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ATCC Announces First Isogenic Cell Line Produced by the CRISPR/Cas9 Technology
Reporter: Stephen J. Williams, PhD.
EML4-ALK Isogenic Cells — New!
ATCC is proud to announce its first product developed using CRISPR/Cas9 technology, the EML4-ALK Fusion-A549 Isogenic Cell Line Human (ATCC® CCL-185IG™). This cell line was derived from the parental A549 (ATCC® CCL-185™) non-small cell lung cancer cell line. EML4-ALK Fusion-A549 Isogenic Cell Line has been intensively validated on the genome, transcript, and protein level, and is otherwise identical to the parental line. This isogenic cell line is more sensitive to ALK inhibitor crizotinib when compared to A549, and serves as a vital model to study cell signaling pathways in cancer as well as in drug screening when used side-by-side with A549 cells.
Further your lung cancer research with the EML4-ALK Fusion-A549 Isogenic Cell Line Human ATCC® CCL-185IG™ derived from A549 ATCC ® CCL-185™ today!
Lung Cancer
Lung cancers are classified by type: small cell lung carcinoma (SCLC) and non-small cell lung carcinoma (NSCLC). SCLCs are associated with smoking and metastasize very early. By contrast, non-smokers usually present with NSCLC, which are further subdivided into squamous cell carcinomas, adenocarcinomas, and large cell carcinomas. Since both SCLC and NSCLC are usually diagnosed after the disease has spread beyond the primary site, the overall survival rates for lung cancers are poor. To breathe new life into your lung cancer research, ATCC provides numerous lung cancer cell lines, a new gene-edited isogenic NSCLC cell line, human primary cells, and h-TERT-immortalized cell lines. And to increase the throughput of your lung cancer experiments, ATCC has lung cancer cell lines organized into tumor cell panels.
All experiments should include physiologically relevant controls. ATCC provides both primary and hTERT-immortalized bronchial epithelial cells and small airway cells that may be used side-by-side with NSCLC or SCLC cells as normal controls. The primary and hTERT-immortalized cells may also be used to create 3D cell culture models to better represent an in vivo environment, ex vivo.
Cambridge Healthtech Institute’s Second Annual New Frontiers in Gene Editing
Reporter: Stephen J. Williams, PhD
Gene editing is rapidly progressing from being a research/screening tool to one that promises important applications downstream in drug development, cell therapy and bioprocessing. Cambridge Healthtech Institute’s second annual symposium on New Frontiers in Gene Editing will bring together experts from all aspects of basic science and clinical research to talk about the progress being made in gene editing and how it’s being applied. Knowing the strengths and limitations of the different tools, how does one decide when to use the CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas system, as opposed to Transcription Activator-Like Effector Nucleases (TALENs), zinc finger nucleases (ZFNs) and other systems? What is being done to overcome some of the inherent challenges with design, delivery and off-target effects, associated with each of these techniques? Experts from pharma/biotech, academic and government labs will share their experiences leveraging the utility of gene editing for diverse applications.
Striving for better design, targeted delivery and performance
CRISPR Screening for drug target identification
Gene editing in stem cells
Gene editing for cell therapy and regenerative medicine
Understanding the pitfalls of gene editing
Dealing with off-target effects
Interested in Gene Editing? You may also want to attend our focused short course:
Cambridge Healthtech Institute 250 First Avenue, Suite 300 | Needham, MA 02494 | 781-972-5400 | www.healthtech.com
2.1.5.26 Cambridge Healthtech Institute’s Second Annual New Frontiers in Gene Editing, SF, 3/10-3/11, 2016, 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
FDA Cellular & Gene Therapy Guidances: Implications for CRSPR/Cas9 Trials, 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
FDA Cellular & Gene Therapy Guidances: Implications for CRSPR/Cas9 Trials
Reporter: Stephen J. Williams, PhD
The recent announcement by Editas CEO Katrine Bosley to pursue a CRSPR/Cas9 gene therapy trial to correct defects in an yet to be disclosed gene to treat one form of a rare eye disease called Leber congenital amaurosis (multiple mutant genes have been linked to the disease) have put an interesting emphasis on the need for a regulatory framework to initiate these trials. Indeed at the 2015 EmTechMIT Conference Editas CEO Katrine Bosley had mentioned this particular issue: the need for discourse with FDA and regulatory bodies to establish guidelines for design of clinical trials using the CRSPR gene editing tool.
To this effect, I have listed below, the multiple FDA Guidance Documents surrounding gene therapy to show that, in the past year, the FDA has shown great commitment to devise a regulatory framework for this therapeutic area.
New CRISPR-non Cas9 proteins, 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
New CRISPR-non Cas9 proteins
Larry H. Bernstein, MD, FCAP, Curator
LPBI
More CRISPR Proteins Discovered
Researchers identify three new proteins that may serve as alternatives to Cas9.
Crystal structure of a Cas9 in complex with an RNA guide and a stretch of target DNAWIKIMEDIA, H. NISHIMASU ET AL.
Scouring genomic databases for sequences with similarity to the components of the CRISPR/Cas9 system and the recently identified CRISPR/Cpf1 system, researchers from the National Center for Biotechnology Information (NCBI), MIT, and Rutgers University have discovered three novel CRISPR systems that could one day provide new gene-editing tools to supplement the currently used CRISPR/Cas9 system. The newly discovered CRISPR systems contain three new proteins, C2c1, C2c2, and C2c3 (named for “Class 2 candidate x”), one of which may cleave RNA.
“This work shows a path to discovery of novel CRISPR/Cas systems with diverse properties, which are demonstrated here in direct experiments,” coauthor Eugene Koonin of NCBI told GenomeWeb. “The most remarkable aspect of the story is how evolution has achieved a broad repertoire of biological activities, a feat we can take advantage of for new genome manipulation tools.” The group published its results yesterday (October 22) in Molecular Cell.
Using such sequence-based techniques, the researchers predict that there are even more CRISPR systems to be discovered, added study coauthor Konstantin Severinov of Rutgers. “There are multiple ways to modify the search algorithm. So more exciting and distinct CRISPR/Cas mechanisms should be expected soon.”
Using CRISPR as a High-Throughput Cancer Screening and Modeling Tool
Using CRISPR/Cas9, scientists created a new high-throughput screening tool for studying the development and progression of liver cancer in mice. [Ernesto del Aguila III, NHGRI]
A contingent of researchers from the UK, Germany, and Spain have recently developed a novel CRISPR/Cas9 system that they believe can be utilized as a multiplexed screening approach to study and model cancer development in mice. In the current study, the investigators directly mutated genes within adult mouse livers to elucidate their role in cancer development and progression—simultaneously uncovering the gene combinations that coordinate to cause liver cancer.
“We reasoned that, by targeting mutations directly to adult liver cells using CRISPR/Cas9, we could better study and understand the biology of this important cancer,” explained co-author Mathias Friedrich, Ph.D., research scientist at the Wellcome Trust Sanger Institute. “Other approaches to engineer mutations in mice, such as stem cell manipulation, are limited by the laborious process, the long time frames and large numbers of animals needed. And, our method better mimics important aspects of human cancer biology than many “classic” mouse models: as in most human cancers, the mutations occur in the adult and only affect a few cells”.
The findings from this study were published online recently in PNAS through an article entitled “CRISPR/Cas9 somatic multiplex-mutagenesis for high-throughput functional cancer genomics in mice.”
This new approach is rapid, scalable, and extremely efficient, allowing the researchers to examine an array of genes or large regions of the genome concurrently. Moreover, this methodology affords scientists the ability to distinguish between cancer driver mutations and passenger mutations—those that occur as side-effects of cancer development.
The research team developed a list of up to eighteen genes with known or unknown evidence for their importance in two forms of liver cancer. They then introduced the CRISPR/Cas9 molecules, targeting various combinations of these genes into mice, which subsequently developed liver or bile duct cancer within a few months.
“Our approach enables us to simultaneously target multiple putative genes in individual cells,” noted co-author Roland Rad, Ph.D., project leader at the Technical University of Munich and the German Cancer Research Center Heidelberg. “We can now rapidly and efficiently screen which genes are cancer-causing and which ones are not. And, we can study how genes work together to cause cancers—a crucial piece of the puzzle we must solve to understand and tackle the disease.”
The investigators were able to confirm that a set of DNA-binding proteins called ARID (AT-rich interactive domain), influence the organization of chromosomes and are important for liver cancer development. Furthermore, mutations in a second protein, TET2, were found to be causative in bile duct cancer: although TET2 has not been found to be mutated in human biliary cancers, the proteins that it interacts with have been, showing that the CRISPR/Cas9 method can identify human cancer genes that are not mutated, but whose function is disturbed by other events.
“The new tools of targeting genes in combination and inducing insertions or deletions in chromosomes change our ability to identify new cancer-causing genes and to understand their role in cancer,” stated senior group leader and co-author Allan Bradley, Ph.D., director emeritus from the Sanger Institute. “Our results show that this approach is feasible and productive in liver cancer; we will now continue to study our new findings and try to extend the approach to other cancer types.”
This CRISPR/Cas9 approach may also be favorable for an in-depth examination of genomic deserts —regions within the human genome that appear to be devoid of genes. Yet, recent data from the ENCODE Project suggests that deserts can be populated, if not by genes, then by DNA regulatory regions that influence the activity of genes.
“Liver cancer has many DNA alterations in regions lacking genes: we don’t know which of these might be important for the disease,” said Dr. Rad. “However, we could show that it is now possible to delete such regions to systematically determine their role in liver cancer development.”
Three newly discovered enzymes—provisionally named C2c1, C2c3, and C2c3—promise to expand the CRISPR genome-editing toolbox beyond the well-known Cas9. They had been hiding inside NIH genomic databases, but they were eventually found out, thanks to the application of computational approaches developed by two groups of researchers. These researchers also initiated experimental work to explore the function of the bioinformatically identified enzymes.
One of the groups was led by Eugene Koonin, Ph.D., senior investigator at the National Center for Biotechnology Information (NCBI), National Library of Medicine (NLM), part of the NIH. The other group was led by Feng Zhang, Ph.D., of the MIT-Harvard Broad Institute.
According to the researchers, the three newly discovered enzymes are all naturally occurring, and all share some features with Cas9. In addition, these three enzymes have unique properties that could be exploited for novel genome-editing applications.
“This work shows a path to discovery of novel CRISPR-Cas systems with diverse properties, which are demonstrated here in direct experiments,” said Dr. Koonin. “The most remarkable aspect of the story is how evolution has achieved a broad repertoire of biological activities, a feat we can take advantage of for new genome-manipulation tools.”
This comment highlights how the researchers’ work, which appeared October 22 in the journal Molecular Cell, included information about potential evolutionary pathways. The researchers also emphasized that their work might lead to additional enzyme discoveries.
“There are multiple ways to modify the search algorithm, so more exciting and distinct CRISPR-Cas mechanisms should be expected soon,” said Konstantin Severinov, Ph.D., one of the researchers. He is affiliated with Rutgers and the Skolkovo Institute of Science and Technology. “These new mechanisms will undoubtedly attract the attention of basic and applied scientists alike.”
The Koonin and Zhang groups also recently collaborated on a project that resulted in the characterization of Cpf1, a class II CRISPR endonuclease, like Cas9. This work was described last month in an article, published in Cell, suggesting that the newly found enzyme’s distinct features pointed to unique genome-editing possibilities.
In his comments about this earlier work, Dr. Zhang made a point that presaged the current work: “Our goal is to develop tools that can accelerate research and eventually lead to new therapeutic applications. We see much more to come, even beyond Cpf1 and Cas9, with other enzymes that may be repurposed for further genome-editing advances.”
2.1.5.27 CRISPR/Cas-mediated Genome Engineering, 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
Cpf1 Is a Single RNA-Guided Endonuclease of a Class 2 CRISPR-Cas System
•Cpf1 is a CRISPR-associated two-component RNA-programmable DNA nuclease
•Targeted DNA is cleaved as a 5-nt staggered cut distal to a 5′ T-rich PAM
•Two Cpf1 orthologs exhibit robust nuclease activity in human cells
Summary
The microbial adaptive immune system CRISPR mediates defense against foreign genetic elements through two classes of RNA-guided nuclease effectors. Class 1 effectors utilize multi-protein complexes, whereas class 2 effectors rely on single-component effector proteins such as the well-characterized Cas9. Here, we report characterization of Cpf1, a putative class 2 CRISPR effector. We demonstrate that Cpf1 mediates robust DNA interference with features distinct from Cas9. Cpf1 is a single RNA-guided endonuclease lacking tracrRNA, and it utilizes a T-rich protospacer-adjacent motif. Moreover, Cpf1 cleaves DNA via a staggered DNA double-stranded break. Out of 16 Cpf1-family proteins, we identified two candidate enzymes from Acidaminococcus and Lachnospiraceae, with efficient genome-editing activity in human cells. Identifying this mechanism of interference broadens our understanding of CRISPR-Cas systems and advances their genome editing applications.
One-Step Generation of Mice Carrying Mutations in Multiple Genes by CRISPR/Cas-Mediated Genome Engineering
Haoyi Wang6, Hui Yang6, Chikdu S. Shivalila6, Meelad M. Dawlaty, Albert W. Cheng, Feng Zhang, Rudolf Jaenisch
Cell May 2013; 153: 4(9), p910–918 http://dx.doi.org/10.1016/j.cell.2013.04.025
Highlights
•CRISPR/Cas9-mediated simultaneous targeting of five genes in mES cells
•Generation of Tet1/Tet2 double-mutant mice in one step
•Generation of Tet1/Tet2 double-mutant mice with predefined mutations in one step
Summary
Mice carrying mutations in multiple genes are traditionally generated by sequential recombination in embryonic stem cells and/or time-consuming intercrossing of mice with a single mutation. The CRISPR/Cas system has been adapted as an efficient gene-targeting technology with the potential for multiplexed genome editing. We demonstrate that CRISPR/Cas-mediated gene editing allows the simultaneous disruption of five genes (Tet1, 2, 3, Sry, Uty – 8 alleles) in mouse embryonic stem (ES) cells with high efficiency. Coinjection of Cas9 mRNA and single-guide RNAs (sgRNAs) targeting Tet1 and Tet2 into zygotes generated mice with biallelic mutations in both genes with an efficiency of 80%. Finally, we show that coinjection of Cas9 mRNA/sgRNAs with mutant oligos generated precise point mutations simultaneously in two target genes. Thus, the CRISPR/Cas system allows the one-step generation of animals carrying mutations in multiple genes, an approach that will greatly accelerate the in vivo study of functionally redundant genes and of epistatic gene interactions.
