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Human gene editing continues to hold a major fascination within a biomedical and biopharmaceutical industries. It’s extraordinary potential is now being realized but important questions as to who will be the beneficiaries of such breakthrough technologies remained to be answered. The session will discuss whether gene editing technologies can alleviate some of the most challenging unmet medical needs. We will discuss how research advances often never reach minority communities and how diverse patient populations will gain access to such breakthrough technologies. It is widely recognize that there are patient voids in the population and we will explore how community health centers might fill this void to ensure that state-of-the-art technologies can reach the forgotten patient groups . We also will touch ethical questions surrounding germline editing and how such research and development could impact the community at large.

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CRISPR cuts turn gels into biological watchdogs

Reporter: Irina Robu, PhD

Genome editing if of significant interest in the prevention and treatment of human diseases including single-gene disorders such as cystic fibrosis, hemophilia and sickle cell disease. It also shows great promise for the prevention and treatment of diseases such as cancer, heart disease, mental illness and human immunodeficiency virus infection. However, ethical concerns arise when genome editing, using technologies such as CRISPR-Cas9 is used to alter human genomes.

James Collins, bioengineer at MIT and his team worked with water-filled polymers that are held together by strands of DNA, known as DNA hydrogels. To alter the properties of these materials, these scientists turned to a form of CRISPR that uses a DNA-snipping enzyme called Cas12a, which can be programed to recognize a specific DNA sequence. The enzyme then cuts its target DNA strand, then severs single strands of DNA nearby. This property lets scientists to build a series of CRISPR-controlled hydrogels encapsulating a target DNA sequence and single strands of DNA, which break up after Cas12a identifies the target sequence in a stimulus. The break-up of the single DNA strands activates the hydrogels to change shape or completely dissolve, releasing a payload.

According to Collins and his team, the programmed hydrogels will release enzymes, small molecules and human cells as part of a smart therapy in response to stimuli. However, in order to make it a smart therapeutic, the researchers in collaboration with Dan Luo, bioengineer at Cornell University placed the CRISPR- controlled hydrogels into electric circuits. The circuit is switched off in response to the detection of the genetic material of harmful pathogens such as Ebola virus and methicillin-resistant Staphylococcus aureus. The team used these hydrogels to develop a prototype diagnostic tool that sends a wireless signal to identify Ebola in lab samples.

Yet, it is evident that these CRISPR-controlled hydrogels show great potential for the prevention and treatment of diseases.

SOURCE

https://www.nature.com/articles/d41586-019-02542-3?utm_source=Nature+Briefing

 

 

 

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Optimization of CRISPR Gene Editing with Gold Nanoparticles

Reporter: Irina Robu, PhD

The CRISPR-Cas9 gene editing system has been welcomed as a hopeful solution to a range of genetic diseases, but the expertise has proven hard to deliver into cells. One plan is to open the cell membrane using an electric shock, but that can accidentally kill the cell. Another is to use viruses as couriers. Problem is, viruses can cause off-target side effects.

CRISPR-Cas9 is a unique technology that enables geneticists and medical researchers to edit parts of the genome by removing, adding or altering sections of DNA sequence. It is faster, cheaper and more accurate than previous techniques of editing DNA and can have a wide range of potential applications.

The CRISPR-Cas9 system consists of two key molecules that introduce a change into the DNA. One is an enzyme called Cas9 which acts as a pair of molecular scissors that can cut the two strands of DNA at a specific location in the genome where bits of DNA can be added or removed. The other one, is a piece of RNA which consists of a small piece of pre-designed RNA sequence located within a longer RNA scaffold. The scaffold part binds to the DNA and pre-designed sequence which contains Cas9. The RNA sequence is designed to find and locate specific sequence in the DNA. The Cas9 trails the guide RNA to the same location in the DNA sequence and makes a cut across both strands of DNA. At this point the cell distinguishes that the DNA is damaged and tries to repair it.

Researchers at Fred Hutchinson Cancer Research Center published new findings in Nature Materials suggested an alternative delivery method such as gold nanoparticles. The gold nanoparticles are packed with all the CRISPR components necessary to make clean gene edits. When the gold nanoparticles were tested in lab models of inherited blood disorders and HIV, between 10% and 20% of the targeted cells were effectively edited, with no toxic side effects.

The researchers use gold nanoparticles to deliver CRISPR to blood stem cells. Each gold nanoparticle contains four CRISPR components, including the enzyme needed to make the DNA cuts. But Fred Hutchinson researchers chose Cas12a, which they believed would lead to more efficient edits. Plus, Cas12a only needs one molecular guide, while Cas9 requires two.
In one experiment, they sought to disturb the gene CCR5 to make cells resistant to HIV. In the second, they created a gene mutation that can protect against blood disorders, including sickle cell disease. They observed the cells encapsulated the nanoparticles within six hours and began the gene-editing process within 48 hours. In mice, gene editing peaked eight weeks after injection, and the edited cells were still in circulation 22 weeks after the treatment.
Researchers at Fred Hutchinson are now working on improving the efficiency of the gold-based CRISPR delivery system so that 50% or more of the targeted cells are edited and are also looking for a commercial partner to bring the technology to clinical phase in the next few years.

