNAcuttingwilladvancethe field by enabling efficient insertion of new or corrected sequences into cells and organisms. In addition, the rapid advance of the field has raised excitement about commercial applications of CRISPR-Cas9.

CRISPR Needle with DNA Nanoclews 

GEN 2015 Aug

A team of researchers from North Carolina State University (NC State) and the University of North Carolina at Chapel Hill (UNC-CH) have created and utilized a nanoscale vehicle composed of DNA to deliver the CRISPR-Cas9 gene editing complex into cells both in vitro and in vivo.

When the nanoclew comes into contact with a cell, the cell absorbs the nanoclew completely—swallowing it and wrapping it an endosome. Nanoclews are coated with a positively charged polymer that breaks down the endosome, setting the nanoclew free inside the cell, thus allowing CRISPR-Cas9 to make its way to the nucleus. [North Carolina State University]

• “Traditionally, researchers deliver DNA into a targeted cell to make the CRISPR RNA and Cas9 inside the cell itself—but that limits control over its dosage,” explained co-senior author Chase Beisel, Ph.D., assistant professor in the department of chemical and biomolecular engineering at NC State. “By directly delivering the Cas9 protein itself, instead of turning the cell into a Cas9 factory, we can ensure that the cell receives the active editing system and can reduce problems with unintended editing.”
• The findings from this study were published recently in Angewandte Chemie through an article entitled “Self-Assembled DNA Nanoclews for the Efficient Delivery of CRISPR-Cas9 for Genome Editing.”

• The nanoclews are made of a single, tightly-wound strand of DNA. The DNA is engineered to partially complement the relevant CRISPR RNA it will carry, allowing the CRISPR-Cas9 complex to loosely attach itself to the nanoclew. “Multiple CRISPR-Cas complexes can be attached to a single nanoclew,” noted lead author Wujin Sun, a Ph.D. student in Dr. Gu’s laboratory.
• When the nanoclew comes into contact with a cell, the cell absorbs the nanoclew completely through typical endocytic mechanisms. The nanoclews are coated with a positively charged polymer, in order to break down the endosomal membrane and set the nanoclew free inside the cell. The CRISPR-Cas9 complexes will then free themselves from the nanoclew structure to make their way to the nucleus. Once the CRISPR-Cas9 complex reaches the nucleus than the gene editing can begin.
• In order to test their delivery method, the investigators created fluorescently labeled cancer cells in culture and within mice. The CRISPR nanoclew was then designed to target the gene generating fluorescent protein in the cells—if the glowing stopped than the nanoclews worked. “And they did work. More than one-third of cancer cells stopped expressing the fluorescent protein,” Dr. Beisel stated.

Imitating Viruses to Deliver Drugs to Cells

2015 Aug – by CNRS (Délégation Paris Michel-Ange)

Figure (not shown). Assembly of the artificial virus and protein delivery: the virus consists of an initial polymer (pGi-Ni2+, left) on which the proteins to be delivered bind. It is encapsulated (right) by a second polymer (πPEI), which binds to the cell surface.

Viruses are able to redirect the functioning of cells in order to infect them. Inspired by their mode of action, scientists from the CNRS and Université de Strasbourg have designed a “chemical virus” that can cross the double lipid layer that surrounds cells, and then disintegrate in the intracellular medium in order to release active compounds. To achieve this, the team used two polymers they had designed, which notably can self-assemble or dissociate, depending on the conditions. This work, the result of collaborative efforts by chemists, biologists and biophysicists, is published in the 1st September issue of Angewandte Chemie International Edition.

Biotechnological advances have offered access to a wealth of compounds with therapeutic potential.  Many of these compounds are only active inside human cells but remain unusable because the lipid membrane surrounding these cells is a barrier they cannot cross. The challenge is therefore to find transfer solutions that can cross this barrier.

By imitating the ability of viruses to penetrate into cells, chemists in the Laboratoire de Conception et Application de Molécules Bioactives (CNRS/Université de Strasbourg) sought to design particles capable of releasing macromolecules that are only active inside cells. To achieve this, these particles must comply with several, often contradictory, constraints. They must remain stable in the extracellular medium, they must be able to bind to the cells so that they be internalized, but they must be more fragile inside the cells so that they can release their content. Using two polymers designed by the team, the scientists succeeded in creating a “chemical virus” that meets the conditions necessary for the direct delivery of active proteins into cells.