Generating genetically modified mice using CRISPR/Cas-mediated genome engineering
Mice with specific gene modifications are valuable tools for studying development and disease. Traditional gene targeting in mice using embryonic stem (ES) cells, although suitable for generating sophisticated genetic modifications in endogenous genes, is complex and time-consuming. We have recently described CRISPR/Cas-mediated genome engineering for the generation of mice carrying mutations in multiple genes, endogenous reporters, conditional alleles or defined deletions. Here we provide a detailed protocol for embryo manipulation by piezo-driven injection of nucleic acids into the cytoplasm to create gene-modified mice. Beginning with target design, the generation of gene-modified mice can be achieved in as little as 4 weeks. We also describe the application of the CRISPR/Cas technology for the simultaneous editing of multiple genes (five genes or more) after a single transfection of ES cells. The principles described in this protocol have already been applied in rats and primates, and they are applicable to sophisticated genome engineering in species in which ES cells are not available.
Our long-range goals are to understand epigenetic regulation of gene expression in mammalian development and disease. An important question is to understand the different epigenetic conformations that distinguish differentiated cell states and to define strategies to transdifferentiate one differentiated cell type into another. Embryonic stem cells are of major significance because they have the potential to generate any cell type in the body and, therefore, are of great interest for regenerative medicine. A major focus of our work is to understand the molecular mechanisms that allow the reprogramming of somatic cells to an embryonic pluripotent state and to use the potential of patient specific pluripotent cells to study complex human diseases.
One-Step Generation of Mice Carrying Mutations in Multiple Genes by CRISPR/Cas-Mediated Genome Engineering
CRISPR/Cas9-mediated simultaneous targeting of five genes in mES cells
Generation of Tet1/Tet2 double-mutant mice in one step
Generation of Tet1/Tet2 double-mutant mice with predefined mutations in one step
Mice carrying mutations in multiple genes are traditionally generated by sequential recombination in embryonic stem cells and/or time-consuming intercrossing of mice with a single mutation. The CRISPR/Cas system has been adapted as an efficient gene-targeting technology with the potential for multiplexed genome editing. We demonstrate that CRISPR/Cas-mediated gene editing allows the simultaneous disruption of five genes (Tet1, 2, 3, Sry, Uty – 8 alleles) in mouse embryonic stem (ES) cells with high efficiency. Coinjection of Cas9 mRNA and single-guide RNAs (sgRNAs) targeting Tet1 and Tet2 into zygotes generated mice with biallelic mutations in both genes with an efficiency of 80%. Finally, we show that coinjection of Cas9 mRNA/sgRNAs with mutant oligos generated precise point mutations simultaneously in two target genes. Thus, the CRISPR/Cas system allows the one-step generation of animals carrying mutations in multiple genes, an approach that will greatly accelerate the in vivo study of functionally redundant genes and of epistatic gene interactions.
Genetically modified mice represent a crucial tool for understanding gene function in development and disease. Mutant mice are conventionally generated by insertional mutagenesis (Copeland and Jenkins, 2010; Kool and Berns, 2009) or by gene-targeting methods (Capecchi, 2005). In conventional gene-targeting methods, mutations are introduced through homologous recombination in mouse embryonic stem (ES) cells. Targeted ES cells injected into wild-type (WT) blastocysts can contribute to the germline of chimeric animals, generating mice containing the targeted gene modification (Capecchi, 2005). It is costly and time consuming to produce single-gene knockout mice and even more so to make double-mutant mice. Moreover, in most other mammalian species, no established ES cell lines are available that contribute efficiently to chimeric animals, which greatly limits the genetic studies in many species.
Alternative methods have been developed to accelerate the process of genome modification by directly injecting DNA or mRNA of site-specific nucleases into the one-cell embryo to generate DNA double-strand break (DSB) at a specified locus in various species (Bogdanove and Voytas, 2011; Carroll et al., 2008; Urnov et al., 2010). DSBs induced by these site-specific nucleases can then be repaired by error-prone nonhomologous end joining (NHEJ) resulting in mutant mice and rats carrying deletions or insertions at the cut site (Carbery et al., 2010; Geurts et al., 2009; Sung et al., 2013;Tesson et al., 2011). If a donor plasmid with homology to the ends flanking the DSB is coinjected, high-fidelity homologous recombination can produce animals with targeted integrations (Cui et al., 2011; Meyer et al., 2010). Because these methods require the complex designs of zinc finger nucleases (ZNFs) or Transcription activator-like effector nucleases (TALENs) for each target gene and because the efficiency of targeting may vary substantially, no multiplexed gene targeting in animals has been reported to date. To dissect the functions of gene family members with redundant functions or to analyze epistatic relationships in genetic pathways, mice with two or more mutated genes are required, prompting the development of efficient technology for the generation of animals carrying multiple mutated genes.
Recently, the type II bacterial CRISPR/Cas system has been demonstrated as an efficient gene-targeting technology with the potential for multiplexed genome editing. Bacteria and archaea have evolved an RNA-based adaptive immune system that uses CRISPR (clustered regularly interspaced short palindromic repeat) and Cas (CRISPR-associated) proteins to detect and destroy invading viruses and plasmids (Horvath and Barrangou, 2010; Wiedenheft et al., 2012). Cas proteins, CRISPR RNAs (crRNAs), andtrans-activating crRNA (tracrRNA) form ribonucleoprotein complexes, which target and degrade foreign nucleic acids, guided by crRNAs ( Gasiunas et al., 2012; Jinek et al., 2012). It was shown that the Cas9 endonuclease from Streptococcus pyogenes type II CRISPR/Cas system can be programmed to produce sequence-specific DSB in vitro by providing a synthetic single-guide RNA (sgRNA) consisting of a fusion of crRNA and tracrRNA ( Jinek et al., 2012). More intriguingly, Cas9 and sgRNA are the only components necessary and sufficient for induction of targeted DNA cleavage in cultured human cells ( Cho et al., 2013; Cong et al., 2013; Mali et al., 2013) as well as in zebrafish (Chang et al., 2013; Hwang et al., 2013). A recent report also demonstrated disruption of a GFP transgene in mice using the CRISPR/Cas system ( Shen et al., 2013). The ease of design, construction, and delivery of multiple sgRNAs suggest the possibility of multiplexed genome editing in mammals. Indeed, one study demonstrated that two loci separated by 119 bp could be cleaved simultaneously in cultured human cells at a low efficiency ( Cong et al., 2013). The extent of achievable multiplexed genome editing has yet to be demonstrated in stem cells as well as in animals. Here, we use the CRISPR/Cas system to drive both NHEJ-based gene disruption and homology directed repair (HDR)-based precise gene editing to achieve highly efficient and simultaneous targeting of multiple genes in stem cells and mice.
Simultaneous Targeting up to Five Genes in ES Cells
To test whether the CRISPR/Cas system could produce targeted cleavage in the mouse genome, we transfected plasmids expressing both the mammalian-codon-optimized Cas9 and a sgRNA targeting each gene ( Cong et al., 2013; Mali et al., 2013) into mouse ES cells and determined the targeted cleavage efficiency by the Surveyor assay ( Guschin et al., 2010). All three Cas9-sgRNA transfections produced cleavage at target loci with high efficiency of 36% at Tet1, 48% atTet2, and 36% at Tet3 ( Figure 1B). Because each target locus contains a restriction enzyme recognition site ( Figure 1A), we PCR amplified an ∼500 bp fragment around each target site and digested the PCR products with the respective enzyme. A correctly targeted allele will lose the restriction site, which can be detected by failure to cleave upon enzyme treatment. Using this restriction fragment length polymorphism (RFLP) assay, we screened 48 ES cell clones from each single-targeting experiment. Consistent with the Surveyor analysis, a high percentage of ES cell clones were targeted, with a high probability of having both alleles mutated ( Figure S1A available online). The results summarized in Table 1 demonstrate that between 65% and 81% of the tested ES cell clones carried mutations in the Tet genes with up to 77% having mutations in both alleles.
Figure 1.
Multiplexed Gene Targeting in mouse ES cells
(A) Schematic of the Cas9/sgRNA-targeting sites in Tet1, 2, and 3. The sgRNA-targeting sequence is underlined, and the protospacer-adjacent motif (PAM) sequence is labeled in green. The restriction sites at the target regions are bold and capitalized. Restriction enzymes used for RFLP and Southern blot analysis are shown, and the Southern blot probes are shown as orange boxes.
(B) Surveyor assay for Cas9-mediated cleavage at Tet1, 2, and 3 loci in ES cells.
(C) Genotyping of triple-targeted ES cells, clones 51, 52, and 53 are shown. Upper: RFLP analysis. Tet1PCR products were digested with SacI, Tet2 PCR products were digested with EcoRV, and Tet3 PCR products were digested with XhoI. Lower: Southern blot analysis. For the Tet1 locus, SacI digested genomic DNA was hybridized with a 5′ probe. Expected fragment size: WT = 5.8 kb, TM (targeted mutation) = 6.4 kb. For the Tet2 locus, SacI, and EcoRV double-digested genomic DNA was hybridized with a 3′ probe. Expected fragment size: WT = 4.3 kb, TM = 5.6 kb. For the Tet3 locus, BamHI and XhoI double-digested genomic DNA was hybridized with a 5′ probe. Expected fragment size: WT = 3.2 kb, TM = 8.1 kb.
(D) The sequence of six mutant alleles in triple-targeted ES cell clone 14 and 41. PAM sequence is labeled in red.
(E) Analysis of 5hmC levels in DNA isolated from triple-targeted ES cell clones by dot blot assay using anti-5hmC antibody. A previously characterized DKO clone derived using traditional method is used as a control. See also Figure S1.
Single-, Triple-, and Quintuple-Gene Targeting in mES Cells, Related to Figure 1
(A) RFLP analysis of clones from each single-targeting experiment (1 to 17 are shown).
(B) RFLP analysis of triple-gene-targeted clones (37 to 53 are shown). Tet1 PCR products were digested with SacI, Tet2 PCR products were digested with EcoRV, and Tet3 PCR products were digested with XhoI. WT control is shown in the last lane. Genotyping of clone 51, 52, and 53 are also shown in Figure 1C.
(C) Schematic of the Cas9/sgRNA-targeting sites in Sry and Uty. The sgRNA-targeting sequence is underlined, and the protospacer-adjacent motif (PAM) sequence is labeled in green. The restriction sites at the target regions are bold and capitalized. Restriction enzymes used for RFLP analysis are shown.
(D) RFLP analysis of quintuple-gene-targeted clones (1 to 10 are shown). Sry PCR products were digested with BsaJI, Uty PCR products were digested with AvrII. WT control is shown in the last lane. RFLP analysis of Tet1, 2, 3 loci are not shown.
CRISPR/Cas-Mediated Gene Targeting in V6.5 ES Cells
Mutant Alleles per Clone / Total Clones Tested
Gene
6
5
4
3
2
1
0
Tet1
N/A
27/48
4/48
17/48
Tet2
37/48
2/48
9/48
Tet3
32/48
3/48
13/48
Tet1+ Tet2 + Tet3
20/96
16/96
2/96
2/96
1/96
0/96
55/96
Plasmids encoding Cas9 and sgRNAs targeting Tet1, Tet2, and Tet3 were transfected separately (single targeting) or in a pool (triple targeting) into ES cells. The number of total alleles mutated in each ES cell clone is listed from 0 to 2 for single-targeting experiment, and 0 to 6 for triple-targeting experiment. The number of clones containing each specific number of mutated alleles is shown in relation to the total number of clones screened in each experiment. See also Table S1.
The high efficiency of single-gene modification prompted us to test the possibility of targeting all three genes simultaneously. For this we cotransfected ES cells with the constructs expressing Cas9 and three sgRNAs targeting Tet1, 2, and 3. Of 96 clones screened using the RFLP assay, 20 clones were identified as having mutations in all six alleles of the three genes ( Figures 1C and S1B and Table 1). To exclude that a PCR bias could give false positive results, we performed Southern blot analysis and confirmed complete agreement with the RFLP results ( Figure 1C). We subcloned and sequenced the PCR products of Tet1-, Tet2-, and Tet3-targeted regions to verify that all of eight tested clones carried biallelic mutations in all three genes with most clones displaying two mutant alleles for each gene with small insertions or deletions (indels) at the target site ( Figure 1D). To test whether these mutant alleles would abolish the function of Tet proteins, we compared the 5hmC level of targeted clones to WT ES cells. Previously, we reported a depletion of 5hmC in Tet1/Tet2 double-knockout ES cells derived using traditional gene-targeting methods ( Dawlaty et al., 2013). As expected from loss of function alleles, we found a significant reduction of 5hmC levels in all clones carrying biallelic mutations in the three genes ( Figure 1E).
To further test the potential of multiplexed gene targeting by CRISPR/Cas system, we designed sgRNAs targeting two Y-linked genes, Sry and Uty ( Figure S1C). Short PCR products encoding sgRNAs targeting all five genes (Tet1, Tet2, Tet3, Sry, and Uty) were pooled and cotransfected with a Cas9 expressing plasmid and the PGK puroR cassette into ES cells. Of 96 clones that were screened using the RFLP assay, 10% carried mutations in all eight alleles of the five genes ( Figure S1D and Table S1), demonstrating the capacity of the CRISP/Cas9 system for highly efficient multiplexed gene targeting.
One-Step Generation of Single-Gene Mutant Mice by Zygote Injection
We tested whether mutant mice could be generated in vivo by direct embryo manipulation. Capped polyadenylated Cas9 mRNA was produced by in vitro transcription and coinjected with sgRNAs. Initially, to determine the optimal concentration of Cas9 mRNA for targeting in vivo, we microinjected varying amounts of Cas9-encoding mRNA with Tet1 targeting sgRNA at constant concentration (20 ng/μl) into pronuclear (PN) stage one-cell mouse embryos and assessed the frequency of altered alleles at the blastocyst stage using the RFLP assay. As expected, higher concentration of Cas9 mRNA led to more efficient gene disruption ( Figure S2A). Nevertheless, even embryos injected with the highest amount of Cas9 mRNA (200 ng/μl) showed normal blastocyst development, suggesting low toxicity.
Figure S2.
One-Step Generation of Single-Gene Mutant Mice by Zygote Injection, Related to Figure 2
(A) RFLP analysis of blastocysts injected with different concentration of Cas9 mRNA and Tet1 sgRNA at 20 ng/μl. Tet1 PCR products were digested with SacI.
(B) Commonly recovered Tet1 and Tet2 alleles resulted from MMEJ. PAM sequence of each targeting sequence is labeled in green. Microhomology flanking the DSB is bold and underlined in WT sequence.
(C) RFLP analysis of eight Tet3-targeted blastocysts demonstrated high targeting efficiency (embryo 3 and 5 failed to amplify). Tet3 PCR products were digested with XhoI.