SOURCE:

https://www.fiercebiotech.com/research/fred-hutch-team-uses-gold-nanoparticles-to-improve-crispr-gene-editing

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Advances in Gene Editing and Gene Silencing | September 20-21, 2016 | Boston, MA

Kinase Inhibitor Discovery September 21-22, 2016 Boston

KEYNOTE SESSION: GENOME EDITING FOR IN VIVO APPLICATIONS

Part 1 (of a two-part conference) will cover the use of CRISPR/Cas9 and RNAi for identifying new drug targets and therapies. It will bring together experts from all aspects of basic science and clinical research to talk about how and where gene editing and RNAi can be best applied. What are the different tools that can be used and what are their strengths and limitations? How does the CRISPR/Cas system compare to RNAi and other gene editing tools, such as Transcription Activator-like Effector Nucleases (TALENs) and zinc finger nucleases (ZFNs), and do they have any complementary uses? Scientists and clinicians from pharma/biotech as well as from academic and government labs will share their experiences leveraging the utility of gene editing for target discovery, disease modeling, and for creating cell and viral therapies. Learn more atDiscoveryOnTarget.com/RNAi-screens-functional-genomics

Advance Registration Discount Available!
Register by August 12 Week to Save up to $200

Keynote Session: Genome Editing for In Vivo Applications

AAV for Gene Therapy and Genome Editing
James Wilson, M.D., Ph.D., Professor, Department of Pathology and Laboratory Medicine, Perelman School of Medicine; Director, Orphan Disease Center and Director, Gene Therapy Program, University of Pennsylvania
In vivo delivery of nucleic acid therapeutics remains the primary barrier to success. My lab has focused on the use of vectors based on adeno-associated virus (AAV) for achieving success in pre-clinical and clinical applications of gene replacement therapy. Most of the current academic and commercial applications of in vivo gene replacement therapy are based on endogenous AAVs we discovered as latent viral genomes in primates. These vectors are reasonably safe and efficient for application of gene replacement therapy. The emergence of genome editing methods has suggested more precise and effective methods to treat inherited diseases in which genes are silenced or mutations are corrected. AAV vectors have been the most efficient platform for achieving genome editing in vivo. We will review our attempts to achieve therapeutic genome editing in animal models of liver disease using AAV.

Using CRISPR/Cas to Target and Destroy Viral DNA Genomes
Bryan R. Cullen, Ph.D., James B. Duke Professor of Molecular Genetics and Microbiology and Director, Center for Virology, Duke University
A number of pathogenic human DNA viruses, including HBV, HIV-1 and HSV1, cause chronic diseases in humans that remain refractory to cure, though these diseases can be controlled by antivirals. In addition the DNA virus HPV causes tumors that depend on the continued expression of viral genes. Here, I will present data demonstrating that several of these viruses can be efficiently cleaved and destroyed using viral vectors that express Cas9 and virus-specific guide RNAs, thus providing a potential novel approach to treatment.

Targeted Endonucleases as Antiviral Agents: Promises and Pitfalls
Keith R. Jerome, M.D., Ph.D., Member, Vaccine and Infectious Disease Division, Fred Hutchinson Cancer Research Center; Professor and Head, Virology Division, Department of Laboratory Medicine, University of Washington
Genome editing offers the prospect of cure for infections such as HIV, hepatitis B virus, herpes simplex, and human papillomavirus, by disruption of essential viral nucleic acids or the human genes encoding receptors needed for viral entry. This talk will highlight the most recent laboratory data and the challenges still ahead in bringing this technology to the clinic.

Nucleic Acid Delivery Systems for RNA Therapy and Gene Editing
Daniel Anderson, Ph.D., Professor, Department of Chemical Engineering, Institute for Medical Engineering & Science, Harvard-MIT Division of Health Sciences & Technology and David H. Koch Institute for Integrative Cancer Research, Massachusetts Institute of Technology
High throughput, combinatorial approaches have revolutionized small molecule drug discovery. Here we describe our high throughput methods for developing and characterizing RNA delivery and gene editing systems. Libraries of degradable polymers and lipid-like materials have been synthesized, formulated and screened for their ability to deliver RNA, both in vitro and in vivo. A number of delivery formulations have been developed with in vivo efficacy, and show potential applications for the treatment of genetic diseases, viral infections and cancers.

PANEL DISCUSSION: CRISPR/Cas: A Realistic and Practical Look at What the Future Could Hold
Moderator: Bryan R. Cullen, Ph.D., James B. Duke Professor of Molecular Genetics and Microbiology and Director, Center for Virology, Duke University
Participants: Session Speakers
Each speaker will spend a few minutes sharing their viewpoints and experiences on where things stand with using the CRISPR/Cas system for in vivo applications. Attendees will have an opportunity to ask questions and share their opinions.

About the Conference

Cambridge Healthtech Institute’s 13th annual two-part conference on Advances in Gene Editing and Gene Silencing will cover the latest in the use of CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas9-based gene editing and RNA interference (RNAi) for use in drug discovery and for developing novel drug therapies.


For sponsorship and exhibit sales information including podium presentation opportunities, contact:
Jon Stroup | T: +1 781-972-5483 | E: jstroup@healthtech.com


Recommended All Access Package:
Includes access to 1 Symposium and 2 Conferences

September 19 Symposium: 
Understanding CRISPR: Mechanisms and Applications

September 20-21 Conference:
Advances in Gene Editing and Gene Silencing – Part 1

September 21-22 Conference: 
Advances in Gene Editing and Gene Silencing – Part 2


Cambridge Healthtech Institute, 250 First Avenue, Suite 300, Needham, MA, USA

Tel: 781-972-5400 | Fax: 781-972-5425 | www.healthtech.com
This email is being sent to sjwilliamspa@comcast.net for marketing purposes. If it is not of interest to you, please disregard and we apologize for any inconvenience this may have caused.

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CRISPR’s Unwanted off-target effects: Need for safety study designs with Gene-Editing

Reporter: Stephen J. Williams, Ph.D.

From CafePharma at https://www.statnews.com/2016/07/18/crispr-off-target-effects/

Do CRISPR enthusiasts have their head in the sand about the safety of gene editing?