In practice, the first polymer (pGi-Ni2+) serves as a substrate for the proteins that bind to it. The second, recently patented polymer (πPEI), encapsulates this assembly thanks to its positive charges, which bind to the negative charges of pGi-Ni2+. The particles obtained (30-40 nanometers in diameter) are able to recognize the cell membrane and bind to it. This binding activates a cellular response: the nanoparticle is surrounded by a membrane fragment and enters the intracellular compartment, called the endosome. Although they remain stable outside the cell, the assemblies are attacked by the acidity that prevails within this new environment.  Furthermore, this drop in pH allows the πPEI to burst the endosome, releasing its content of active compounds.

Thanks to this assembly, the scientists were able to concentrate enough active proteins within the cells to achieve a notable biological effect. Thus by delivering a protein called caspase 3 into cancer cell lines, they succeeded in inducing 80% cell death.¹

The in vitro results are encouraging, particularly since this “chemical virus” only becomes toxic at a dose ten times higher than that used during the study. Furthermore, preliminary results in the mouse have not revealed any excess mortality. However, elimination by the body of the two polymers remains an open question. The next stage will consist in testing this method in-depth and in vivo, in animals. In the short term, this system will serve as a research tool to vectorize² recombinant and/or chemically modified proteins into cells. In the longer term, this work could make it possible to apply pharmaceutical proteins to intracellular targets and contribute to the development of innovative drugs.

This work was made possible by the collaboration of biophysicists and biologists. The skills in electron cryomicroscopy available at the Institut de Génétique et de Biologie Moléculaire et Cellulaire (CNRS/Université de Strasbourg/Inserm), and the expertise in atomic force microscopy of the Laboratoire de Biophotonique et Pharmacologie (CNRS/Université de Strasbourg) enabled highly precise characterization of the molecular assemblies. The Laboratoire Biotechnologie et Signalisation Cellulaire (CNRS/Université de Strasbourg) supplied the recombinant proteins encapsulated in the artificial virus.

A CRISPR view of development

Melissa M. Harrison,1 Brian V. Jenkins,2 Kate M. O’Connor-Giles,3,4 and Jill Wildonger2
1Department of Biomolecular Chemistry, University of Wisconsin School of Medicine and Public Health, Madison, Wisconsin 53706, USA; 2Biochemistry Department, University of Wisconsin-Madison, Madison, Wisconsin 53706, USA; 3Laboratory of Genetics, 4Laboratory of Cell and Molecular Biology, University of Wisconsin-Madison, Madison, Wisconsin 53706, USA
GENES & DEVELOPMENT 2015 Aug; 28:1859–1872
http://www.genesdev.org/cgi/doi/10.1101/gad.248252.114.

The CRISPR (clustered regularly interspaced short palindromic repeat)–Cas9 (CRISPR-associated nuclease 9) system is poised to transform developmental biology by providing a simple, efficient method to precisely manipulate the genome of virtually any developing organism. This RNA-guided nuclease (RGN)-based approach already has been effectively used to induce targeted mutations in multiple genes simultaneously, create conditional alleles, and generate endogenously tagged proteins. Illustrating the adaptability of RGNs, the genomes of >20 different plant and animal species as well as multiple cell lines and primary cells have been successfully modified. Here we review the current and potential uses of RGNs to investigate genome function during development.

Through the regulated process of development, a single cell divides and differentiates into the multitude of specialized cells that compose a mature organism. This process is controlled in large part by differential gene expression, which generates cells with distinct identities and phenotypes despite nearly identical genomes. Recent advances in genome engineering provide the opportunity to efficiently introduce almost any targeted modification in genomic DNA and, in so doing, the unprecedented ability to probe genome function during development in a diverse array of systems.

The CRISPR–Cas9 system has propelled genome editing from being a technical possibility to a practical reality for developmental biology studies due to the simplicity with which the Cas9 nuclease is recruited to a specific DNA sequence by a small, easily generated guide RNA (gRNA) that recognizes its genomic target via standard Watson-Crick base-pairing.