(D) Some Tet3-targeted mice show smaller size and all homozygous mutants died within 1 day after birth.
(E) RFLP analysis of Tet3 single-targeted newborn mice. Mouse 8 and 14 survived after birth. Sample 2 and 6 failed to amplify.
(F) Sequences of both Tet3 alleles of surviving Tet3-targeted mouse 14. PAM sequences are labeled in red.
To investigate whether postnatal mice carrying targeted mutations could be generated, we coinjected sgRNAs targeting Tet1or Tet2 with different concentrations of Cas9 mRNA. Blastocysts derived from the injected embryos were transplanted into foster mothers and newborn pups were obtained. As summarized in Table 2, about 10% of the transferred blastocysts developed to birth independent of the RNA concentrations used for injection suggesting low fetal toxicity of the Cas9 mRNA and sgRNA. RFLP, Southern blot, and sequencing analysis demonstrated that between 50 and 90% of the postnatal mice carried biallelic mutations in either target gene ( Figures 2A, 2B, and 2C and Table 2).
Table 2.
CRISPR/Cas-Mediated Single-Gene Targeting in BDF2 Mice
Cas9 mRNA and sgRNAs targeting Tet1, Tet2, or Tet3 were injected into fertilized eggs. The blastocysts derived from injected embryos were transplanted into foster mothers and newborn pups were obtained and genotyped. The number of total alleles mutated in each mouse is listed from 0 to 2. The number of mice containing each specific number of mutated alleles is shown in relation to the total number of mice screened in each experiment. See also Table S2. A Some of the pups were cannibalized.
Surprisingly, specific Δ9 Tet1 and specific Δ8 and Δ15 Tet2 mutant alleles were repeatedly recovered in independently derived mice. Preferential generation of these alleles is likely caused by a short sequence repeat flanking the DSB (see Figure S2B) consistent with previous reports demonstrating that perfect microhomology sequences flanking the cleavage sites can generate microhomology-mediated precise deletions by end repair mechanism (MMEJ) ( McVey and Lee, 2008; Symington and Gautier, 2011) (Figure S2B). A similar observation was also made when TALEN mRNA was injected into one-cell rat embryos ( Tesson et al., 2011).
We also derived blastocysts from zygotes injected with Cas9 mRNA and Tet3 sgRNA. Genotyping of the blastocysts demonstrated that of eight embryos three were homozygous and three were heterozygous Tet3 mutants (two failed to amplify) (Figure S2C). Some blastocysts were implanted into foster mothers and, upon C section, we readily identified multiple mice of smaller size ( Figure S2D), many of which died soon after delivery. Genotyping shown in Figure S2E indicated that all pups with mutations in both Tet3 alleles died neonatally. Only 2 out of 15 mice survived that were either Tet3heterozygous mutants or WT ( Figure S2F). These results are consistent with the lethal neonatal phenotype of Tet3 knockout mice generated using traditional methods ( Gu et al., 2011), although we have not yet established which of the Tet3 mutations produced loss of function rather than hypomorphic alleles.
One-Step Generation of Double-Gene Mutant Mice by Zygote Injection
To test whether Tet1/Tet2 double-mutant mice could be produced from single embryos, we coinjected Tet1 and Tet2 sgRNAs with 20 or 100 ng/μl Cas9 mRNA into zygotes. A total of 28 pups were born from 144 embryos transferred into foster mothers (21% live-birth rate) that had been injected at the zygote stage with high concentrations of RNA (Cas9 mRNA at 100 ng/μl, sgRNAs at 50 ng/μl), consistent with low or no toxicity of the Cas9 mRNA and sgRNAs ( Table 3). RFLP, Southern blot analysis, and sequencing identified 22 mice carrying targeted mutations at all four alleles of the Tet1 and Tet2genes ( Figures 2D and 2E) with the remaining mice carrying mutations in a subset of alleles ( Table 3). Injection of zygotes with low concentration of RNA (Cas9 mRNA at 20 ng/μl, sgRNAs at 20 ng/μl) yielded 19 pups from 75 transferred embryos (25% live-birth rate), which is a higher survival rate than from embryos injected with 100 ng/μl of Cas9 RNA. Nevertheless, more than 50% of the pups were biallelic Tet1/Tet2 double mutants ( Table 3). These results demonstrate that postnatal mice carrying biallelic mutations in two different genes can be generated within one month with high efficiency (Figure 2F).
Table 3.
CRISPR/Cas-Mediated Double-Gene Targeting in BDF2 Mice
Cas9 mRNA and sgRNAs targeting Tet1and Tet2 were coinjected into fertilized eggs. The blastocysts derived from the injected embryos were transplanted into foster mothers and newborn pups were obtained and genotyped. The number of total alleles mutated in each mouse is listed from 0 to 4 for Tet1 and Tet2. The number of mice containing each specific number of mutated alleles is shown in relation to the number of total mice screened in each experiment. A Some of the pups were cannibalized.
Although the high live-birth rate and normal development of mutant mice suggest low toxicity of CRISPR/Cas9 system, we sought to determine the off-target effects in vivo. Previous work in vitro, in bacteria, and in cultured human cells suggested that the protospacer-adjacent motif (PAM) sequence NGG and the 8 to 12 base “seed sequence” at the 3′ end of the sgRNA are most important for determining the DNA cleavage specificity (Cong et al., 2013; Jiang et al., 2013; Jinek et al., 2012). Based on this rule, only three and four potential off targets exist in mouse genome for Tet1 and Tet2 sgRNA, respectively ( Table S2 and Experimental Procedures), with each of them perfectly matching the 12 bp seed sequence at the 3′ end and the NGG PAM sequence of the sgRNA (there is no potential off-target site for Tet3 sgRNA using this prediction rule). From seven double-mutant mice produced from injection with high RNA concentration we PCR amplified 400 to 500 bp fragments from all seven potential off-target loci and found no cleavage in the Surveyor assay ( Figure S3), suggesting a high specificity of CRISPR/Cas system.
Figure S3.
Off-Target Analysis of Double-Mutant Mice, Related to Figure 2
(A) Three potential off targets of Tet1 sgRNA and four potential off targets of Tet2 sgRNA are shown. The 12 bp perfect matching seed sequence is labeled in blue, and NGG PAM sequence is labeled in red.
(B) Surveyor assay of all seven potential off-target loci in seven double-mutant mice derived with high concentration of Cas9 mRNA (100 ng/μl) injection. WT control is included as the eighth sample. The weak cleavage activity at Ubr1 locus is not due to off-target effect because sequences of these PCR products show no mutations.
Multiplexed Precise HDR-Mediated Genome Editing In Vivo
The NHEJ-mediated gene mutations described above produced mutant alleles with different and unpredictable insertions and deletions of variable size. We explored the possibility of precise homology directed repair (HDR)-mediated genome editing by coinjecting Cas9 mRNA, sgRNAs, and single-stranded DNA oligos into one-cell embryos. For this we designed an oligo targeting Tet1 so as to change two base pairs of a SacI restriction site and creating instead an EcoRI site and a second oligo targetingTet2 with two base pair changes that would convert an EcoRV site into an EcoRI site (Figure 3A). Blastocysts were derived from zygotes injected with Cas9 mRNA and sgRNAs and oligos targeting Tet1 or Tet2, respectively. DNA was isolated, amplified, and digested with EcoRI to detect oligo-mediated HDR events. Six out of nine Tet1-targeted embryos and 9 out of 15 Tet2-targeted embryos incorporated an EcoRI site at the respective target locus, with several embryos having both alleles modified (Figure S4A). When Cas9 mRNA, sgRNAs, and single-stranded DNA oligos targeting both Tet1 and Tet2 were coinjected into zygotes, out of 14 embryos, four were identified that were targeted with the oligo at the Tet1 locus, seven that were targeted with the oligo at the Tet2 locus and one embryo (2) that had one allele of each gene correctly modified (Figure S4B). All four alleles of embryo 2 were sequenced, confirming that one allele of each gene contained the 2 bp changes directed by the oligo, whereas the other alleles were disrupted by NHEJ-mediated deletion and insertion ( Figure S4C).
Figure 3.
Multiplexed HDR-Mediated Genome Editing In Vivo
(A) Schematic of the oligo-targeting sites at Tet1 and Tet2 loci. The sgRNA-targeting sequence is underlined, and the PAM sequence is labeled in green. Oligo targeting each gene is shown under the target site, with 2 bp changes labeled in red. Restriction enzyme sites used for RFLP analysis are bold and capitalized.
(B) RFLP analysis of double oligo injection mice with HDR-mediated targeting at the Tet1 and Tet2 loci.
(C) The sequences of both alleles of Tet1 and Tet2 in mouse 5 and 7 show simultaneously HDR-mediated targeting at one allele or two alleles of each gene, and NHEJ-mediated disruption at the other alleles. See also Figure S4.
Multiplexed Precise HDR-Mediated Genome Editing In Vivo, Related to Figure 3
(A) RFLP analysis of single oligo injection embryos with HDR-mediated targeting at Tet1 and Tet2 locus.
(B) RFLP analysis of double oligo injection embryos with multiplexed HDR-mediated targeting at both Tet1and Tet2 loci.
(C) Sequences of both alleles of Tet1 and Tet2 in embryo 2 show simultaneously HDR-mediated targeting at one allele of both genes, and NHEJ-mediated gene disruption at the other allele of each gene.
Blastocysts with double oligo injections were implanted into foster mothers and a total of 10 pups were born from 48 embryos transferred (21% live-birth rate). Upon RFLP analysis using EcoRI, we identified seven mice containing EcoRI sites at the Tet1 locus and eight mice containing EcoRI sites at the Tet2 locus, with six mice containing EcoRI sites at both Tet1 and Tet2 loci ( Figure 3B). We also applied RFLP analysis using SacI and EcoRV to Tet1 and Tet2 loci, respectively, showing that all alleles not targeted by oligos contained disruptions, which is in consistent with the high biallelic mutation rate by Cas9 mRNA and sgRNAs injection. These results were confirmed by sequencing demonstrating mutations in all four alleles of mouse 5 and 7 ( Figure 3C). Our results demonstrate that mice with HDR-mediated precise mutations in multiple genes can be generated in one step by CRISPR/Cas-mediated genome editing.
Discussion
The genetic manipulation of mice is a crucial approach for the study of development and disease. However, the generation of mice with specific mutations is labor intensive and involves gene targeting by homologous recombination in ES cells, the production of chimeric mice, and, after germline transmission of the targeted ES cells, the interbreeding of heterozygous mice to produce the homozygous experimental animals, a process that may take 6 to 12 months or longer (Capecchi, 2005). To produce mice carrying mutations in several genes requires time-consuming intercrossing of single-mutant mice. Similarly, the generation of ES cells carrying homozygous mutations in several genes is usually achieved by sequential targeting, a process that is labor intensive, necessitating multiple consecutive cloning steps to target the genes and to delete the selectable markers.
As summarized in Figure 4, we have established three different approaches for the generation of mice carrying multiple genetic alterations. We demonstrate that CRISPR/Cas-mediated genome editing in ES cells can generate the simultaneous mutations of several genes with high efficiency, a single-step approach allowing the production of cells with mutations in five different genes (Figure 4A). We chose the threeTet genes as targets because the respective mutant phenotypes have been well defined previously ( Dawlaty et al., 2011, 2013; Gu et al., 2011). Cells mutant for Tet1, 2 and 3were depleted of 5hmC as would be expected for loss of function mutations of the genes (Dawlaty et al., 2013). However, we have not as yet established, which of the Cas9-mediated gene mutations produced loss of function rather than hypomorphic alleles.
Figure 4.
Mutiplexed Genome Editing in ES Cells and Mouse
(A) Multiple gene targeting in ES cells.
(B) One-step generation of mice with multiple mutations. Upper: multiple targeted mutations with random indels introduced through NHEJ. Lower: multiple predefined mutations introduced through HDR-mediated repair.
We also show that mouse embryos can be directly modified by injection of Cas9 mRNA and sgRNA into the fertilized egg resulting in the efficient production of mice carrying biallelic mutations in a given gene. More significantly, coinjection of Cas9 with Tet1 andTet2 sgRNAs into zygotes produced mice that carried mutations in both genes (Figure 4B, upper). We found that up to 95% of newborn mice were biallelic mutant in the targeted gene when single sgRNA was injected and when coinjected with two different sgRNAs, up to 80% carried biallelic mutations in both targeted genes. Thus, mice carrying multiple mutations can be generated within 4 weeks, which is a much shorter time frame than can be achieved by conventional consecutive targeting of genes in ES cells and avoids time-consuming intercrossing of single-mutant mice.
The introduction of DSBs by CRISPR/Cas generates mutant alleles with varying deletions or insertions in contrast to designed precise mutations created by homologous recombination. The introduction of point mutations into human ES cells, cancer cell lines, and mouse by ZNF or TALEN along with DNA oligo has been demonstrated previously (Chen et al., 2011; Soldner et al., 2011; Wefers et al., 2013). We demonstrate that CRISPR/Cas-mediated targeting is useful to generate mutant alleles with predetermined alterations, and coinjection of single-stranded oligos can introduce designed point mutations into two target genes in one step, allowing for multiplexed gene editing in a strictly controlled manner (Figure 4B, lower). It will be of great interest to assess whether this targeting system allows for the production of conditional alleles, or precise insertion of larger DNA fragments such as GFP markers so as to generate conditional knockout and reporter mice for specific genes.
There are several potential limitations of the CRISPR/Cas technology. First, the requirement for a NGG PAM sequence of S. pyogenes Cas9 limits the target space in the mouse genome. It has been shown that the Streptococcus thermophilus LMD-9 Cas9 using different PAM sequence can also induce targeted DNA cleavage in mammalian cells ( Cong et al., 2013). Therefore, exploiting different Cas9 proteins may enable to target most of the mouse genome. Second, although the sgRNAs used here showed high targeting efficiency, much work is needed to elucidate the rules for designing sgRNAs with consistent high targeting efficiency, which is essential for multiplexed genome engineering. Third, although our off-target analysis for the seven most likely off targets of Tet1 and Tet2 sgRNAs failed to detect mutations in these loci, it is possible that other mutations were induced following as yet unidentified rules. A more thorough sequencing analysis for a large number of sgRNAs will provide more information about the potential off-target cleavage of the CRISPR/Cas system and lead to a better prediction of potential off-target sites. Last, oligo-mediated repair allows for precise genome editing, but the other allele is often mutated through NHEJ ( Figures 3B, 3C, andS4C). We have shown that using lower Cas9 mRNA concentration generates more mice with heterozygous mutations. Therefore, it may be possible to optimize the system for more efficient generation of mice with only one oligo -modified allele. In addition, employment of Cas9 nickase will likely avoid this complication because it mainly induces DNA single-strand break, which is typically repaired through HDR ( Cong et al., 2013;Mali et al., 2013).