WASHINGTON — At scientific meetings on genome-editing, you’d expect researchers to show pretty slides of the ribbony 3-D structure of the CRISPR-Cas9 molecules neatly snipping out disease-causing genes in order to, everyone hopes, cure illnesses from cancer to muscular dystrophy. Less expected: slides of someone kneeling on a beach with his head in the sand.

Yet that is what Dr. J. Keith Joung of Massachusetts General Hospital showed at the American Society of Hematology’s workshop on genome-editing last week in Washington. While the 150 experts from industry, academia, the National Institutes of Health, and the Food and Drug Administration were upbeat about the possibility of using genome-editing to treat and even cure sickle cell disease, leukemia, HIV/AIDS, and other blood disorders, there was a skunk at the picnic: an emerging concern that some enthusiastic CRISPR-ers are ignoring growing evidence that CRISPR might inadvertently alter regions of the genome other than the intended ones.

“In the early days of this field, algorithms were generated to predict off-target effects and [made] available on the web,” Joung said. Further research has shown, however, that such algorithms, including one from MIT and one calledE-CRISP, “miss a fair number” of off-target effects. “These tools are used in a lot of papers, but they really aren’t very good at predicting where there will be off-target effects,” he said. “We think we can get off-target effects to less than 1 percent, but we need to do better,” especially if genome-editing is to be safely used to treat patients.

Off-target effects occur because of how CRISPR works. It has two parts. RNA makes a beeline for the site in a genome specified by the RNA’s string of nucleotides, and an enzyme cuts the genome there. Trouble is, more than one site in a genome can have the same string of nucleotides. Scientists might address CRISPR to the genome version of 123 Main Street, aiming for 123 Main on chromosome 9, only to find CRISPR has instead gone to 123 Main on chromosome 14.

In one example Joung showed, CRISPR is supposed to edit a gene called VEGFA (which stimulates production of blood vessels, including those used by cancerous tumors) on chromosome 6. But, studies show, this CRISPR can also hit genes on virtually every one of the other 22 human chromosomes. The same is true for CRISPRs aimed at other genes. Although each CRISPR has zero to a dozen or so “known” off-target sites (where “known” means predicted by those web-based algorithms), Joung said, there can be as many as 150 “novel” off-target sites, meaning scientists had no idea those errors were possible.

One reason for concern about off-target effects is that genome-editing might disable a tumor-suppressor gene or activate a cancer-causing one. It might also allow pieces of two different chromosomes to get together, a phenomenon called translocation, which is the cause of chronic myeloid leukemia, among other problems.

Many researchers, including those planning clinical trials, are using web-based algorithms to predict which regions of the genome might get accidentally CRISPR’d. They include the scientists whose proposal to use CRISPR in patients was the first to be approved by an NIH committee. When scientists assure regulators that they looked for off-target effects in CRISPR’d cells growing in lab dishes, what they usually mean is that they looked for CRISPR’ing of genes that the algorithms flagged.

As a result, off-target effects might be occurring but, because scientists are doing the equivalent of the drunk searching for their lost keys only under the lamppost, they’re not being found.

Other articles on CRISPR and Gene Editing on this Open Access Journal Include:

FDA Cellular & Gene Therapy Guidances: Implications for CRSPR/Cas9 Trials

CRISPR/Cas9 Finds Its Way As an Important Tool For Drug Discovery & Development

CRISPR, the Genome Editing Technology is Nearing Human Trials: Human T cells will soon be modified using the CRISPR technique in a clinical trial to attack cancer cells

Use of CRISPR & RNAi for Drug Discovery, CHI’s World PreClinical Congress – Europe, November 14-15, 2016, Lisbon, Portugal

CRISPR: A Podcast from Nature.com on Gene Editing

AND Please See Our Following ebooks available on Amazon containing interviews with Dr. Jennifer Duodna

Volume One: Genomics Orientations for Personalized Medicine

Volume Two: Latest in Genomics Methodologies for Therapeutics: Gene Editing, NGS & BioInformatics, Simulations and the Genome Ontology

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Gene Editing with CRISPR gets Crisper

Curators: Larry H. Bernstein, MD, FCAP and Aviva Lev-Ari, PhD, RN

 

 

CRISPR Moves from Butchery to Surgery   

More Genomes Are Going Under the CRISPR Knife, So Surgical Standards Are Rising

http://www.genengnews.com/gen-articles/crispr-moves-from-butchery-to-surgery/5759/

  • The Dharmacon subsidary of GE Healthcare provides the Edit-R Lentiviral Gene Engineering platform. It is based on the natural S. pyrogenes system, but unlike that system, which uses a single guide RNA (sgRNA), the platform uses two component RNAs, a gene-specific CRISPR RNA (crRNA) and a universal trans-activating crRNA (tracrRNA). Once hybridized to the universal tracrRNA (blue), the crRNA (green) directs the Cas9 nuclease to a specific genomic region to induce a double- strand break.

    Scientists recently convened at the CRISPR Precision Gene Editing Congress, held in Boston, to discuss the new technology. As with any new technique, scientists have discovered that CRISPR comes with its own set of challenges, and the Congress focused its discussion around improving specificity, efficiency, and delivery.

    In the naturally occurring system, CRISPR-Cas9 works like a self-vaccination in the bacterial immune system by targeting and cleaving viral DNA sequences stored from previous encounters with invading phages. The endogenous system uses two RNA elements, CRISPR RNA (crRNA) and trans-activating RNA (tracrRNA), which come together and guide the Cas9 nuclease to the target DNA.