Cas9 enzymes from type II CRISPR–Cas systems are emerging as the sequence-specific nucleases of choice for genome engineering for several reasons. Most notably, as anRNA-guidednuclease(RGN),Cas9isguidedbyasingle gRNA that is readily engineered. In the case of the most commonly used Cas9, derived from Streptococcus pyogenes, the gRNA targeting sequence comprises 20 nucleotides (nt) that can be ordered as a pair of oligonucleotides and rapidly cloned. In contrast, generating an effective ZFN or TALEN is labor-intensive (see Box 1). ZFNs and TALENs are proteins that combine uniquely designed and generated DNA-binding sequences with the FokI nuclease cleavage domain. FokI is an obligate dimer, necessitating the generation of two novel proteins per editing experiment compared with a single gRNA for CRISPR–Cas9-mediated targeting.

Figure 1. (not shown) The flexibility and adaptability of the CRISPR–Cas9 system offers vast potential for genome manipulations. (A) Overview of the CRISPR–Cas9 system. At its simplest, the system consists of the chimeric gRNA (purple), which guides the Cas9 nuclease to the genomic target site (red). The genomic target site is composed of 20 base pairs (bp) of homology with the gRNA (red) and a PAM sequence (white). Cleavage (scissors) occurs 3 bp 59 of the PAM. (B) Components required for RGN-mediated genome editing. The CRISPR–Cas9 components can be delivered as DNA, RNA, or protein, as indicated, and introduced into the cell or embryo through injection, transfection, electroporation, or infection. Organisms and cells expressing transgenic Cas9 are available, and in Drosophila, both the transgenic Cas9-expressing strains and those expressing transgenic gRNA have been shown to increase targeting efficacy. To introduce designer mutations and/or exogenous sequence, a ssDNA or dsDNA donor template is included. (C) Genome engineering outcomes. Cas9-induced DSBs can be repaired by either NHEJ or HDR. (Top left) The DSB generated by a single gRNA can be repaired by NHEJ to generate indels. (Bottom left, dashed box) With the use of two gRNAs, NHEJ can result in larger deletions. If the gRNAs target sequences on different chromosomes, it is possible to generate chromosomal translocations and inversions. (Right) With the inclusion of a researcher-designed donor template, HDR makes it possible to generate conditional alleles (top), fluorescently or epitope tagged proteins (middle), specific mutations (bottom), or any combination thereof. The donor template can also be designed to correct a mutation in the organism or cell or replace a gene. (D) Catalytically inactive dCas9 provides a platform for probing genomic function. dCas9 can be fused to any number of different effectors to allow for the visualization of where specific DNA sequences localize, the repression or activation of transcription, or the immunoprecipitation of the bound chromatin.

Box: 1. A miniguide to genome engineering techniques

Zinc finger nucleases (ZFNs), transcriptional activator-like effector nucleases (TALENs), and CRISPR (clustered regularly interspaced short palindromic repeat)–Cas9 (CRISPR-associated nuclease 9) all function on a similar principle: A nuclease is guided to a specific sequence within the genome to induce a double strand DNA break (DSB). Once a DSB is generated, the cell’s intrinsic DNA repair machinery is set in motion, and it is during the repair of the DSB that the genome is modified. DSBs are typically repaired by either non-homologous end joining (NHEJ) or homology-directed repair (HDR) (Fig. 1C). In NHEJ, the two cleaved ends of the DSB are ligated together. During this process, DNA of varying sizes, generally on the order of a few base pairs, is occasionally inserted and/or deleted randomly. When a DSB is targeted to a coding exon, these insertions or deletions (indels) can result in a truncated gene product. If two DSBs are induced, NHEJ can generate deletions, eliminating an entire gene or region. HDR uses homologous sequence as a template to repair the DSB. Researchers can take advantage of this repair pathway to introduce designer mutations or exogenous sequence, such as genetically encoded tags, by supplying the cell with a donor DNA template that has homology with the sequence flanking the DSB. Note that cells can also use endogenous DNA as a template, in which case the DSB is repaired without incorporation of the donor-supplied edits. It is important to keep in mind that although the researcher directs where the DSB occurs in the genome, the cell is in control of how the DSB is repaired, which determines the ultimate outcome of a genome-editing experiment.