It is likely that a much larger number of genomic loci than targeted in the present work can be modified simultaneously when pooled sgRNAs are introduced. The methods presented here open up the possibility of systematic genome engineering in mice, facilitating the investigation of entire signaling pathways, of synthetic lethal phenotypes or of genes that have redundant functions. A particularly interesting application is the possibility to produce mice carrying multiple alterations in candidate loci that have been identified in GWAS studies to play a role in the genesis of multigenic diseases. In summary, CRISPR/Cas-mediated genome editing makes possible the generation of ES cells and mice carrying multiple genetic alterations and will facilitate the genetic dissection of development and complex diseases.
One-Step Generation of Mice Carrying Reporter and Conditional Alleles by CRISPR/Cas-Mediated Genome Engineering
One-step generation of mice with reporters in endogenous genes
One-step generation of conditional mutant mice
Off-target analysis suggests high specificity of the CRISPR/Cas9 system
The type II bacterial CRISPR/Cas system is a novel genome-engineering technology with the ease of multiplexed gene targeting. Here, we created reporter and conditional mutant mice by coinjection of zygotes with Cas9 mRNA and different guide RNAs (sgRNAs) as well as DNA vectors of different sizes. Using this one-step procedure we generated mice carrying a tag or a fluorescent reporter construct in the Nanog, the Sox2, and the Oct4 gene as well as Mecp2 conditional mutant mice. In addition, using sgRNAs targeting two separate sites in the Mecp2 gene, we produced mice harboring the predicted deletions of about 700 bps. Finally, we analyzed potential off-targets of five sgRNAs in gene-modified mice and ESC lines and identified off-target mutations in only rare instances.
Mice with specific gene modification are valuable tools for studying development and disease. Traditional gene targeting in embryonic stem (ES) cells, although suitable for generating sophisticated genetic modifications in endogenous genes, is complex and time-consuming (Capecchi, 2005). The production of genetically modified mice and rats has been greatly accelerated by novel approaches using direct injection of DNA or mRNA of site-specific nucleases into the one-cell-stage embryo, generating DNA double-strand breaks (DSB) at specified sequences leading to targeted mutations (Carbery et al., 2010, Geurts et al., 2009, Shen et al., 2013, Sung et al., 2013, Tesson et al., 2011 and Wang et al., 2013). Coinjection of a single-stranded or double-stranded DNA template containing homology to the sequences flanking the DSB can produce mutant alleles with precise point mutations or DNA inserts (Brown et al., 2013, Cui et al., 2011, Meyer et al., 2010, Wang et al., 2013 and Wefers et al., 2013). Recently, pronuclear injection of two pairs of ZFNs and two double-stranded donor vectors into rat fertilized eggs produced rat containing loxP-flanked (floxed) alleles (Brown et al., 2013). However, the complex and time-consuming design and generation of ZFNs and double-stranded donor vectors limit the application of this method.
Previously, we used the type II bacterial CRISPR/Cas system as an efficient tool to generate mice carrying mutations in multiple genes in one step (Wang et al., 2013). However, this study left a number of issues unresolved. For example, neither the efficiency of using the CRISPR/Cas gene-editing approach for the insertion of DNA constructs into endogenous genes nor its utility to create conditional mutant mice was clarified. Here, we report the one-step generation of mice carrying reporter constructs in three different genes as well as the derivation of conditional mutant mice. In addition, we performed an extensive off-target cleavage analysis and show that off-target mutations are rare in targeted mice and ES cells derived from CRISPR/Cas zygote injection.
Finally, we designed sgRNA targeting the Oct4 3′ UTR, which was coinjected with a published donor vector designed to integrate the 3 kb transgene cassette (IRES-eGFP-loxP-Neo-loxP; Figure 2D) at the 3′ end of the Oct4 gene ( Lengner et al., 2007). Blastocysts were derived from injected zygotes, inspected for GFP expression, and explanted to derive ES cells. About 20% (47/254) of the blastocysts displayed uniform GFP expression in the ICM region. Three of nine derived ES cell lines expressed GFP (Figure 2E), including one showed mosaic expression ( Table S2). Three out of ten live-born mice contained the targeted allele ( Table 1). Correct targeting in mice and ES cell lines was confirmed by Southern blot analysis ( Figure 2F).
Conventionally, transgenic mice are generated by pronuclear instead of cytoplasmic injection of DNA. To optimize the generation of CRISPR/Cas9-targeted embryos, we compared different concentrations of RNA and the Nanog-mCherry or the Oct4-GFP DNA vectors as well as three different delivery modes: (1) simultaneous injection of all constructs into the cytoplasm, (2) simultaneous injection of the RNA and the DNA into the pronucleus, and (3) injection of Cas9/sgRNA into the cytoplasm followed 2 hr later by pronuclear injection of the DNA vector. Table S1 shows that simultaneous injection of all constructs into the cytoplasm at a concentration of 100 ng/μl Cas9 RNA, 50 ng/μl of sgRNA and 200 ng/μl of vector DNA was optimal, resulting in 9% (86/936) to 19% (47/254) of targeted blastocysts. Similarly, the simultaneous injection of 5 ng/μl Cas9 RNA, 2.5 ng/μl of sgRNA, and 10 ng/μl of DNA vector into the pronucleus yielded between 9% (7/75) and 18% (13/72) targeted blastocysts. In contrast, the two-step procedure with Cas9 and sgRNA simultaneous injected into the cytoplasm followed 2 hr later by pronuclear injection of different concentrations of DNA vector yielded no or at most 3% (1/34) positive blastocysts. Thus, our results suggest that simultaneous injection of RNA and DNA into the cytoplasm or nucleus is the most efficient procedure to achieve targeted insertion.
A total of 98 E13.5 embryos and mice were generated from zygotes injected with Cas9 mRNA, sgRNAs, and DNA oligos targeting the L2 and R1 sites. Genomic DNA was digested with both NheI and EcoRI, and analyzed by Southern blot using exon 3 and exon 4 probes (Figures 3A and 3B). The L2 and R1 oligos contained, in addition to the loxP site, different restriction sites (NheI or EcoRI). Thus, single loxP site integration at L2 or R1 will produce either a 3.9 kb or a 2 kb band, respectively, when hybridized with the exon3 probe (Figures 3A and B). We found that about 50% (45/98) of the embryos and mice carried a loxP site at the L2 site and about 25% (25/98) at the R1 site. Importantly, integration of both loxP sites on the same DNA molecule, generating a floxed allele, produces a 700 bp band as detected by exon 3 probe hybridization (Figures 3A and 3B). RFLP analysis, sequencing (Figures S4A and S4B) and Southern blot analysis (Figure 3B) showed that 16 out of the 98 mice tested contained two loxP sites flanking exon 3 on the same allele. Table 2 summarizes the frequency of all alleles and shows that the overall insertion frequency of an L2 or R1 insertion was slightly higher in females (21/38) than in males (28/60) consistent with the higher copy number of the X-linkedMecp2 gene in females. To confirm that the floxed allele was functional, we used genomic DNA for in vitro Cre-mediated recombination. Upon Cre treatment, both the deletion and circular products were detected by PCR in targeted mice, but not in DNA from wild-type mice ( Figure 3C). The PCR products were sequenced and confirmed the precise Cre-loxP-mediated recombination ( Figure S4C).
In this study, we demonstrate that CRISPR/Cas technology can be used for efficient one-step insertions of a short epitope or longer fluorescent tags into precise genomic locations, which will facilitate the generation of mice carrying reporters in endogenous genes. Mice and embryos carrying reporter constructs in the Sox2, the Nanog and the Oct4 gene were derived from zygotes injected with Cas9 mRNA, sgRNAs, and DNA oligos or vectors encoding a tag or a fluorescent marker. Moreover, microinjection of two Mecp2-specific sgRNAs, Cas9 mRNA, and two different oligos encoding loxP sites into fertilized eggs allowed for the one-step generation of conditional mutant mice. In addition, we show that the introduction of two spaced sgRNAs targeting the Mecp2 gene can produce mice carrying defined deletions of about 700 bp. Though all RNA and DNA constructs were injected into the cytoplasm or nucleus of zygotes, the gene modification events could happen at the one-cell stage or later. Indeed, Southern analyses revealed mosaicism in 17% (1/6) to 40% (20/49) of the targeted mice and ES cell lines indicating that the insertion of the transgenes had occurred after the zygote stage ( Table S2).
More…
In summary, CRISPR/Cas-mediated genome editing represents an efficient and simple method of generating sophisticated genetic modifications in mice such as conditional alleles and endogenous reporters in one step. The principles described in this study could be directly adapted to other mammalian species, opening the possibility of sophisticated genome engineering in many species where ES cells are not available.
Multiplexed activation of endogenous genes by CRISPR-on, an RNA-guided transcriptional activator system
Albert W Cheng1,2,*, Haoyi Wang1,*, Hui Yang1, Linyu Shi1, Yarden Katz1,3, Thorold W Theunissen1, Sudharshan Rangarajan1, Chikdu S Shivalila1,4, Daniel B Dadon1,4 and Rudolf Jaenisch1,4 Cell Research (2013) 23:1163–1171. http://dx.doi.org:/10.1038/cr.2013.122
Technologies allowing for specific regulation of endogenous genes are valuable for the study of gene functions and have great potential in therapeutics. We created the CRISPR-on system, a two-component transcriptional activator consisting of a nuclease-dead Cas9 (dCas9) protein fused with a transcriptional activation domain and single guide RNAs (sgRNAs) with complementary sequence to gene promoters. We demonstrate that CRISPR-on can efficiently activate exogenous reporter genes in both human and mouse cells in a tunable manner. In addition, we show that robust reporter gene activation in vivo can be achieved by injecting the system components into mouse zygotes. Furthermore, we show that CRISPR-on can activate the endogenous IL1RN, SOX2, and OCT4genes. The most efficient gene activation was achieved by clusters of 3-4 sgRNAs binding to the proximal promoters, suggesting their synergistic action in gene induction. Significantly, when sgRNAs targeting multiple genes were simultaneously introduced into cells, robust multiplexed endogenous gene activation was achieved. Genome-wide expression profiling demonstrated high specificity of the system.
Gene expression is strictly controlled in many biol-ogical processes, such as development and diseases. Transcription factors regulate gene expression by binding to specific DNA sequences at the enhancer and promoter regions of target genes, and modulate transcription through their effector domains1. Based on the same principle, artificial transcription factors (ATFs) have been generated by fusing various functional domains to a DNA binding domain engineered to bind to the genes of interest, thereby modulating their expression2,3. The capability of regulating endogenous gene expression using ATFs may facilitate the study of the transcriptional network underlying complex biological processes and provide new therapeutic options for diseases. Significant efforts and progress have been made to engineer DNA binding domains with defined specificities. The decipherment of the “code” of DNA binding specificity of zinc finger proteins and transcription activator-like effectors (TALE) has led to the rational design of DNA binding domains to recognize specific nucleotides with certain probability4,5,6,7,8,9,10. However, binding specificity of these ATFs is usually degenerate, can be difficult to predict and the complex and time-consuming design and generation limits their applications. To study the transcriptional network in a systematic manner, regulating multiple endogenous genes is required, prompting the development of efficient technology for simultaneous regulation of multiple endogenous genes.
CRISPR (clustered regularly interspaced short palin-dromic repeat) and Cas (CRISPR-associated) proteins are utilized by bacteria and archea to defend against viral pathogens11,12. Because the binding of Cas protein is guided by the simple base-pair complementarities between the engineered single guide RNA (sgRNA) and a target genomic DNA sequence, Cas9 could be directed to specific genomic locus or multiple loci simultaneously, by providing the engineered sgRNAs13,14,15,16,17,18,19,20. A recent study described the CRISPRi (CRISPR interference) system, in which the nuclease-deficient dCas9 (D10A; H840A) proteins blocked the transcription apparatus when directed to promoters or gene bodies in bacteria21. A subsequent study demonstrated a more efficient gene repression in eukaryotes by dCas9 fused with a transcription repression domain or exogenous transgene activation when fused with an activation domain22. Two most recent studies showed single endogenous gene activation using dCas9-based activators9,10. To what extent multiple endogenous genes could be regulated simultaneously has not been explored. In this study we report the generation of an RNA-programmable CRISPR-on system, which enables the simultaneous activation of multiple endogenous genes with a defined stoichiometry.
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We show here that the CRISPR-on system can be used for the simultaneous induction of at least three different endogenous genes. More significantly, we demonstrated that the stoichiometry of gene induction of multiple genes can be tuned by adjusting the relative amount of their cognate sgRNAs. Simultaneous activation of multiple endogenous genes with defined stoichiometry opens up novel opportunities for systems biology as it allows for the predictable manipulation of transcriptional networks.
Finally, with the ease of design and synthesis, a library of sgRNAs could be generated. When introduced into a cell line constitutively expressing dCas9 activator, gene activation screens mediated by RNA (RNAa) could be achieved. As the specificity components (sgRNA) can be separately designed and constructed from the effector component (Cas fusion proteins), the same library of sgRNAs could be used with different dCas9 fusions (e.g., VP160 domain for transactivation, KRAB domain for transcriptional repression, chromatin modifier domains for specific histone modification) to exert different functions at particular genomic loci.
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Maeder ML, Linder SJ, Reyon D, et al. Robust, synergistic regulation of human gene expression using TALE activators. Nat Methods 2013; 10:243–245. | Article | PubMed | CAS |
Perez-Pinera P, Ousterout DG, Brunger JM, et al. Synergistic and tunable human gene activation by combinations of synthetic transcription factors.Nat Methods 2013; 10:239–242. | Article | PubMed | CAS |
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Our long-range goals are to understand epigenetic regulation of gene expression in mammalian development and disease. An important question is to understand the different epigenetic conformations that distinguish differentiated cell states and to define strategies to transdifferentiate one differentiated cell type into another. Embryonic stem cells are of major significance because they have the potential to generate any cell type in the body and, therefore, are of great interest for regenerative medicine. A major focus of our work is to understand the molecular mechanisms that allow the reprogramming of somatic cells to an embryonic pluripotent state and to use the potential of patient specific pluripotent cells to study complex human diseases.