    Early publications that demonstrated CRISPR gene editing in mammalian cells combined the crRNA and tracrRNA sequences to form one long transcript called asingle-guide RNA (sgRNA). However, an alternative approach is being explored by scientists at the Dharmacon subsidiary of GE Healthcare. These scientists have a system that mimics the endogenous system through a synthetic two-component approach thatpreserves individual crRNA and tracrRNA. The tracrRNA is universal to any gene target or species; the crRNA contains the information needed to target the gene of interest.

    Predesigned Guide RNAs

    In contrast to sgRNAs, which are generated through either in vitro transcription of a DNA template or a plasmid-based expression system, synthetic crRNA and tracrRNA eliminate the need for additional cloning and purification steps. The efficacy of guide RNA (gRNA), whether delivered as a sgRNA or individual crRNA and tracrRNA, depends not only on DNA binding, but also on the generation of an indel that will deliver the coup de grâce to gene function.

    “Almost all of the gRNAs were able to create a break in genomic DNA,” said Louise Baskin, senior product manager at Dharmacon. “But there was a very wide range in efficiency and in creating functional protein knock-outs.”

    To remove the guesswork from gRNA design, Dharmacon developed an algorithm to predict gene knockout efficiency using wet-lab data. They also incorporated specificity as a component of their algorithm, using a much more comprehensive alignment tool to predict potential off-target effects caused by mismatches and bulges often missed by other alignment tools. Customers can enter their target gene to access predesigned gRNAs as either two-component RNAs or lentiviral sgRNA vectors for multiple applications.

    “We put time and effort into our algorithm to ensure that our guide RNAs are not only functional but also highly specific,” asserts Baskin. “As a result, customers don’t have to do any design work.”

    Donor DNA Formats

    http://www.genengnews.com/Media/images/Article/thumb_MilliporeSigma_CRISPR3120824917.jpg
    MilliporeSigma’s CRISPR Epigenetic Activator is based on fusion of a nuclease-deficient Cas9 (dCas9) to the catalytic histone acetyltransferase (HAT) core domain of the human E1A-associated protein p300. This technology allows researchers to target specific DNA regions or gene sequences. Researchers can localize epigenetic changes to their target of interest and see the effects of those changes in gene expression.

    Knockout experiments are a powerful tool for analyzing gene function. However, for researchers who want to introduce DNA into the genome, guide design, donor DNA selection, and Cas9 activity are paramount to successful DNA integration.MilliporeSigma offers two formats for donor DNA: double-stranded DNA (dsDNA) plasmids and single-stranded DNA (ssDNA) oligonucleotides. The most appropriate format depends on cell type and length of the donor DNA. “There are some cell types that have immune responses to dsDNA,” said Gregory Davis, Ph.D., R&D manager, MilliporeSigma.

  • The ssDNA format can save researchers time and money, but it has a limited carrying capacity of approximately 120 base pairs.In addition to selecting an appropriate donor DNA format, controlling where, how, and when the Cas9 enzyme cuts can affect gene-editing efficiency. Scientists are playing tug-of-war, trying to pull cells toward the preferred homology-directed repair (HDR) and away from the less favored nonhomologous end joining (NHEJ) repair mechanism.One method to achieve this modifies the Cas9 enzyme to generate a nickase that cuts only one DNA strand instead of creating a double-strand break. Accordingly, MilliporeSigma has created a Cas9 paired-nickase system that promotes HDR, while also limiting off-target effects and increasing the number of sequences available for site-dependent gene modifications, such as disease-associated single nucleotide polymorphisms (SNPs).“The best thing you can do is to cut as close to the SNP as possible,” advised Dr. Davis. “As you move the double-stranded break away from the site of mutation you get an exponential drop in the frequency of recombination.”

 

  • Ribonucleo-protein Complexes

    Another strategy to improve gene-editing efficiency, developed by Thermo Fisher, involves combining purified Cas9 protein with gRNA to generate a stable ribonucleoprotein (RNP) complex. In contrast to plasmid- or mRNA-based formats, which require transcription and/or translation, the Cas9 RNP complex cuts DNA immediately after entering the cell. Rapid clearance of the complex from the cell helps to minimize off-target effects, and, unlike a viral vector, the transient complex does not introduce foreign DNA sequences into the genome.

    To deliver their Cas9 RNP complex to cells, Thermo Fisher has developed a lipofectamine transfection reagent called CRISPRMAX. “We went back to the drawing board with our delivery, screened a bunch of components, and got a brand-new, fully  optimized lipid nanoparticle formulation,” explained Jon Chesnut, Ph.D., the company’s senior director of synthetic biology R&D. “The formulation is specifically designed for delivering the RNP to cells more efficiently.”

    Besides the reagent and the formulation, Thermo Fisher has also developed a range of gene-editing tools. For example, it has introduced the Neon® transfection system for delivering DNA, RNA, or protein into cells via electroporation. Dr. Chesnut emphasized the company’s focus on simplifying complex workflows by optimizing protocols and pairing everything with the appropriate up- and downstream reagents.

From Mammalian Cells to Microbes

One of the first sources of CRISPR technology was the Feng Zhang laboratory at the Broad Institute, which counted among its first licensees a company called GenScript. This company offers a gene-editing service called GenCRISPR™ to establish mammalian cell lines with CRISPR-derived gene knockouts.

“There are a lot of challenges with mammalian cells, and each cell line has its own set of issues,” said Laura Geuss, a marketing specialist at GenScript. “We try to offer a variety of packages that can help customers who have difficult-to-work-with cells.” These packages include both viral-based and transient transfection techniques.