ZFNs

ZFNs are fusion proteins comprised of DNA-binding C2H2 zinc fingers fused to the nonspecific DNA cleavage domain of the nuclease Fok1 (for review, see Carroll 2011). Each zinc finger can be engineered to recognize a nucleotide triplet, and multiple (typically three to six) zinc fingers are joined in tandem to target specific genome sequences. Because the Fok1 cleavage domain must dimerize to be active, two ZFNs are required to create a DSB. This technique, which was first  successfully used in fruit flies more than a decade ago (Bibikova et al. 2002), has since been used to modify the genomes of many different organisms, including those that had not previously been developed as genetic model systems.

TALENs

Similar to ZFNs, TALENs are chimeric proteins comprised of a programmable DNA-binding domain fused to the Fok1 nuclease domain (for review, see Joung and Sander 2013). TALEs are naturally occurring proteins that are secreted by the bacteria Xanthamonas and bind to sequences in the host plant genome, activating transcription. The TALE DNA binding domain is composed of multiple repeats, each of which are 33–35 amino acids long. Each repeat recognizes a single nucleotide in the target DNA sequence. Nucleotide specificity is conferred by a two-amino-acid hypervariable region present in each repeat. Sequence-specific TALENs are generated by modifying the two residues in the hypervariable region and concatenating multiple TALE repeats together. Because the TALE DNA-binding domain is fused to Fok1, TALENs, like ZFNs, must also be used as dimers to generate DSBs.

RGNs hold great potential for dissecting how the genome functions during development. Since the CRISPR–Cas9 system has been recently described in detail elsewhere (Hsu et al. 2014; Sander and Joung 2014), we provide just a brief overview of the system (Box1; Fig.1A–C) and focus here on a few practical considerations for using RGNs to edit the genome of a developing organism.

The CRISPR–Cas9 system

The CRISPR–Cas9 genome-editing method is derived from a prokaryotic RNA-guided defense system (Gasiunas et al. 2012; Jinek et al. 2012, 2013; Cong et al. 2013; Mali et al. 2013c). CRISPR repeats were first discovered in the Escherichia coli genome as an unusual repeat locus (Ishino et al. 1987). The significance of this structure was appreciated later when investigators realized that phage and plasmid sequences are similar to the spacer sequences in CRISPR loci (Bolotin et al. 2005; Mojica et al. 2005; Pourcel et al. 2005). Soon afterward, it was shown that spacers are derived from viral genomic sequence (Barrangou et al. 2007). In the CRISPR–Cas system, short sequences (referred to as ‘‘protospacers’’) from an invading viral genome are copied as‘‘spacers’’ between repetitive sequences in the CRISPR locus of the host genome. The CRISPR locus is transcribed and processed into short CRISPR RNAs (crRNAs) that guide the Cas to the complementary genomic target sequence. There are at least eleven different CRISPR– Cas systems, which have been grouped into three major types (I–III). In the type I and II systems, nucleotides adjacent to the protospacer in the targeted genome comprise the protospacer adjacent motif (PAM). The PAM is essential for Cas to cleave its target DNA, enabling the CRISPR–Cas system to differentiate between the invading viral genome and the CRISPR locus in the host genome, which does not incorporate the PAM. For additional details on this fascinating prokaryotic adaptive immune response, see recent reviews (Sorek et al. 2013; Terns and Terns 2014). Type II CRISPR–Cas systems have been adapted as a genome-engineering tool. In this system, crRNA teams up with a second RNA, called trans-acting CRISPR RNA (tracrRNA), which is critical for crRNA maturation and recruiting the Cas9 nuclease to DNA (Deltcheva et al. 2011; Jinek et al. 2012). The RNA that guides Cas9 uses a short (;20-nt) sequence to identify its genomic target. This three-component system was simplified by fusing together crRNA and tracrRNA, creating a single chimeric ‘‘guide’’ RNA (abbreviated as sgRNA or simply gRNA) (Gasiunas et al. 2012; Jinek et al. 2012). While some early experiments indicated that a gRNA may not cleave a subset of targets as efficiently as a crRNA in combination with tracrRNA (Mali et al. 2013c), the ease of using a single RNA has led to the widespread adoption of gRNAs for genome engineering. A number of resources for designing experiments using the CRISPR–Cas9 system are freely available online. (A comprehensive list is available at http://www. geewisc.wisc.edu.)