Three years ago, no one knew or cared about much about a protein called Cpf1 produced by a bacterial gene. Now, it shows potential for making a fast-developing genome editing technique called CRISPR easier and more accurate. Bioinformaticians identified this protein and its potential connection to CRISPR by scanning the public database of genome sequences. Their colleagues now show that two of 16 versions of this protein tested can delete a gene in a human cell. Cpf1 has other advantages as well-being smaller than one of the popular Cas9 proteins used and depending on a smaller piece of RNA to find its target DNA. But its utility for editing genomes of human and other cells needs further testing.
CRISPR/Cas9 Finds Its Way As an Important Tool For Drug Discovery & Development
UPDATED 6/11/2021
CRISPR Diagnostics: CRISPR-dx Comes of Age: Tool in Drug Development
The past five years has seen a rapid expansion of the ability of CRISPR based tools toward diagnostic testing. Recently, CRISPR has been used to detect SARS-CoV-2 in patients. An article in the journal Science describes the different classes of CRISPR diagnostics in use today .
Update near end of post
UPDATED 8/08/2020
Association to Causation: Using GWAS to Identify Druggable Targets
A Gen Webinar Thursday, August 6, 2020; 11:00am – 12:30pm EST
See at end of post
Curator: Stephen J. Williams, Ph.D.
Article 21.4.8- CRISPRCas9 Finds Its Way As an Important Tool For Drug Discovery & Development
CRISPR/Cas9 Finds Its Way As an Important Tool For Drug Discovery & Development, 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
The RNA-guided Cas9 nuclease from the microbial clustered regularly interspaced short palindromic repeats (CRISPR)adaptive immune system can be used to facilitate efficient genome engineering in eukaryotic cells by simply specifying a 20-nt targeting sequence within its guide RNA.
CRISPR/Cas systems are part of the adaptive immune system of bacteria and archaea, protecting them against invading nucleic acids such as viruses by cleaving the foreign DNA in a sequence-dependent manner. Although CRISPR arrays were first identified in the Escherichia coli genome in 1987 (Ishino et al., 1987), their biological function was not understood until 2005, when it was shown that the spacers were homologous to viral and plasmid sequences suggesting a role in adaptive immunity (Bolotin et al., 2005; Mojica et al., 2005; Pourcel et al., 2005). Two years later, CRISPR arrays were confirmed to provide protection against invading viruses when combined with Cas genes (Barrangou et al., 2007). The mechanism of this immune system based on RNA-mediated DNA targeting was demonstrated shortly thereafter (Brounset al., 2008; Deltcheva et al., 2011; Garneau et al., 2010; Marraffini and Sontheimer, 2008).
Jennifer Doudna, PhD Professor of Molecular and Cell Biology and Chemistry, University of California, Berkeley Investigator, Howard Hughes Medical Institute has recently received numerous awards and accolades for the discovery of CRISPR/Cas9 as a tool for mammalian genetic manipulation as well as her primary intended research target to understand bacterial resistance to viral infection.
A good post on the matter and Dr. Doudna can be seen below:
In Delineating a Role for CRISPR-Cas9 in Pharmaceutical Targeting inheritable metabolic disorders in which may benefit from a CRISPR-Cas9 mediated therapy is discussed. However this curation is meant to focus on CRISPR/CAS9 AS A TOOL IN PRECLINICAL DRUG DEVELOPMENT.
Three Areas of Importance of CRISPR/Cas9 as a TOOL in Preclinical Drug Discovery Include:
Gene-Function Studies: CRISPR/CAS9 ability to DEFINE GENETIC LESION and INSERTION SITE
CRISPR/CAS9 Use in Developing Models of Disease
CRISPR/CAS9 Use as aDiagnostic Tool
Using CRISPR/Cas9 in PRECLINICAL TOXICOLOGY STUDIES
I. Gene-Function Studies: CRISPR/CAS9 ability to DEFINE GENETIC LESION and INSERTION SITE
The advent of the first tools for manipulating genetic material (cloning, PCR, transgenic technology, and before microarray and other’omic methods) allowed scientists to probe novel, individual gene functions as well as their variants and mutants in a “one-gene-at-a time” process. In essence, a gene (or mutant gene) was sequenced, cloned into expression vectors and transfected into recipient cells where function was evaluated.
However, some of the experimental issues with this methodology involved
Lack of knowledge of insertion site of the transgene – this leads to off-target effects usually due to insertion of a transgene in front of unwanted promoters or insertion at a site resulting in gene disruption or even mutagenesis. In an extreme case, such as transposon-induced mutagenesis may lead to transformation as described in an earlier post on this site How Mobile Elements in “Junk” DNA Promote Cancer – Part 1: Transposon-mediated Tumorigenesis
Most transfections experiments result in NON ISOGENIC cell lines – by definition the insertion of a transgene alters the genetic makeup of a cell line. Simple transfection experiments with one transgene compared to a “null” transfectant compares non-isogenic lines, possibly confusing the interpretation of gene-function studies. Therefore a common technique is to develop cell lines with inducible gene expression, thereby allowing the investigator to compare a gene’s effect in ISOGENIC cell lines.
Use of CRSPR in Highthrough-put Screening of Genetic Function
A very nice presentation and summary of CRSPR’s use in determining gene function in a high-throughput manner can be found below
an approach for global detection of DNA double-stranded breaks (DSBs) introduced by RGNs and potentially other nucleases. This method, called genome-wide, unbiased identification of DSBs enabled by sequencing (GUIDE-seq), relies on capture of double-stranded oligodeoxynucleotides into DSBs. Application of GUIDE-seq to 13 RGNs in two human cell lines revealed wide variability in RGN off-target activities and unappreciated characteristics of off-target sequences. The majority of identified sites were not detected by existing computational methods or chromatin immunoprecipitation sequencing (ChIP-seq). GUIDE-seq also identified RGN-independent genomic breakpoint ‘hotspots’.
1) In this talk Dr. Tyler Jacks discussed his use of CRSPR to generate a mouse model of liver tumor in an immunocompetent mouse. Some notes from this talk are given below
B) Engineering Cancer Genomes: Tyler Jacks, Ph.D.; Director, Koch Institute for Integrative Cancer Research
Cancer GEM’s (genetically engineered mouse models of cancer) had moved from transgenics to defined oncogenes
Observation that p53 -/- mice develop spontaneous tumors (lymphomas)
then GEMs moved to Cre/Lox systems to generate mice with deletions however these tumor models require lots of animals, much time to create, expensive to keep;
figured can use CRSPR/Cas9 as rapid, inexpensive way to generate engineered mice and tumor models
he used CRSPR/Cas9 vectors targeting PTEN to introduce PTEN mutations in-vivo to hepatocytes; when they also introduced p53 mutations produced hemangiosarcomas; took ONLY THREE months to produce detectable tumors
also produced liver tumors by using CRSPR/Cas9 to introduce gain of function mutation in β-catenin
2) In the Upcoming Meeting New Frontiers in Gene Editing multiple uses of CRISPR technology is discussed in relation to gene knockout/function studies, tumor model development and
New Frontiers in Gene Editing
Session Spotlight: BUILDING IN VIVO MODELS FOR DRUG DISCOVERY
Genome Editing Animal Models in Drug Discovery Myung Shin, Ph.D., Senior Principal Scientist, Biology-Discovery, Genetics and Pharmacogenomics, Merck Research Laboratories
Recent advances in genome editing have greatly accelerated and expanded the ability to generate animal models. These tools allow generating mouse models in condensed timeline compared to that of conventional gene-targeting knock-out/knock-in strategies. Moreover, the genome editing methods have expanded the ability to generate animal models beyond mice. In this talk, we will discuss the application of ZFN and CRISPR to generate various animal models for drug discovery programs.
In vivo Cancer Modeling and Genetic Screening Using CRISPR/Cas9 Sidi Chen, Ph.D., Postdoctoral Fellow, Laboratories of Dr. Phillip A. Sharp and Dr. Feng Zhang, Koch Institute for Integrative Cancer Research at MIT and Broad Institute of Harvard and MIT
Here we describe a genome-wide CRISPR-Cas9-mediated loss-of-function screen in tumor growth and metastasis. We mutagenized a non-metastatic mouse cancer cell line using a genome-scale library. The mutant cell pool rapidly generates metastases when transplanted into immunocompromised mice. Enriched sgRNAs in lung metastases and late stage primary tumors were found to target a small set of genes, suggesting specific loss-of-function mutations drive tumor growth and metastasis.
FEATURED PRESENTATION: In vivo Chromosome Engineering Using CRISPR-Cas9 Andrea Ventura, M.D., Ph.D., Assistant Member, Cancer Biology and Genetics Program, Memorial Sloan Kettering Cancer Center
We will discuss our experience using somatic genome editing to engineer oncogenic chromosomal rearrangements in vivo. More specifically, we will present the results of our ongoing efforts aimed at modeling cancers driven by chromosomal rearrangements using viral mediated delivery of Crispr-Cas9 to adult animals.
RNAi and CRISPR/Cas9-Based in vivo Models for Drug Discovery Christof Fellmann, Ph.D., Postdoctoral Fellow, Laboratory of Dr. Jennifer Doudna, Department of Molecular and Cell Biology, The University of California, Berkeley
Genetically engineered mouse models (GEMMs) are a powerful tool to study disease initiation, treatment response and relapse. By combining CRISPR/Cas9 and “Sensor” validated, tetracycline-regulated “miR-E” shRNA technology, we have developed a fast and scalable platform to generate RNAi GEMMs with reversible gene silencing capability. The synergy of CRISPR/Cas9 and RNAi enabled us to not only model disease pathogenesis, but also mimic drug therapy in mice, providing us capability to perform preclinical studies in vivo.
In vivo Genome Editing Using Staphylococcus aureus Cas9 Fei Ann Ran, Ph.D., Post-doctoral Fellow, Laboratory of Dr. Feng Zhang, Broad Institute and Junior Fellow, Harvard Society of Fellows
The RNA-guided Cas9 nuclease from the bacterial CRISPR/Cas system has been adapted as a powerful tool for facilitating targeted genome editing in eukaryotes. Recently, we have identified an additional small Cas9 nuclease from Staphylococcus aureus that can be packaged with its guide RNA into a single adeno-associated virus (AAV) vector for in vivo applications. We demonstrate the use of this system for effective gene modification in adult animals and further expand the Cas9 toolbox for in vivo genome editing.
OriGene, Making the Right Tools for CRISPR Research Xuan Liu, Ph.D., Senior Director, Marketing, OriGene
CRISPR technology has quickly revolutionized the scientific community. Its simplicity has democratized the genome editing technology and enabled every lab to consider its utility in gene function research. As the largest tool box for gene functional research, OriGene created a large collection of CRISPR-related tools, including various all-in-one vectors for gRNA cloning, donor vector backbones, genome-wide knockout kits, AAVS1 insertion vectors, etc. OriGene’s high quality products will accelerate CRISPR research.
A critical component of producing transgenic animals is the ability of each successive generations to pass on the transgene. In her post on this site, A NEW ERA OF GENETIC MANIPULATION Dr. Demet Sag discusses the molecular biology of Cas9 systems and their efficiency to cause point mutations which can be passed on to subsequent generations
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This group developed a new technology for editing genes that can be transferable change to the next generation by combining microbial immune defense mechanism, CRISPR/Cas9 that is the latest ground breaking technology for translational genomics with gene therapy-like approach.
In short, this so-called “mutagenic chain reaction” (MCR) introduces a recessive mutation defined by CRISPR/Cas9 that lead into a high rate of transferable information to the next generation. They reported that when they crossed the female MCR offspring to wild type flies, the yellow phenotype observed more than 95 percent efficiency.
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The advantage of CRISPR/Cas9 over ZFNs or TALENs is its scalability and multiplexibility in that multiple sites within the mammalian genome can be simultaneously modified, providing a robust, high-throughput approach for gene editing in mammalian cells.
Applied StemCell, Inc. offers various services related to animal models including conventional transgenic rats, and phenotype analysis using knock-in, knock-out strategies.
Further explanation of their use of CRSPR can be found at the site below:
III. Using CRISPR/Cas9 in PRECLINICAL TOXICOLOGY STUDIES
As of now it is unclear as to the strategy of pharma in how to use this technology for toxicology testing however a few companies have licensed the technology to use across their R&D platforms including
A recent paper used a sister technique TALEN to generate knock-in pigs which suggest that it would be possible to generate pigs with human transgenes, especially in human liver isozymes in orer to study hepatotoxicity of drugs.
Associate Professor UCSF Investigator Chan Zuckerberg Biohub
Kevin Holden, PhD
Head of Science Synthego
Abhi Saharia, PhD
VP, Commercial Development Synthego
Human genetics provides perhaps the single best opportunity to innovate and improve clinical success rates, through the identification of novel drug targets for complex disease. Even as correlation identifies multiple genetic variants associated with disease, it is challenging to conduct requisite functional studies to identify the causal variants, especially since most association signals map to non-coding regions of the genome.
Genetic editing technologies, such as CRISPR, have enabled the modeling of associated variants at their native loci, including non-coding loci, empowering the identification of underlying biological mechanisms of disease with potential causal genes. However, genome editing is largely manual today severely limiting scale, and forcing the use of rational filters to prioritize which variants to investigate functionally.
In this GEN webinar, we will discuss several strategies enabling large-scale functional investigation of disease-associated variants in a cost- and time-effective manner, including different types of pooled CRISPR-based screens and the development of a fully automated genome engineering platform. We will also review how optimization of genome engineering on this platform enables the engineering of disease-associated variants at scale in pluripotent cells.
They will be presenting on use of wide scale CRSPR screens to validate druggable targets
The presenters will also discuss new platforms for these wide scale screens
Big gap between accumulation of genetic variant information and functions of these variants
CRSPRi or CRSPa (siRNA coupled or enhancer coupled CRSPR guides)
Arrayed screens: multiplate guide RNAs and phenotype measured (phenotype can be morphology, complex biological systems like organoids or non autonomous functions
Using pooled screens and use of suitable cell model critical for this strategy
For example in iPSC vs. neurons has different expression patterns upon same CRSPR of UBA1
Advantage is using CRSPR to take iPSC from diseased variant patient to make a corrected isogenic control then introduce gRNAs and use modifier screens to determine phenotypes
Generated a platform called CRISPRbrain.org to do bioinformatics on various experiments with different guide RNAs (CRSPRs)
Nature medicine paper did GWAS and found 27 SNV associated with high risk disease and a rational filter focused on 1 SNV in noncoding region but why study a single variant and if studied all 27 would they have been able to identify a more representative druggable set?