However, the most distinctive service offered by GenScript is its microbial genome-editing service for bacteria (Escherichia coli) and yeast (Saccharomyces cerevisiae). The company’s strategy for gene editing in bacteria can enable seamless knockins, knockouts, or gene replacements by combining CRISPR with lambda red recombineering. Traditionally one of the most effective methods for gene editing in microbes, recombineering allows editing without restriction enzymes through in vivo homologous recombination mediated by a phage-based recombination system such as lambda red.

On its own, lambda red technology cannot target multiple genes, but when paired with CRISPR, it allows the editing of multiple genes with greater efficiency than is possible with CRISPR alone, as the lambda red proteins help repair double-strand breaks in E. coli. The ability to knockout different gene combinations makes Genscript’s microbial editing service particularly well suited for the optimization of metabolic pathways.

Pooled and Arrayed Library Strategies

Scientists are using CRISPR technology for applications such as metabolic engineering and drug development. Yet another application area benefitting from CRISPR technology is cancer research. Here, the use of pooled CRISPR libraries is becoming commonplace. Pooled CRISPR libraries can help detect mutations that affect drug resistance, and they can aid in patient stratification and clinical trial design.

Pooled screening uses proliferation or viability as a phenotype to assess how genetic alterations, resulting from the application of a pooled CRISPR library, affect cell growth and death in the presence of a therapeutic compound. The enrichment or depletion of different gRNA populations is quantified using deep sequencing to identify the genomic edits that result in changes to cell viability.

MilliporeSigma provides pooled CRISPR libraries ranging from the whole human genome to smaller custom pools for these gene-function experiments. For pharmaceutical and biotech companies, Horizon Discovery offers a pooled screening service, ResponderSCREEN, which provides a whole-genome pooled screen to identify genes that confer sensitivity or resistance to a compound. This service is comprehensive, taking clients from experimental design all the way through to suggestions for follow-up studies.

Horizon Discovery maintains a Research Biotech business unit that is focused on target discovery and enabling translational medicine in oncology. “Our internal backbone gives us the ability to provide expert advice demonstrated by results,” said Jon Moore, Ph.D., the company’s CSO.

In contrast to a pooled screen, where thousands of gRNA are combined in one tube, an arrayed screen applies one gRNA per well, removing the need for deep sequencing and broadening the options for different endpoint assays. To establish and distribute a whole-genome arrayed lentiviral CRISPR library, MilliporeSigma partnered with the Wellcome Trust Sanger Institute. “This is the first and only arrayed CRISPR library in the world,” declared Shawn Shafer, Ph.D., functional genomics market segment manager, MilliporeSigma. “We were really proud to partner with Sanger on this.”

Pooled and arrayed screens are powerful tools for studying gene function. The appropriate platform for an experiment, however, will be determined by the desired endpoint assay.

Detection and Quantification of Edits

 

The QX200 Droplet Digital PCR System from Bio-Rad Laboratories can provide researchers with an absolute measure of target DNA molecules for EvaGreen or probe-based digital PCR applications. The system, which can provide rapid, low-cost, ultra-sensitive quantification of both NHEJ- and HDR-editing events, consists of two instruments, the QX200 Droplet Generator and the QX200 Droplet Reader, and their associated consumables.

Finally, one last challenge for CRISPR lies in the detection and quantification of changes made to the genome post-editing. Conventional methods for detecting these alterations include gel methods and next-generation sequencing. While gel methods lack sensitivity and scalability, next-generation sequencing is costly and requires intensive bioinformatics.

To address this gap, Bio-Rad Laboratories developed a set of assay strategies to enable sensitive and precise edit detection with its Droplet Digital PCR (ddPCR) technology. The platform is designed to enable absolute quantification of nucleic acids with high sensitivity, high precision, and short turnaround time through massive droplet partitioning of samples.

Using a validated assay, a typical ddPCR experiment takes about five to six hours to complete. The ddPCR platform enables detection of rare mutations, and publications have reported detection of precise edits at a frequency of <0.05%, and of NHEJ-derived indels at a frequency as low as 0.1%. In addition to quantifying precise edits, indels, and computationally predicted off-target mutations, ddPCR can also be used to characterize the consequences of edits at the RNA level.

According to a recently published Science paper, the laboratory of Charles A. Gersbach, Ph.D., at Duke University used ddPCR in a study of muscle function in a mouse model of Duchenne muscular dystrophy. Specifically, ddPCR was used to assess the efficiency of CRISPR-Cas9 in removing the mutated exon 23 from the dystrophin gene. (Exon 23 deletion by CRISPR-Cas9 resulted in expression of the modified dystrophin gene and significant enhancement of muscle force.)

Quantitative ddPCR showed that exon 23 was deleted in ~2% of all alleles from the whole-muscle lysate. Further ddPCR studies found that 59% of mRNA transcripts reflected the deletion.

“There’s an overarching idea that the genome-editing field is moving extremely quickly, and for good reason,” asserted Jennifer Berman, Ph.D., staff scientist, Bio-Rad Laboratories. “There’s a lot of exciting work to be done, but detection and quantification of edits can be a bottleneck for researchers.”

The gene-editing field is moving quickly, and new innovations are finding their way into the laboratory as researchers lay the foundation for precise, well-controlled gene editing with CRISPR.

 

Are Current Cancer Drug Discovery Methods Flawed?

GEN May 3, 2016   http://www.genengnews.com/gen-news-highlights/are-current-cancer-drug-discovery-methods-flawed/81252682/

 

Researchers utilized a systems biology approach to develop new methods to assess drug sensitivity in cells. [The Institute for Systems Biology]

Understanding how cells respond and proliferate in the presence of anticancer compounds has been the foundation of drug discovery ideology for decades. Now, a new study from scientists at Vanderbilt University casts significant suspicion on the primary method used to test compounds for anticancer activity in cells—instilling doubt on methods employed by the entire scientific enterprise and pharmaceutical industry to discover new cancer drugs.