The current methods of producing the CRISPR–Cas9 components provide great flexibility in terms of expression and delivery, and biologists can exploit these options to control when and where DSBs are generated in an organism. To introduce DSBs and generate modifications early in development, the CRISPR–Cas9 components can be injected as DNA, RNA, or protein into most developing organisms. This approach, which has been widely used, generates mosaic organisms for analysis. To gain control over which tissues are affected, a plasmid expressing Cas9 under the control of tissue-specific enhancers can be used. Since each cell has a choice of whether to repair a breakthrough NHEJ or HDR, a variety of different repair events will be present in the injected organism (and in individual cells). The frequency at which both alleles of a gene are affected has been reported to be high enough to visualize null phenotypes in developing mice and zebrafish (Jao et al. 2013; Wang et al. 2013a; Yasue et al. 2014; Yen et al. 2014).

Genome engineering with RGNs enables the direct manipulation of nearly any sequence in the genome to determine its role in development. The major limitation as to which genomic loci can be targeted is the requirement of a specific protospacer adjacent motif (PAM). The PAM is a short DNA motif adjacent to the Cas9 recognition sequence in the target DNA and is essential for cleavage. The most commonly used S. pyogenes Cas9 requires the PAM sequence 59-NGG (in cell lines, other PAMs are recognized, including 59-NAG, but at a lower frequency) (Jinek et al. 2012; Esvelt et al. 2013; Hsu et al. 2013; Jiang et al. 2013a; Zhang et al. 2014). The PAM is critical for cleavage and increases target specificity but, conversely, can also make some segments of the genome refractory to Cas9 cleavage. For example, AT-rich genomic sequences may contain fewer PAM sites that would be recognized and cleaved by S. pyogenes Cas9. Thus, some poly(dA-dT) tracts, which are implicated in nucleosome positioning (for review, see Struhl and Segal 2013), may be difficult to manipulate using S. pyogenes Cas9.

With RGNs, a variety of genomic manipulations are brought within reach of developmental biologists studying a diversity of organisms (Table 1 [nt shown]). This approach also makes it possible to readily generate mutations in different genetic strains, making it easier to control genetic background and eliminating the need to carry out multigenerational mating schemes to bring different mutations together in the same animal. While the CRISPR–Cas9 system has been widely used to introduce indels and deletions, HDR makes it possible to introduce more precise gene mutations, deletions, and exogenous sequences, such as loxP sites and green fluorescent protein (GFP).

Multiplexing advantages

Genes that have essential roles in development are often functionally redundant, and thus the effects of mutating a single gene can be masked by the presence of another gene. Due to the ease and efficiency with which gRNAs can be generated, multiple gRNAs can be used in a single experiment to simultaneously mutate multiple genes, overcoming issues of redundancy. Recent technical innovations now make it possible to express multiple gRNAs from a single transcript (Nissim et al. 2014; Tsai et al. 2014), making RGN multiplexing experiments even easier to carry out. Such multiplexing experiments will also facilitate multifaceted experiments, including epistasis tests and manipulating genes that are physically very close together in the genome. Multiplexing has already been used successfully to simultaneously disrupt both Tet1 and Tet2 in developing mice following injection into zygotes (Wang et al. 2013a). The CRISPR– Cas9 system has also been used to eliminate two genes in monkeys (Niu et al. 2014b).

Many gene products of interest to developmental biologists are essential early in development, and mutations in these genes are lethal to an animal before it reaches later developmental stages. Conditional alleles provide spatial and temporal control over gene inactivation and therefore have been invaluable tools for working with genes that cause early lethality. Conditional alleles have also been used to determine where and when a gene is acting during development. The utility of exerting conditional control over gene activity is widely recognized, and an international consortium is currently working to create a library of conditional alleles for  ~ 20,000 genes in the mouse genome (Skarnes et al. 2011). Since the expression of the conditional allele reflects the expression pattern of the recombinase, it is advantageous to have a variety of lines that express recombinase in specific tissues or at discrete developmental stages. The CRISPR– Cas9 system was recently used to generate two different Cre recombinase-expressing lines in rats (Ma et al. 2014b). Thus, RGNs are being used to rapidly generate the tools necessary to probe gene function in a tissue- and time-dependent manner.