Goal is to reduce or eliminate these rational filters
Syntheco uses an electroporation with ribonucleic proteins (RNP) to give highest efficiency and minimizes off target as complex is only in cells for a short period of time
They confirm they are doing single cell cloning by using automated microscopy to confirm single cell growth in each cloning well
Engineering iPSc genetically modified cells at scale
The closer you get to your target site the more efficient your CRSPR so a big factor when making guides, especially for knock-in CRSPR
Adding a small molecule non homologous end joining inhibitor increases efficiency to 95%
Cold shocking the cells also assists in homologous repair
Use cleavage resistant templates
III. CRISPR/CAS9 AS A DIAGNOSTIC TOOL
In the journal Science, Omar Abudayyeh and Jonathan Gootenberg discuss how CRISPR-based diagnostic (CRISPR-dx) tools offer a solution, and multiple CRISPR-dx products for detection of the SARS-CoV-2 RNA genome have been authorized by the US Food and Drug Administration (FDA). In addition they discuss the work by Jiao et al. in combining this technique to develop a rapid and sensitive SARS-CoV2 diagnostic test.
Omar O. Abudayyeh, Jonathan S. Gootenberg. Science 28 May 2021: CRISPR Diagnostics Vol. 372, Issue 6545, pp. 914-915; DOI: 10.1126/science.abi9335
Summary
Although clinical diagnostics take many forms, nucleic acid–based testing has become the gold standard for sensitive detection of many diseases, including pathogenic infections. Quantitative polymerase chain reaction (qPCR) has been widely adopted for its ability to detect only a few DNA or RNA molecules that can unambiguously specify a particular disease. However, the complexity of this technique restricts application to laboratory settings. The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) pandemic has underscored the need for the development and deployment of nucleic acid tests that are economical, easily scaled, and capable of being run in low-resource settings, without sacrifices in speed, sensitivity or specificity. CRISPR-based diagnostic (CRISPR-dx) tools offer a solution, and multiple CRISPR-dx products for detection of the SARS-CoV-2 RNA genome have been authorized by the US Food and Drug Administration (FDA). On page 941 of this issue, Jiao et al. (1) describe a new CRISPR-based tool to distinguish several SARS-CoV-2 variants in a single reaction.
Although clinical diagnostics take many forms, nucleic acid–based testing has become the gold standard for sensitive detection of many diseases, including pathogenic infections. Quantitative polymerase chain reaction (qPCR) has been widely adopted for its ability to detect only a few DNA or RNA molecules that can unambiguously specify a particular disease. However, the complexity of this technique restricts application to laboratory settings. The severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) pandemic has underscored the need for the development and deployment of nucleic acid tests that are economical, easily scaled, and capable of being run in low-resource settings, without sacrifices in speed, sensitivity or specificity. CRISPR-based diagnostic (CRISPR-dx) tools offer a solution, and multiple CRISPR-dx products for detection of the SARS-CoV-2 RNA genome have been authorized by the US Food and Drug Administration (FDA). On page 941 of this issue, Jiao et al. (1) describe a new CRISPR-based tool to distinguish several SARS-CoV-2 variants in a single reaction.
There are multiple types of CRISPR systems comprising basic components of a single protein or protein complex, which cuts a specific DNA or RNA target programmed by a complementary guide sequence in a CRISPR-associated RNA (crRNA). The type V and VI systems and the CRISPR-associated endonucleases Cas12 (2, 3) and Cas13 (4, 5) bind and cut DNA or RNA, respectively. Furthermore, upon recognizing a target DNA or RNA sequence, Cas12 and Cas13 proteins exhibit “collateral activity” whereby any DNA or RNA, respectively, in the sample is cleaved regardless of its nucleic acid sequence (4, 6). Thus, reporter DNAs or RNAs, which allow for visual or fluorescent detection upon cleavage, can be added to a sample to infer the presence or absence of specific DNA or RNA species (4–8).
Initial versions of CRISPR-dx utilizing Cas13 alone were sensitive to the low picomolar range, corresponding to a limit of detection of millions of molecules in a microliter sample. To improve sensitivity, preamplification methods, such as recombinase polymerase amplification (RPA), PCR, loop-mediated isothermal amplification (LAMP), or nucleic acid sequence–based amplification (NASBA), can be used with Cas12 or Cas13 to enable a limit of detection down to a single molecule (8). This preamplification approach, applicable to both Cas12 and Cas13 (6, 7), enabled a suite of detection methods and multiplexing up to four orthogonal targets (7). Additional developments expanded CRISPR-dx readouts beyond fluorescence, including lateral flow (7), colorimetric (9), and electronic or material responsive readouts (10), allowing for instrument-free approaches. In addition, post–collateral-cleavage amplification methods, such as the use of the CRISPR-associated enzyme Csm6, have been combined with Cas13 to further increase the speed of CRISPR-dx tests (7). As an alternative to collateral-cleavage–based detection, type III CRISPR systems, which involve large multiprotein complexes capable of targeting both DNA and RNA, have been used for SARS-CoV-2 detection through production of colorimetric or fluorometric readouts (11).
FDA-authorized CRISPR-dx tests are currently only for use in centralized labs, because the most common CRISPR detection protocols require fluid handling steps and two different incubations, precluding their immediate use at the point of care. Single-step formulations have been developed to overcome this limitation, and these “one-pot” versions of CRISPR-dx are simple to run, operate at a single temperature, and run without complex equipment, producing either fluorescence or lateral flow readouts. The programmability of CRISPR makes new diagnostic tests easier to develop, and within months of the release of the SARS-CoV-2 genome, many COVID-19–specific CRISPR tests were reported and distributed around the world.
The broader capability for Cas enzyme–enhanced nucleic acid binding or cleavage has led to several other detection modalities. Cas9-based methods for cleaving nucleic acids in solution for diagnostic purposes have been combined with other detection platforms, such as destruction of undesired amplicons for preparation of next-generation sequencing libraries (12), or selective removal of alleles for nucleotide-specific detection (13). Alternatively, the programmable cleavage event from the Cas nuclease can be used to initiate an amplification reaction (14). Cas9-based DNA targeting has also been used for nucleotide detection in combination with solid-state electronics, promising an amplification-free platform for detection. In this platform, called CRISPR-Chip, the Cas9 protein binds nucleotide targets of interest (often in the context of the native genome) to graphene transistors, where the presence of these targets alters either current or voltage (15). By utilizing additional Cas9 orthologs and specific guide designs, CRISPR-Chip approaches have been tuned for single–base-pair sensitivity (15). Because they are integrated with electronic readers, CRISPR-Chip platforms may allow facile point-of-care detection with handheld devices.
Different classes of CRISPR diagnostics. GRAPHIC: ERIN DANIEL
Jiao et al. use a distinct characteristic of type II CRISPR systems, which involve Cas9, to develop a new type of noncollateral based CRISPR detection. Unlike Cas12s and Cas13, Cas9-crRNA complex formation requires an additional RNA known as the trans-activating CRISPR RNA (tracrRNA). By sequencing RNAs bound to Cas9 from Campylobacter jejuni in its natural host, the authors identified unexpected crRNAs, called noncanonical crRNA (ncrRNA), that corresponded to endogenous transcripts. Upon investigation of this surprising observation, it became clear that the tracrRNA was capable of hybridizing to semi-complementary sequences from a variety of RNA sources, leading to biogenesis of ncrRNAs of various sizes. Recognizing that they could program tracrRNAs to target a transcript of interest, the authors generated a reprogrammed tracrRNA (Rptr) that could bind and cleave a desired transcript, converting a piece of that transcript into a functional guide RNA. By then creating fluorescent DNA sensors that would be cleaved by the Rptr and ncrRNAs, the sensing of RNA by Cas9 could be linked to a detectable readout. This platform, called LEOPARD (leveraging engineered tracrRNAs and on-target DNAs for parallel RNA detection), can be combined with gel-based readouts and enables multiplexed detection of several different sequences in a single reaction (see the figure).
Jiao et al. also combined LEOPARD with PCR in a multistep workflow to detect SARS-CoV-2 genomes from patients with COVID-19. Although more work is needed to integrate this Cas9-based detection modality into a single step with RPA or LAMP to create a portable and sensitive isothermal test, an advantage of this approach is the higher-order multiplexing that can be achieved, allowing multiple pathogens, diseases, or variants to be detected simultaneously. More work is also needed to combine this technology with extraction-free methods for better ease of use; alternative readouts to gel-based readouts, such as lateral flow and colorimetric readouts, would be beneficial for point-of-care detection.
In just 5 years, the CRISPR-dx field has rapidly expanded, growing from a set of peculiar molecular biology discoveries to multiple FDA-authorized COVID-19 tests and spanning four of the six major subtypes of CRISPR systems. Despite the tremendous promise of CRISPR-dx, substantial challenges remain to adapting these technologies for point-of-care and at-home settings. Simplification of the chemistries to operate as a single reaction in a matter of minutes would be revolutionary, especially if the reaction could be run at room temperature without any complex or expensive equipment. These improvements to CRISPR-dx assays can be achieved by identification or engineering of additional Cas enzymes with lower-temperature requirements, higher sensitivity, or faster kinetics, enabling rapid and simple amplification-free detection with single-molecule sensitivity.
Often overlooked is the necessity for a sample DNA or RNA preparation step that is simple enough to be added directly to the CRISPR reaction to maintain a simple workflow for point-of-care testing. In addition, higher-order multiplexing developments would allow for expansive testing menus and approach the possibility of testing for all known diseases. As these advancements are realized, innovative uses of CRISPR-dx will continue in areas such as surveillance, integration with biomaterials, and environmental monitoring. In future years, CRISPR-dx assays may become universal in the clinic and at home, reshaping how diseases are diagnosed.
Where is the most promising avenue to success in Pharmaceuticals with CRISPR-Cas9?
Author: Larry H. Bernstein, MD, FCAP
Article 21.4.5- Where is the most promising avenue to success in Pharmaceuticals with CRISPR-Cas9
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 (RNAi) 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
CRISPR/Cas (clustered regularly interspaced palindromic repeats/CRISPR-associated) systems are a bacterial defence against invading foreign nucleic acids derived from bacteriophages or exogenous plasmids1, 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 degradation5, 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 genes9, 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 pathogens11, 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.
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.
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.”
Delineating a Role for CRISPR-Cas9 in Pharmaceutical Targeting, 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
Abstract
The recent development of advanced methods for genome engineering has superceded methods already in used in recent years of the 21st century. The CRISPR-Cas9 application for genome editing has real potential for pharmaceutical development, and perhaps also for diagnostics. The importance of conjoint development of diagnostics and therapeutics can’t be overstressed. Further, the limitations of the method have to be viewed in the light of the historical development of inborn errors of human metabolism, and current understanding of complex polygenomic and environmental risk factors.
Genome editing technologies enable the deletion, insertion or correction of DNA at specific targeted sites within an organism’s genome. The power of the technology lies in its ability to specifically target any site in the genome and to alter the DNA sequence at that site. This has opened the door to potentially curing diseases caused by genetic defects, whether inherited or acquired.
Genome editing can be applied across many diverse fields of science. It has allowed researchers to gain a much deeper understanding of the role played by individual genes. Researchers working in the biomedical field use these techniques to address diseases that are known to have a genetic origin.
Early genome-editing research focused on the use of zinc finger nucleases and transcription activator-like effector nucleases (TALENs), which laid important foundations in establishing genome engineering as a potential approach for treating human diseases.
The recent discovery of CRISPR-Cas9, followed by work demonstrating its advantages over traditional approaches, promises a step-change in the use of genome editing to develop transformative medicines for serious human diseases.
Cas9* is an endonuclease (an enzyme) that can be easily programmed with RNA to cut DNA at targeted sites within the genome, enabling the deletion, insertion or correction of target genes, including those that cause diseases, with surgical precision. By using CRISPR-Cas9* genome-editing technology, scientists and clinicians are conducting pioneering research with a view to tackling both recessive and dominant genetic defects.
In order to find a place for CRISPR-Cas9 in gene therapy, it becomes necessary to consider inborn errors of metabolism and the evolution of traditional approaches to genetic diseases. Traditional gene therapy approaches to date have only been useful in correcting some recessive genetic disorders. Thanks to its ease of use and broad applicability, CRISPR-Cas9 has truly democratized genome editing and transformed many areas of research. Thousands of academic laboratories across the world are carrying out research using the technology. To this point, the technology known as CRISPR-Cas9 has been a science project, a research tool with enormous potential.
Genetic Disorders
A genetic disorder is a genetic problem caused by one or more abnormalities in the genome, especially a condition that is present from birth (congenital). Most genetic disorders are quite rare and affect one person in every several thousands or millions.
Genetic disorders may or may not be heritable, i.e., passed down from the parents’ genes. In non-heritable genetic disorders, defects may be caused by new mutations or changes to the DNA. In such cases, the defect will only be heritable if it occurs in the germ line. The same disease, such as some forms of cancer, may be caused by an inherited genetic condition in some people, by new mutations in other people, and mainly by environmental causes in still other people. Whether, when and to what extent a person with the genetic defect or abnormality will actually suffer from the disease is almost always affected by the environmental factors and events in the person’s development.
A single-gene disorder is the result of a single mutated gene. Over 4000 human diseases are caused by single-gene defects.[4] Single-gene disorders can be passed on to subsequent generations in several ways. Genomic imprinting and uniparental disomy, however, may affect inheritance patterns. The divisions betweenrecessive and dominant types are not “hard and fast”, although the divisions between autosomal and X-linked types are (since the latter types are distinguished purely based on the chromosomal location of the gene). For example, achondroplasia is typically considered a dominant disorder, but children with two genes for achondroplasia have a severe skeletal disorder of which achondroplasics could be viewed as carriers. Sickle-cell anemia is also considered a recessive condition, but heterozygous carriers have increased resistance to malaria in early childhood, which could be described as a related dominant condition.[5] When a couple where one partner or both are sufferers or carriers of a single-gene disorder wish to have a child, they can do so through in vitro fertilization, which means they can then have a preimplantation genetic diagnosis to check whether the embryo has the genetic disorder.[6]
Genes causing rare heritable childhood diseases are being discovered at an accelerating pace driven by the decreasing cost and increasing accessibility of next-generation DNA sequencing combined with the maturation of strategies for successful gene identification. The findings are shedding light on the biological mechanisms of childhood disease and broadening the phenotypic spectrum of many clinical syndromes. Still, thousands of childhood disease genes remain to be identified, and given their increasing rarity, this will require large-scale collaboration that includes mechanisms for sharing phenotypic and genotypic data sets. Nonetheless, genomic technologies are poised for widespread translation to clinical practice for the benefit of children and families living with these rare diseases.
Single gene defects result in abnormalities in the synthesis or catabolism of proteins, carbohydrates, fats, or complex molecules. Most are due to a defect in an enzyme or transport protein, which results in a block in a metabolic pathway. Effects are due to toxic accumulations of substrates before the block, intermediates from alternative metabolic pathways, defects in energy production and use caused by a deficiency of products beyond the block, or a combination of these metabolic deviations. Nearly every metabolic disease has several forms that vary in age of onset, clinical severity, and, often, mode of inheritance.