“More than 90% of candidate cancer drugs fail in late-stage clinical trials, costing hundreds of millions of dollars,” explained co-senior author Vito Quaranta, M.D., director of the Quantitative Systems Biology Center at Vanderbilt. “The flawed in vitro drug discovery metric may not be the only responsible factor, but it may be worth pursuing an estimate of its impact.”

The Vanderbilt investigators have developed what they believe to be a new metric for evaluating a compound’s effect on cell proliferation—called the DIP (drug-induced proliferation) rate—that overcomes the flawed bias in the traditional method.

The findings from this study were published recently in Nature Methods in an article entitled “An Unbiased Metric of Antiproliferative Drug Effect In Vitro.”

For more than three decades, researchers have evaluated the ability of a compound to kill cells by adding the compound in vitro and counting how many cells are alive after 72 hours. Yet, proliferation assays that measure cell number at a single time point don’t take into account the bias introduced by exponential cell proliferation, even in the presence of the drug.

“Cells are not uniform, they all proliferate exponentially, but at different rates,” Dr. Quaranta noted. “At 72 hours, some cells will have doubled three times and others will not have doubled at all.”

Dr. Quaranta added that drugs don’t all behave the same way on every cell line—for example, a drug might have an immediate effect on one cell line and a delayed effect on another.

The research team decided to take a systems biology approach, a mixture of experimentation and mathematical modeling, to demonstrate the time-dependent bias in static proliferation assays and to develop the time-independent DIP rate metric.

“Systems biology is what really makes the difference here,” Dr. Quaranta remarked. “It’s about understanding cells—and life—as dynamic systems.”This new study is of particular importance in light of recent international efforts to generate data sets that include the responses of thousands of cell lines to hundreds of compounds. Using the

  • Cancer Cell Line Encyclopedia (CCLE) and
  • Genomics of Drug Sensitivity in Cancer (GDSC) databases

will allow drug discovery scientists to include drug response data along with genomic and proteomic data that detail each cell line’s molecular makeup.

“The idea is to look for statistical correlations—these particular cell lines with this particular makeup are sensitive to these types of compounds—to use these large databases as discovery tools for new therapeutic targets in cancer,” Dr. Quaranta stated. “If the metric by which you’ve evaluated the drug sensitivity of the cells is wrong, your statistical correlations are basically no good.”

The Vanderbilt team evaluated the responses from four different melanoma cell lines to the drug vemurafenib, currently used to treat melanoma, with the standard metric—used for the CCLE and GDSC databases—and with the DIP rate. In one cell line, they found a glaring disagreement between the two metrics.

“The static metric says that the cell line is very sensitive to vemurafenib. However, our analysis shows this is not the case,” said co-lead study author Leonard Harris, Ph.D., a systems biology postdoctoral fellow at Vanderbilt. “A brief period of drug sensitivity, quickly followed by rebound, fools the static metric, but not the DIP rate.”

Dr. Quaranta added that the findings “suggest we should expect melanoma tumors treated with this drug to come back, and that’s what has happened, puzzling investigators. DIP rate analyses may help solve this conundrum, leading to better treatment strategies.”

The researchers noted that using the DIP rate is possible because of advances in automation, robotics, microscopy, and image processing. Moreover, the DIP rate metric offers another advantage—it can reveal which drugs are truly cytotoxic (cell killing), rather than merely cytostatic (cell growth inhibiting). Although cytostatic drugs may initially have promising therapeutic effects, they may leave tumor cells alive that then have the potential to cause the cancer to recur.

The Vanderbilt team is currently in the process of identifying commercial entities that can further refine the software and make it widely available to the research community to inform drug discovery.

 

An unbiased metric of antiproliferative drug effect in vitro

Leonard A HarrisPeter L FrickShawn P GarbettKeisha N HardemanB Bishal PaudelCarlos F LopezVito Quaranta & Darren R Tyson
Nature Methods 2 May (2016)
                 doi:10.1038/nmeth.3852

In vitro cell proliferation assays are widely used in pharmacology, molecular biology, and drug discovery. Using theoretical modeling and experimentation, we show that current metrics of antiproliferative small molecule effect suffer from time-dependent bias, leading to inaccurate assessments of parameters such as drug potency and efficacy. We propose the drug-induced proliferation (DIP) rate, the slope of the line on a plot of cell population doublings versus time, as an alternative, time-independent metric.

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  3. Bonnans, C., Chou, J. & Werb, Z. Nat. Rev. Mol. Cell Biol. 15, 786801 (2014).
  4. Garnett, M.J. et al. Nature 483, 570575 (2012)

 

Mapping Traits to Genes with CRISPR

Researchers develop a technique to direct chromosome recombination with CRISPR/Cas9, allowing high-resolution genetic mapping of phenotypic traits in yeast.

By Catherine Offord | May 5, 2016

http://www.the-scientist.com/?articles.view/articleNo/46029/title/Mapping-Traits-to-Genes-with-CRISPR

 

Researchers used CRISPR/Cas9 to make a targeted double-strand break (DSB) in one arm of a yeast chromosome labeled with a green fluorescent protein (GFP) gene. A within-cell mechanism called homologous repair (HR) mends the broken arm using its homolog, resulting in a recombined region from the site of the break to the chromosome tip. When this cell divides by mitosis, each daughter cell will contain a homozygous section in an outcome known as “loss of heterozygosity” (LOH). One of the daughter cells is detectable because, due to complete loss of the GFP gene, it will no longer be fluorescent.REPRINTED WITH PERMISSION FROM M.J. SADHU ET AL., SCIENCE

When mapping phenotypic traits to specific loci, scientists typically rely on the natural recombination of chromosomes during meiotic cell division in order to infer the positions of responsible genes. But recombination events vary with species and chromosome region, giving researchers little control over which areas of the genome are shuffled. Now, a team at the University of California, Los Angeles (UCLA), has found a way around these problems by using CRISPR/Cas9 to direct targeted recombination events during mitotic cell division in yeast. The team described its technique today (May 5) in Science.