RGNs open the door to quickly and easily tagging endogenous genes for developmental studies. Furthermore, because the CRISPR–Cas9 system is amenable to multiplexing, tags could be added simultaneously to multiple genes or different splice isoforms of a single gene. There is an ever-growing number of genetically encoded molecular tags that can be used for functional analysis, protein purification, or protein and RNA localization studies.

One of the first reportsof the use of RGNs for genome engineering demonstrated success in induced pluripotent stem cells (iPSCs) with a frequency of between 2% and 4% when assayed by deep sequencing of bulk culture (Mali et al. 2013c). Recovery of engineered cells is increased when Cas9-expressing cells are marked with a fluorescent marker and selected by cell sorting (Ding et al. 2013). Using this strategy, it was reported that clones containing at least one mutant allele could be isolated at frequencies between 51% and 79%. In comparison, TALENs designed against the same set of genes resulted in between 0% and 34% of clones containing at least one mutant allele.

The relative ease of generating mutant animals will yield many additional animal models of disease and supply a means of testing whether specific polymorphisms are the proximal cause of disease in vivo. Additionally, the CRISPR–Cas9 system is amenable to application in organisms not widely used for genetic studies. Organisms that may be better suited to mimic human disease can now be more easily used to generate disease models. For example, mouse models of the bleeding disorder von Willebrand disease fail to fully recapitulate the human disease.

Apart from point mutations and gene deletions, large chromosomal rearrangements can drive specific cancers. By simultaneously introducing gRNAs targeting two different chromosomes or two widely separated regions of the same chromosome, RGNs have been used to introduce targeted inversions and translocations into otherwise wild-type human cells (Choi and Meyerson 2014; Torres et al. 2014). These engineered cells will ultimately allow for studies of the causative role of these gene fusions in cancer progression. Translocations that drive lung adenocarcinoma (Choi and Meyerson 2014), acute myeloid leukemia, and Ewing’s sarcoma (Torres et al. 2014) have been generated in both HEK293 cells and more physiologically relevant cell types (nontransformed immortalized lung epithelial cells and human mesenchymal stem cells). Additionally, cell lines harboring chromosomal inversions found in lung adenocarcinoma have also been created (Choi and Meyerson 2014).

The first RGN based genetic screens were recently carried out in cultured mammalian cells (Koike-Yusa et al. 2014; Shalem et al. 2014; Wang et al. 2014; Zhou et al. 2014). When carrying out such a screen, it is important to consider both the number of genes targeted by the library and the degree of coverage of each gene. The largest library reported to date is comprised of 90,000 gRNAs designed to target 19,000 genes, which equates to about four to five gRNAs per targeted gene (Koike-Yusa et al. 2014).The screens identified targets affecting the DNA mismatch repair pathway (Koike-Yusa et al. 2014; Wang et al. 2014), resistance to bacterial and chemical toxins (Koike-Yusa et al. 2014; Wang et al. 2014; Zhou et al. 2014), and cell survival and proliferation (Shalem et al. 2014; Wang et al. 2014). The Zheng group (Shalem et al. 2014) also compared the results of their screen for genes involved in resistance to a drug that inhibits B-Raf with a prior RNAi screen that used the same cell line and drug. This comparison revealed that gRNAs identified targets that could be validated more consistently and efficiently than shRNAs, pointing to the potential advantages of using gRNAs to knock out, rather than knock down, gene function in genetic screens.

The question remains whether similar screens can be performed in a developing organism. Excitingly, two recent proof-of-principle studies using worms and mice indicate that RGNs will likely be useful for in vivo genetic screens, including unbiased forward genetic screens (Liu et al. 2014a; Mashiko et al. 2014).