There is a large number of inborn errors of metabolism.
Victor Almon McKusick (October 21, 1921 – July 22, 2008), an internist and medical geneticist, was the University Professor of Medical Genetics and Professor of Medicine at the Johns Hopkins Hospital, Baltimore, MD, USA.[1] He was a proponent of the mapping of the human genome due to its use for studying congenital diseases. He is well known for his studies of the Amish and, what he called, “little people”. He was the original author and, until his death, remained chief editor of Mendelian Inheritance in Man (MIM) and its online counterpart Online Mendelian Inheritance in Man (OMIM). He is widely known as the “father of medical genetics”.[2]
McKusick traveled to Copenhagen to speak about the heritable disorders of connective tissue at the first international congress of human genetics. The meeting looms as the birthplace of the medical genetics field.[2] In the following decades, McKusick created and chaired a new Division of Medical Genetics at Hopkins beginning in 1957. In 1973, he served as Physician-in-Chief, William Osler Professor of Medicine, and Chairman of the Department of Medicine at Johns Hopkins Hospital and School of Medicine.[6] He held concurrent appointments as University Professor of Medical Genetics at the McKusick–Nathans Institute of Genetic Medicine, Professor of Medicine at the Johns Hopkins School of Medicine, Professor of Epidemiology at the Johns Hopkins Bloomberg School of Public Health, and Professor of Biology at Johns Hopkins University.[5] He co-founded Genomics in 1987 with Dr. Frank Ruddle, and served as an editor.[6] He was a lead investigator in determining if Abraham Lincoln had Marfan syndrome.[8]
Elizabeth F. Neufeld
Born in France, Elizabeth Neufeld immigrated to the United States in 1940. She obtained a BS from Queens College, New York and a Ph.D. from the University of California Berkeley. After postdoctoral training in, she moved to the NIH in Bethesda, MD, where she began her studies of a rare group of genetic diseases. She moved back to California in 1984 as Chair of the Department of Biological Chemistry – a position that she occupied till 2004.
The brain in a mouse model of a genetic lysosomal disorder, Sanfilippo syndrome type B
Our interests have long been the cause, consequences and treatment of human genetic diseases due to deficiency of lysosomal enzymes. The disease currently under investigation is the Sanfilippo syndrome type B (MPS III B). It is caused by mutation in the NAGLU gene, with resulting deficiency of the lysosomal enzyme alpha-N-acetyl-glucosaminidase and accumulation of its substrate (heparan sulfate). The disease manifests itself in childhood by severe mental retardation and intractable behavioral problems. The neurologic deterioration progresses to dementia, with death usually in the second decade. We use a mouse knockout model (Naglu -/-) in order to study the pathophysiology of the disease and to develop therapy. Because of the special cell biology of lysosomal enzymes, which can be taken up by receptor-mediated endocytosis, exogenous administration of the enzyme could theoretically cure the disease. Unfortunately, the blood-brain barrier (BBB) prevents the therapeutic enzyme from reaching the brain. Part of our current research is to develop a novel technology to get lysosomal enzymes across the BBB. We also study changes in gene and protein expression in some specific parts of the brain, in which there is accumulation of certain lipids and proteins which seem unrelated biochemically to each other or to the primary defect. We try to understand the cause and consequences of these accumulations. Although they are secondary defects, they may be relevant to the pathophysiology of the dieease and may have represent targets for pharmacologic intervention.
Neufeld began her scientific studies at a time when few women chose science as a career. The historical bias against women in science, compounded with an influx of men coming back from the Second World War and going to college, made positions for women rare; few women could be found in the science faculties of colleges and universities.
When she first began working on Hurler syndrome in 1967, she initially thought the problem might stem from faulty regulation of the sugars, but experiments showed the problem was in fact the abnormally slow rate at which the sugars were broken down. Working with fellow scientist Joseph Fratantoni, Neufeld attempted to isolate the problem by tagging mucopolysaccharides with radioactive sulfate, as well as mixing normal cells with MPS patient cells. Fratantoni inadvertently mixed cells from a Hurler patient and a Hunter patient—and the result was a nearly normal cell culture. The two cultures had essentially “cured” each other.
In 1973 Neufeld was named chief of NIH’s Section of Human Biochemical Genetics, and in 1979 she was named chief of the Genetics and Biochemistry Branch of the National Institute of Arthritis, Diabetes, and Digestive and Kidney Diseases (NIADDK). She served as deputy director in NIADDK’s Division of Intramural Research from 1981 to 1983. In 1984 Neufeld went back to the University of California, this time the Los Angeles campus, as chair of the biological chemistry department.
Neufeld’s research opened the way for prenatal diagnosis of such life-threatening fetal disorders as Hurler syndrome. Neufeld chaired the Scientific Advisory Board of the National MPS Society and was president of the American Society for Biochemistry and Molecular Biology from 1992 to 1993. She was elected to both the National Academy of Sciences (USA) and the American Academy of Arts and Sciences in 1977 and named a fellow of the American Association for Advancement in Science in 1988. In 1990 she was named California Scientist of the Year. She was awarded the Wolf Prize, the Albert Lasker Award for Clinical Medical Research, and was awarded the National Medal of Science in 1994 “for her contributions to the understanding of the lysosomal storage diseases, demonstrating the strong linkage between basic and applied scientific investigation.”[3]
Jarvis “Jay” Edwin Seegmiller, M.D.
“Jay Seegmiller was one of the giants of American medicine,” said Edward Holmes, M.D., Vice Chancellor of Health Sciences and dean of the School of Medicine at UCSD. “He and his trainees have made innumerable contributions to our understanding of the pathogenesis of many human disorders. Seegmiller was one of the country’s leading researchers in intermediary metabolism, with a focus on purine metabolism and inherited metabolism. He worked in the field of human biochemical genetics, with a special interest in the mechanisms by which genetically determined defects of metabolism lead to various forms of arthritis. His laboratory identified a wide range of primary metabolic defects in metabolism responsible for development of gout.
He is perhaps best known for his discovery of the enzyme defect in Lesch-Nyhan Syndrome, a fatal disorder of the nervous system causing severe mental retardation and self-mutilation impulses. As Director of the Human Biochemical Genetics Program at UCSD, Seegmiller’s investigations into the translation of genetic research and methods of prevention, detection and treatment of hereditary diseases led to Congressional testimony on the possibility of controlling genetic disease in the United States. As a result, genetic referral centers have been established throughout the country.
He joined the newly established UCSD School of Medicine in 1969 as head of the Arthritis Division of the Department of Medicine. There, he directed a research program in human biochemical genetics involving senior faculty from five departments within the School of Medicine. While a professor at UCSD, he served as a Macy Scholar both at Oxford University and at the Basel Institute in Switzerland, as well as a Guggenheim Fellow at the Swiss Institute for Experimental Cancer Research in Lausanne.
In 1983, he became the founding director of what is today UCSD’s Stein Institute for Research on Aging (SIRA). Even after his retirement, he continued to serve as Associate Director of SIRA from 1990 until his death.
“He had the foresight of proposing the formation of and then establishing a new Institute on Aging at UCSD before there was any such Institute in the entire UC system,” said Dilip Jeste, M.D., the Estelle and Edgar Levi Chair in Aging, Professor of Psychiatry and Neurosciences and current Director of SIRA. “He was himself a role model of successful aging, and continued working in the SIRA till his very last days.
Seegmiller received his Doctor of Medicine with honors from the University of Chicago in 1948. After he completed his internship at Johns Hopkins Hospital in Baltimore, Maryland, he trained with Bernard Horecker of the National Institute of Arthritis and Metabolic Disease at the National Institutes of Health.
Seegmiller was appointed Senior Investigator of the National Institute of Arthritis and Metabolic Disease in 1954, where he carried out biochemical and clinical studies of human hereditary disease, with a special interest in those causing various forms of arthritis. He became Assistant Scientific Director of the Institute in 1960, and was appointed Chief of the section on Human Biochemical Genetics in 1966, becoming one of several NIH leaders recruited to help launch UC San Diego’s new medical school.
Seegmiller’s clinical activities included studies in life longevity in South America. In 1974, he joined a team of notable scientists and traveled to the remote village of Vilcabamba in Ecuador, to find out what role genetic factors played in the population of the Andean villagers who comprised some of the longest-living people in the world. His later work led to the discovery of free radicals and their damaging effects in the human ability to withstand diseases, bringing forward new investigations on human aging at SIRA.
Seegmiller was a member of the National Academy of Sciences, the American Academy of Arts and Sciences, and was the recipient of numerous prizes and awards in honor of his extraordinary achievements in science and medicine. He received the United States Public Health Distinguished Service Award in 1969; and was honored as Master of the American College of Rheumatology (ACR) in 1992. He was on the advisory boards for the National Genetics Foundation, the City of Hope Medical Center in Duarte, California, the Task Force on Endocrinology and Metabolism for NIH, the Executive Editorial Board for Analytical Biochemistry, and was President of the Western Association of Physicians in 1979.
What has changed?
The 21st century has seen the mapping of the human genome. The huge focus on the genome came after the Watson and Crick discovery put the genome at the center of the translational network with the central hypothesis. What followed was transcription of RNA into placement of an amino acid into protein. The central hypothesis is DNA RNA protein. However, RNAi and non-translational RNA are now also important. RNA has a role in suppressing translation, as do proteins by allosteric effects. In addition, the most common diseases involved in age related change are strongly responsive to extracellular matrix effects, ionic fluxes, effects on the cellular matrix, and involve multicentric genome expression. This mode of expression leads one to think hard about the therapeutic target, or targets. The effect of RNA or of protein interacting with the genome is not an element of the classic construct.
Identifying a part of the problem
Type 2 diabetes mellitus, hypertension, arrhythmias, atherosclerotic plaque development, cancer, congestive heart disease, pulmonary hypertension, pulmonary interstitial sclerosis, and renovascular disease are among the common diseases that develop during a lifetime. The phenotypic presentations may have genomic associations, and there may also be population variants. There is also a cross-talk between these phenotypic expressions. Classically, medical terminology has been based on signs and symptoms of disease. In the increasingly complex experience, the laboratory has played an increased role in the diagnosis as well as prognostication. The laboratory experience with respect to the practice of medicine has heavily relied of either proteins, enzymes, or the products of chemical reactions. The use of genomic profiling has rapidly emerged in the laboratory armamentarium, but has had a slow ascent in practice.
Case in Point. Pompe’s disease
William Canfield is a glycobiologist, chief scientific officer and founder of an Oklahoma City-based biotechnology company, Novazyme, which was acquired by Genzyme in August 2001 and developed, among other things, an enzyme that can stabilize (but not cure) Pompe disease, based on Canfield’s ongoing research since 1998.[1][2]
John Crowley took over a position as a CEO in Novazyme after leaving Bristol-Myers Squibb in March 2000 and together with Dr. Y. T. Chen[4] at Duke University pushed for expedited approval by the U.S. Food and Drug Administration (FDA) of a new drug compound, NZ-1001 under orphan drug designation for the treatment of Glycogen storage disease type II in October 2005. The FDA stated: “We have determined that Novazyme’s recombinant human highly phosphorylated acid alpha-glucosidase (rhHPGAA) qualifies for orphan designation for enzyme replacement therapy in patients with all subtypes of glycogen storage disease type II (Pompe’s disease).” [5][6] Subsequent research at Genzyme on NZ-1001 along with three other potential compounds brought approval of the first enzyme replacement therapy for Pompe’s disease – Alglucosidase alfa (Myozyme or Lumizyme, Genzyme Inc) in 2006.[7]
William Canfields work with Pompes Disease was fictionalized and made the subject of a 2010 movie Extraordinary Measures in which he is called Dr. Robert Stonehill and played by Harrison Ford.[8]
Case in point. Polymorphisms in the long non-coding RNA
Hypertension (HT) is a complex disorder influenced by both genetic and environmental factors. Recent genome-wide association studies have identified a major risk locus for atherosclerosis on chromosome 9p21.3 (chr9p21.3). SNPs within the coding sequences of CDKN2A/B proteins and the long non-coding RNA CDKN2B-AS1 could potentially contribute to HT development. Such a study has now been done. The findings suggest that SNPs rs10757274, rs2383207, rs10757278, and rs1333049, particularly those within the CDKN2B-AS1 gene, and related haplotypes may confer increased susceptibility to HT development. (unpublished)
Case in point. Lipoprotein Lipase and Atherosclerosis
Lipoprotein lipase (LPL) plays a pivotal role in lipids and metabolism of lipoprotein. Dysfunctions of LPL have been found to be associated with dyslipidemia, obesity and insulin resistance.Dyslipidemia, obesity and insulin resistance are risk factor of atherosclerosis. Japanese investigators have hypothesized that elevating LPL activity would cause protection of atherosclerosis. (unpublished).
Case in point. Holocaust survivors pass on stress.
Descendants of Holocaust Survivors Have Altered Stress Hormones
Parents’ traumatic experience may hamper their offspring’s ability to bounce back from trauma
A person’s experience as a child or teenager can have a profound impact on their future children’s lives, new work is showing. Rachel Yehuda, a researcher in the growing field of epigenetics and the intergenerational effects of trauma, and her colleagues have long studied mass trauma survivors and their offspring. Their latest results reveal that descendants of people who survived the Holocaust have different stress hormone profiles than their peers, perhaps predisposing them to anxiety disorders.
Yehuda’s team at the Icahn School of Medicine at Mount Sinai and the James J. Peters Veterans Affairs Medical Center in Bronx, N.Y., and others had previously established that survivors of the Holocaust have altered levels of circulating stress hormones compared with other Jewish adults of the same age. Survivors have lower levels of cortisol, a hormone that helps the body return to normal after trauma; those who suffered post-traumatic stress disorder (PTSD) have even lower levels.