“Current methods rely on events that happen naturally during meiosis,” explained study coauthor Leonid Kruglyak of UCLA. “Whatever rate those events occur at, you’re kind of stuck with. Our idea was that using CRISPR, we can generate those events at will, exactly where we want them, in large numbers, and in a way that’s easy for us to pull out the cells in which they happened.”

Generally, researchers use coinheritance of a trait of interest with specific genetic markers—whose positions are known—to figure out what part of the genome is responsible for a given phenotype. But the procedure often requires impractically large numbers of progeny or generations to observe the few cases in which coinheritance happens to be disrupted informatively. What’s more, the resolution of mapping is limited by the length of the smallest sequence shuffled by recombination—and that sequence could include several genes or gene variants.

“Once you get down to that minimal region, you’re done,” said Kruglyak. “You need to switch to other methods to test every gene and every variant in that region, and that can be anywhere from challenging to impossible.”

But programmable, DNA-cutting champion CRISPR/Cas9 offered an alternative. During mitotic—rather than meiotic—cell division, rare, double-strand breaks in one arm of a chromosome preparing to split are sometimes repaired by a mechanism called homologous recombination. This mechanism uses the other chromosome in the homologous pair to replace the sequence from the break down to the end of the broken arm. Normally, such mitotic recombination happens so rarely as to be impractical for mapping purposes. With CRISPR/Cas9, however, the researchers found that they could direct double-strand breaks to any locus along a chromosome of interest (provided it was heterozygous—to ensure that only one of the chromosomes would be cut), thus controlling the sites of recombination.

Combining this technique with a signal of recombination success, such as a green fluorescent protein (GFP) gene at the tip of one chromosome in the pair, allowed the researchers to pick out cells in which recombination had occurred: if the technique failed, both daughter cells produced by mitotic division would be heterozygous, with one copy of the signal gene each. But if it succeeded, one cell would end up with two copies, and the other cell with none—an outcome called loss of heterozygosity.

“If we get loss of heterozygosity . . . half the cells derived after that loss of heterozygosity event won’t have GFP anymore,” study coauthor Meru Sadhu of UCLA explained. “We search for these cells that don’t have GFP out of the general population of cells.” If these non-fluorescent cells with loss of heterozygosity have the same phenotype as the parent for a trait of interest, then CRISPR/Cas9-targeted recombination missed the responsible gene. If the phenotype is affected, however, then the trait must be linked to a locus in the recombined, now-homozygous region, somewhere between the cut site and the GFP gene.

By systematically making cuts using CRISPR/Cas9 along chromosomes in a hybrid, diploid strain ofSaccharomyces cerevisiae yeast, picking out non-fluorescent cells, and then observing the phenotype, the UCLA team demonstrated that it could rapidly identify the phenotypic contribution of specific gene variants. “We can simply walk along the chromosome and at every [variant] position we can ask, does it matter for the trait we’re studying?” explained Kruglyak.

For example, the team showed that manganese sensitivity—a well-defined phenotypic trait in lab yeast—could be pinpointed using this method to a single nucleotide polymorphism (SNP) in a gene encoding the Pmr1 protein (a manganese transporter).

Jason Moffat, a molecular geneticist at the University of Toronto who was not involved in the work, toldThe Scientist that researchers had “dreamed about” exploiting these sorts of mechanisms for mapping purposes, but without CRISPR, such techniques were previously out of reach. Until now, “it hasn’t been so easy to actually make double-stranded breaks on one copy of a pair of chromosomes, and then follow loss of heterozygosity in mitosis,” he said, adding that he hopes to see the approach translated into human cell lines.

Applying the technique beyond yeast will be important, agreed cell and developmental biologist Ethan Bier of the University of California, San Diego, because chromosomal repair varies among organisms. “In yeast, they absolutely demonstrate the power of [this method],” he said. “We’ll just have to see how the technology develops in other systems that are going to be far less suited to the technology than yeast. . . . I would like to see it implemented in another system to show that they can get the same oomph out of it in, say, mammalian somatic cells.”

Kruglyak told The Scientist that work in higher organisms, though planned, is still in early stages; currently, his team is working to apply the technique to map loci responsible for trait differences between—rather than within—yeast species.

“We have a much poorer understanding of the differences across species,” Sadhu explained. “Except for a few specific examples, we’re pretty much in the dark there.”

M.J. Sadhu, “CRISPR-directed mitotic recombination enables genetic mapping without crosses,” Science, doi:10.1126/science.aaf5124, 2016.

 

CRISPR-directed mitotic recombination enables genetic mapping without crosses

Meru J Sadhu, Joshua S Bloom, Laura Day, Leonid Kruglyak

Thank you, David, for the kind words and comments. We agree that the most immediate applications of the CRISPR-based recombination mapping will be in unicellular organisms and cell culture. We also think the method holds a lot of promise for research in multicellular organisms, although we did not mean to imply that it “will be an efficient mapping method for all multicellular organisms”. Every organism will have its own set of constraints as well as experimental tools that will be relevant when adapting a new technique. To best help experts working on these organisms, here are our thoughts on your questions.