In regards to knocking down gene expression, it remains to be determined how effective CRISPRi and dCas9 chimeras are in comparison with RNAi. Notably, CRISPRi and the dCas9 chimeras designed to inhibit gene expression are reportedly less effective in cultured mammalian cells than in bacteria (Gilbert et al. 2013). Nonetheless, given the ease with which dCas9 and TALE platforms can be programmed and their versatility, the potential application of these approaches to investigating genome dynamics in vivo is enticing to consider.

The majority of RGN-editing experiments have taken advantage of NHEJ to create small indels and larger deletions, which are useful for disrupting gene expression. However, to introduce specific mutations or other tailored modifications (e.g., genetically encoded tags), the HDR pathway must be activated. In most eukaryotic cells, DSBs are repaired more frequently through NHEJ than HDR (for review, see Lieber et al. 2003; Carroll 2014).

Pharma IQ (PiQ), 2015 Sep 1

Pharma IQ spoke to Bhuvaneish, a Post Doctorate Fellow in neurodegenerative disorders.

Bhuvaneish T.S joined the Scottish Centre for Regenerative Medicine – University of Edinburgh, almost  two years ago to establish and drive the use of CRISPR Cas9 within the University’s lab and apply it as a model for different disorders

Aim: To model motor neuron diseases using human pleuripotent stem cells

Bhuvaneish notes: “The disease modelling of neurodegenerative disorders, using human IPS (Induced Pluripotent Stem Cells), is quite challenging because of the technical variability in generating the IPS lines between different patient samples and also the varied genetic background between the donors. So this is a complex problem and leads to [difficulties when] interpreting the results and it’s also possible to generate erroneous results rather than proper scientific results because of the variations.

“One way to overcome this problem is using multiple lines for our study. So instead of using two or three patient donors, increasing their sample number to five or six, which is a tedious process.

“The other option, which [is] the ideal scenario, is to generate isogenic stem cells that differ only in the disease causing genetic variant.  So that’s where the CRISPR Cas9 comes in and it’s a quite handy tool for us.

“In a nutshell what you could do is take patients’ stem cells and then perform a gene correction in CRISPR Cas9. So now we have two types of cell, one is the mutant and the other is the gene corrected. Both are pretty much identical apart from the disease variant. It could be either a point mutation, [or] an expansion repeat, etc. This allows us to nail different phenotypes for motor neurone disorders.

“So generally we generate motor neurones from these two lines and model the disease in a dish, which also helps us to understand the mechanism of the disease.”

Bhuvaneish’s lab also generates different knock outs, which is highly efficient with the CRISPR technique.

Challenges with CRISPR Cas9

With Bhuvaneish leading the use of this technique in the lab, he encountered various challenges regarding the delivery system into the stem cells.

These challenges include off target effects and the efficiency of CRISPR Cas9.

On the latter point, he explains: “Although people say that the efficiency of CRISPR is much better than other gene editing systems like TAL effectors or zinc fingers, it is still pretty low. I mean, the efficiencies you are talking about is 2%, so it is still low.

He continues:  “These are the two challenges which we have and I think it’s a challenge the entire world has at the moment with this technology. And we’ve been trying to increase efficiencies with certain drugs, which has also been published recently. I haven’t got any data to back it up myself but looks promising, though.”

“So that itself is a really good thing because now I can dissect the disease causing phenotypes which we see in our culture and that has been reversed after gene correction. You can completely reverse the phenotype. So that itself is proof of concept that the disease causing the mutation is causing this phenotype.”

“In the research field it’s a really, really important tool but for gene therapy as a therapeutic we are still very behind because of the ethical issues.  The big challenge is in how to deliver these Cas9 proteins and the guide RNAs to the required donor. It could be that the disease has affected only one particular organ rather than the whole body so you would try to target those particular organs. And it’s a challenge in delivering those Cas9 and the guide RNAs to the particular organ because it’s quite a huge protein compared to conventional proteins which have been used for gene therapy.

“Although it’s highly efficient when compared to the others, for therapeutics we need precise targeting with very, very minimal off target mutations. So that would be CRISPR’s bottleneck coming into the medicine field as a therapeutic.

“For the research it is great at the moment. It has enabled most of the researchers to do the genome editing in human stem cells, which was virtually impossible before.”

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