Case in point. Genome engineering with CRISPR-Cas9
The new frontier of genome engineering with CRISPR-Cas9
BACKGROUND: Technologies for making and manipulating DNA have enabled advances in biology ever since the discovery of the DNA double helix. But introducing site-specific modifications in the genomes of cells and organisms remained elusive. Early approaches relied on the principle of site-specific recognition of DNA sequences by oligonucleotides, small molecules, or self-splicing introns. More recently, the site-directed zinc finger nucleases (ZFNs) and TAL effector nucleases (TALENs) using the principles of DNAprotein recognition were developed. However, difficulties of protein design, synthesis, and validation remained a barrier to
SUMMARY
The field of biology is now experiencing a transformative phase with the advent of facile genome engineering in animals and plants using RNA-programmable CRISPR-Cas9. The CRISPR-Cas9 technology originates from type II CRISPR-Cas systems, which provide bacteria with adaptive immunity to viruses and plasmids. The CRISPR associated protein Cas9 is an endonuclease that uses a guide sequence within an RNA duplex, tracrRNA:crRNA, to form base pairs with DNA target sequences, enabling Cas9 to introduce a site-specific double-strand break in the DNA. The dual tracrRNA:crRNA was engineered as a single guide RNA (sgRNA) that retains two critical features: a sequence at the 5 side that determines the DNA target site by Watson-Crick base-pairing and a duplex RNA structure at the 3 side that binds to Cas9. This finding created a simple two-component system in which changes in the guide sequence of the sgRNA program Cas9 to target any DNA sequence of interest. The simplicity of CRISPR-Cas9 programming, together with a unique DNA cleaving mechanism, the capacity for multiplexed target recognition, and the existence of many natural type II CRISPR-Cas system variants, has enabled remarkable developments using this cost-effective and easy-to-use technology to precisely and efficiently target, edit, modify, regulate, and mark genomic loci of a wide array of cells and organisms.
Figure (not shown)
The Cas9 enzyme (blue) generates breaks in double-stranded DNA by using its two catalytic centers (blades) to cleave each strand of a DNA target site (gold) next to a PAM sequence (red) and matching the 20-nucleotide sequence (orange) of the single guide RNA (sgRNA). The sgRNA includes a dual-RNA sequence derived from CRISPR RNA (light green) and a separate transcript (tracrRNA, dark green) that binds and stabilizes the Cas9 protein. Cas9-sgRNA–mediated DNA cleavage produces a blunt double-stranded break that triggers repair enzymes to disrupt or replace DNA sequences at or near the cleavage site. Catalytically inactive forms of Cas9 can also be used for programmable regulation of transcription and visualization of genomic loci.
This Review illustrates the power of the technology to systematically analyze gene functions in mammalian cells, study genomic rearrangements and the progression of cancers or other diseases, and potentially correct genetic mutations responsible for inherited disorders. CRISPR-Cas9 is having a major impact on functional genomics conducted in experimental systems. Its application in genome-wide studies will enable large-scale screening for drug targets and other phenotypes and will facilitate the generation of engineered animal models that will benefit pharmacological studies and the understanding of human diseases. CRISPR-Cas9 applications in plants and fungi also promise to change the pace and course of agricultural research. Future research directions to improve the technology will include engineering or identifying smaller Cas9 variants with distinct specificity that may be more amenable to delivery in human cells. Understanding the homology-directed repair mechanisms that follow Cas9-mediated DNA cleavage will enhance insertion of new or corrected sequences into genomes. The development of specific methods for efficient and safe delivery of Cas9 and its guide RNAs to cells and tissues will also be critical for applications of the technology in human gene therapy.
Case in point.
ZFN, TALEN and CRISPR/Cas-based methods for genome engineering
Thomas Gaj1,2,3, Charles A. Gersbach4,5, and Carlos F. Barbas III1,2,3 1The Skaggs Institute for Chemical Biology, The Scripps Research Institute, La Jolla, CA, USA 2Department of Molecular Biology, The Scripps Research Institute, La Jolla, CA, USA 3Department of Chemistry, The Scripps Research Institute, La Jolla, CA, USA 4Department of Biomedical Engineering, Duke University, Durham, NC, USA 5Institutes for Genome Sciences and Policy, Duke University, Durham, NC, USA
Abstract Zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs) comprise a powerful class of tools that are redefining the boundaries of biological research. These chimeric nucleases are composed of programmable, sequence-specific DNA-binding modules linked to a non-specific DNA cleavage domain. ZFNs and TALENs enable a broad range of genetic modifications by inducing DNA double-strand breaks that stimulate error-prone nonhomologous end joining or homology-directed repair at specific genomic locations. Here, we review achievements made possible by site-specific nuclease technologies and discuss applications of these reagents for genetic analysis and manipulation. In addition, we highlight the therapeutic potential of ZFNs and TALENs and discuss future prospects for the field, including the emergence of CRISPR/Cas-based RNA-guided DNA endonucleases.
Classical and contemporary approaches for establishing gene function With the development of new and affordable methods for whole-genome sequencing, and the design and implementation of large-scale genome annotation projects, scientists’ are poised to deliver upon the promises of the Genomic Revolution to transform basic science and personalized medicine. The resulting wealth of information presents researchers with a new primary challenge of converting this enormous amount of data into functionally and clinically relevant knowledge. Central to this problem is the need for efficient and reliable methods that enable investigators to determine how genotype influences phenotype. Targeted gene inactivation via homologous recombination is a powerful method capable of providing conclusive information for evaluating gene function.
Several factors impede the use of these methods:
the low efficiency at which engineered constructs are correctly inserted into the chromosomal target site,
the need for time-consuming and labor-insensitive selection/screening strategies, and
the potential for adverse mutagenic effects.
Targeted gene knockdown by RNA interference (RNAi) has provided researchers with a rapid, inexpensive and high-throughput alternative to homologous recombination. However, knockdown by RNAi is incomplete, varies between experiments and laboratories, has unpredictable off-target effects, and provides only temporary inhibition of gene function. These restrictions impede researchers’ ability to directly link phenotype to genotype and limit the practical application of RNAi technology.
In the past decade, a new approach has emerged that enables investigators to directly manipulate virtually any gene in a diverse range of cell types and organisms. This core technology – commonly referred to as “genome editing” – is based on the use of engineered nucleases composed of sequence-specific DNA-binding domains fused to a non-specific DNA cleavage module. These chimeric nucleases enable efficient and precise genetic modifications by inducing targeted DNA double-strand breaks (DSBs) that stimulate the cellular DNA repair mechanisms, including error-prone non-homologous end joining (NHEJ) and homology-directed repair (HDR).
Case in point.
CRISPR/Cas9 and Targeted Genome Editing: A New Era in Molecular Biology
The development of efficient and reliable ways to make precise, targeted changes to the genome of living cells is a long-standing goal for biomedical researchers. Recently, a new tool based on a bacterial CRISPR-associated protein-9 nuclease (Cas9) from Streptococcus pyogenes has generated considerable excitement. This follows several attempts over the years to manipulate gene function, including homologous recombination and RNA interference (RNAi).
RNAi, in particular, became a laboratory staple enabling inexpensive and high-throughput interrogation of gene function, but it is hampered by providing only temporary inhibition of gene function and unpredictable off-target effects. Other recent approaches to targeted genome modification – zinc-finger nucleases (ZFNs), and transcription-activator like effector nucleases (TALENs) – enable researchers to generate permanent mutations by introducing double stranded breaks to activate repair pathways. These approaches are costly and time-consuming to engineer, limiting their widespread use, particularly for large scale, high-throughput studies.
The Biology of Cas9
The functions of CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats) and CRISPR-associated (Cas) genes are essential in adaptive immunity in select bacteria and archaea, enabling the organisms to respond to and eliminate invading genetic material. These repeats were initially discovered in the 1980s in E. coli, but their function wasn’t confirmed until 2007 by Barrangou and colleagues, who demonstrated that S. thermophilus can acquire resistance against a bacteriophage by integrating a genome fragment of an infectious virus into its CRISPR locus.
Three types of CRISPR mechanisms have been identified, of which type II has been the most studied. In this case, invading DNA from viruses or plasmids is cut into small fragments and incorporated into a CRISPR locus amidst a series of short repeats (around 20 bps). The loci are transcribed, and transcripts are then processed to generate small RNAs (crRNA – CRISPR RNA), which are used to guide effector endonucleases that target invading DNA based on sequence complementarity (Figure 1) (not shown).
In the acquisition phase, foreign DNA is incorporated into the bacterial genome at the CRISPR loci. CRISPR loci is then transcribed and processed into crRNA during crRNA biogenesis. During interference, Cas9 endonuclease complexed with a crRNA and separate tracrRNA cleaves foreign DNA containing a 20-nucleotide crRNA complementary sequence adjacent to the PAM sequence.
Investment in CRISPR technology
CRISPR Therapeutics is a biopharmaceutical company created to translate CRISPR-Cas9, a breakthrough genome-editing technology, into transformative medicines for serious human diseases. We are uniquely positioned to translate CRISPR-Cas9 technology into human therapeutics, thanks to its multi-disciplinary team of world-renowned academics, clinicians and drug developers.
CRISPR Therapeutics’ vision is to cure serious human diseases at the molecular level using CRISPR-Cas9. The company is headquartered in Basel, Switzerland and has operations in London, UK and Cambridge, Massachusetts.
The biopharmaceutical company that is focused on translating CRISPR-Cas9 gene-editing technology into transformative medicines for serious human diseases, congratulates its scientific founder, Dr. Emmanuelle Charpentier, for being named to TIME Magazine’s TIME 100 Most Influential People in the World alongside fellow CRISPR-Cas9 discoverer, Dr. Jennifer Doudna. In addition, Dr. Emmanuelle was awarded the Louis Jeantet Prize for Medicine, considered the most prestigious European award for researchers in the life sciences, for her discovery of the CRISPR-Cas9 gene editing tool. She will receive the award in a ceremony in Geneva, Switzerland, on April 22, 2015.
Dr. Charpentier has received numerous additional awards for her research, including in 2014 the Alexander von Humboldt Professorship, the Dr Paul Janssen Award, the Grand-Prix Jean-Pierre Lecocq (French Academy of Sciences), the Göran Gustafsson Prize (Royal Swedish Academy of Sciences) and in 2015 the Breakthrough Prize in Life Sciences. She was also selected as one of the American Foreign Policy magazine’s 100 Leading Global Thinkers for 2014.
Cambridge-based Editas Medicine announced a $120 million Series B round led by Bill Gates’s chief advisor for science and technology, Boris Nikolic. The list of financiers teaming with Nikolic reads like a rolodex of so-called crossover investors. Nikolic, who joined Editas’ board, made the investment through what’s been called “bng0,” a new U.S.-based investment company backed by “large family offices with a global presence and long-term investment horizon” and formed specifically to invest in Editas. CEO Katrine Bosley confirmed that Gates is one of the individuals investing in Editas alongside Nikolic. Editas has become the first of the group not only to attract crossover backers, but to begin discussing the diseases that its targeting.
Caribou Biosciences, one of the biotech startups working to advance a much-watched new technology for precise gene editing, has raised an $11 million Series A round from venture capital firms and Swiss drug giant Novartis.
The money will help Berkeley, CA-based Caribou speed up its efforts to adapt a versatile genome editing technique co-discovered by one of its founders, UC Berkeley professor Jennifer Doudna, for a range of uses, including drug research and development, and industrial technology.
Doudna and her collaborator, Emmanuelle Charpentier of the Helmholtz Center for Infection Research in Braunschweig, Germany, and Umeå University in Sweden, figured out how to transform a bacterial defense against viral infection into a tool to edit out abnormal sections of genes, such as those that cause hereditary diseases.
Caribou’s gene editing platform is based on two elements of that bacterial molecular machinery: a guiding mechanism called CRISPR (clustered, regularly interspaced palindromic repeats), and an enzyme called Cas9, or CRISPR-associated protein 9, molecular scissors that cut a segment of DNA. Caribou was founded in 2011 to commercialize the work from Doudna’s lab.
Researchers have hijacked a defense system normally used by bacteria to fend off viral infections and redirected it against the human papillomavirus (HPV), the virus that causes cervical, head and neck, and other cancers.
Using the genome editing tool known as CRISPR, the Duke University researchers were able to selectively destroy two viral genes responsible for the growth and survival of cervical carcinoma cells, causing the cancer cells to self-destruct.
The findings, published in the Journal of Virology, give credence to an approach only recently attempted in mammalian cells, and could pave the way toward antiviral strategies targeted against other DNA-based viruses like hepatitis B and herpes simplex.
“Because this approach is only going after viral genes, there should be no off-target effects on normal cells,” said Bryan R. Cullen, Ph.D., senior study author and professor of molecular genetics and microbiology at Duke University School of Medicine. “You can think of this as targeting a missile that will destroy a certain target. You put in a code that tells the missile exactly what to hit, and it will only hit that, and it won’t hit anything else because it doesn’t have the code for another target.”
In this study, Cullen decided to target the human papillomavirus (HPV), which causes almost all cervical cancers and about half of head and neck cancers. Specifically, he and his colleagues went after the viral genes E6 and E7, two “oncogenes” that block the host’s own efforts to keep cancer cells at bay.
To run CRISPR against the virus, the researchers needed two ingredients. First, they needed the target code for E6 or E7, consisting of a short strip of RNA sequence, the chemical cousin of DNA. To this “guide RNA” they added the Cas9 protein, which would cut any DNA that could line up and bind to that RNA sequence.
The carcinoma cells that received the anti-HPV guide RNA/Cas9 combination immediately stopped growing. In contrast, cells that had received a control virus, containing a random guide RNA sequence, continued on their path to immortality. The researchers then dug down to the molecular level to investigate the consequences of destroying E6 or E7 in cancer cells. E6 normally blocks a protein called p53, known as the guardian of the genome because it can turn on suicide pathways in the cell when it senses that something has gone awry. In this study, targeting E6 enabled p53 to resume its normal function, spurring death of the cancer cell.
E7 works in a similar way, blocking another protein called retinoblastoma or Rb that can trigger growth arrest and senescence, another form of cell death. As expected, the researchers found that targeting E7 also set this second “tumor suppressor” back in motion.
“As soon as you turn off E6 or E7, the host defense mechanisms are allowed to come back on again, because they have been there this whole time, but they have been turned off by HPV,” Cullen said. “What happens is the cell immediately commits suicide.”
Cullen and his colleagues are now working on developing a different viral vector, based on the adeno-associated virus, to deliver their CRISPR cargo into cancer cells. Once they are happy with their delivery system, they will begin to test this approach in animal models.
“What we would hope to see in an HPV-induced cancer is rapid induction of tumor necrosis caused by loss of E6 or E7,” Cullen said. “This method has the potential to be a single hit treatment that will dramatically reduce tumor load without having any effect on normal cells.”
The researchers are also targeting other viruses that use DNA as their genetic material, including the hepatitis B virus and herpes simplex virus.
Reference: “Inactivation of the human papillomavirus E6 or E7 gene in cervical carcinoma cells using a bacterial CRISPR/Cas RNA-guided endonuclease,” Edward M. Kennedy, Anand V. R. Kornepati, Michael Goldstein, Hal P. Bogerd, Brigid C. Poling, Adam W. Whisnant, Michael B. Kastan and Bryan R. Cullen.Journal of Virology, August 6, 2014. DOI 10.1128/JVI.01879-14.
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