You asked about mutagenesis during recombination. We Sanger sequenced 72 of our LOH lines at the recombination site and did not observe any mutations, as described in the supplementary materials. We expect the absence of mutagenesis is because we targeted heterozygous sites where the untargeted allele did not have a usable PAM site; thus, following LOH, the targeted site is no longer present and cutting stops. In your experiments you targeted sites that were homozygous; thus, following recombination, the CRISPR target site persisted, and continued cutting ultimately led to repair by NHEJ and mutagenesis.

As to the more general question of the optimal mapping strategies in different organisms, they will depend on the ease of generating and screening for editing events, the cost and logistics of maintaining and typing many lines, and generation time, among other factors. It sounds like in Drosophila today, your related approach of generating markers with CRISPR, and then enriching for natural recombination events that separate them, is preferable. In yeast, we’ve found the opposite to be the case. As you note, even in Drosophila, our approach may be preferable for regions with low or highly non-uniform recombination rates.

Finally, mapping in sterile interspecies hybrids should be straightforward for unicellular hybrids (of which there are many examples) and for cells cultured from hybrid animals or plants. For studies in hybrid multicellular organisms, we agree that driving mitotic recombination in the early embryo may be the most promising approach. Chimeric individuals with mitotic clones will be sufficient for many traits. Depending on the system, it may in fact be possible to generate diploid individuals with uniform LOH genotype, but this is certainly beyond the scope of our paper. The calculation of the number of lines assumes that the mapping is done in a single step; as you note in your earlier comment, mapping sequentially can reduce this number dramatically.

This is a lovely method and should find wide applicability in many settings, especially for microorganisms and cell lines. However, it is not clear that this approach will be, as implied by the discussion, an efficient mapping method for all multicellular organisms. I have performed similar experiments in Drosophila, focused on meiotic recombination, on a much smaller scale, and found that CRISPR-Cas9 can indeed generate targeted recombination at gRNA target sites. In every case I tested, I found that the recombination event was associated with a deletion at the gRNA site, which is probably unimportant for most mapping efforts, but may be a concern in some specific cases, for example for clinical applications. It would be interesting to know how often mutations occurred at the targeted gRNA site in this study.

The wider issue, however, is whether CRISPR-mediated recombination will be more efficient than other methods of mapping. After careful consideration of all the costs and the time involved in each of the steps for Drosophila, we have decided that targeted meiotic recombination using flanking visible markers will be, in most cases, considerably more efficient than CRISPR-mediated recombination. This is mainly due to the large expense of injecting embryos and the extensive effort and time required to screen injected animals for appropriate events. It is both cheaper and faster to generate markers (with CRISPR) and then perform a large meiotic recombination mapping experiment than it would be to generate the lines required for CRISPR-mediated recombination mapping. It is possible to dramatically reduce costs by, for example, mapping sequentially at finer resolution. But this approach would require much more time than marker-assisted mapping. If someone develops a rapid and cheap method of reliably introducing DNA into Drosophila embryos, then this calculus might change.

However, it is possible to imagine situations where CRISPR-mediated mapping would be preferable, even for Drosophila. For example, some genomic regions display extremely low or highly non-uniform recombination rates. It is possible that CRISPR-mediated mapping could provide a reasonable approach to fine mapping genes in these regions.

The authors also propose the exciting possibility that CRISPR-mediated loss of heterozygosity could be used to map traits in sterile species hybrids. It is not entirely obvious to me how this experiment would proceed and I hope the authors can illuminate me. If we imagine driving a recombination event in the early embryo (with maternal Cas9 from one parent and gRNA from a second parent), then at best we would end up with chimeric individuals carrying mitotic clones. I don’t think one could generate diploid animals where all cells carried the same loss of heterozygosity event. Even if we could, this experiment would require construction of a substantial number of stable transgenic lines expressing gRNAs. Mapping an ~20Mbp chromosome arm to ~10kb would require on the order of two-thousand transgenic lines. Not an undertaking to be taken lightly. It is already possible to perform similar tests (hemizygosity tests) using D. melanogaster deficiency lines in crosses with D. simulans, so perhaps CRISPR-mediated LOH could complement these deficiency screens for fine mapping efforts. But, at the moment, it is not clear to me how to do the experiment.

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CRISPR-Cas9 Screening by Horizon Discovery, Cambridge, UK – HDx™ Reference Standards

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https://www.horizondiscovery.com/resources/webinars/how-crispr-cas9-screening-will-revolutionise-your-drug-development-programs?_cldee=ZC5vcmNoYXJkLXdlYmJAYmF0aC5lZHU%3d&utm_source=ClickDimensions&utm_medium=email&utm_campaign=201605%20Email%20-%207%20must-read%20CRISPR%20screening%20papers&urlid=3

Seven essential CRISPR screening papers

https://www.horizondiscovery.com/knowledge-base/essential-crispr-screening-papers?_cldee=ZC5vcmNoYXJkLXdlYmJAYmF0aC5lZHU%3d&utm_source=ClickDimensions&utm_medium=email&utm_campaign=201605%20Email%20-%207%20must-read%20CRISPR%20screening%20papers&urlid=1

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Watch our short introductory video to Horizon’s CRISPR screening platform CLICK HERE
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https://www.horizondiscovery.com/knowledge-base/essential-crispr-screening-papers?_cldee=ZC5vcmNoYXJkLXdlYmJAYmF0aC5lZHU%3d&utm_source=ClickDimensions&utm_medium=email&utm_campaign=201605%20Email%20-%207%20must-read%20CRISPR%20screening%20papers&urlid=1

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Scientific Literature

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