Posts Tagged ‘CRISPR’

Reporter and Curator: Dr. Sudipta Saha, Ph.D.


Scientists think excessive population growth is a cause of scarcity and environmental degradation. A male pill could reduce the number of unintended pregnancies, which accounts for 40 percent of all pregnancies worldwide.


But, big drug companies long ago dropped out of the search for a male contraceptive pill which is able to chemically intercept millions of sperm before they reach a woman’s egg. Right now the chemical burden for contraception relies solely on the female. There’s not much activity in the male contraception field because an effective solution is available on the female side.


Presently, male contraception means a condom or a vasectomy. But researchers from Center for Drug Discovery at Baylor College of Medicine, USA are renewing the search for a better option—an easy-to-take pill that’s safe, fast-acting, and reversible.


The scientists began with lists of genes active in the testes for sperm production and motility and then created knockout mice that lack those genes. Using the gene-editing technology called CRISPR, in collaboration with Japanese scientists, they have so far made more than 75 of these “knockout” mice.


They allowed these mice to mate with normal (wild type) female mice, and if their female partners don’t get pregnant after three to six months, it means the gene might be a target for a contraceptive. Out of 2300 genes that are particularly active in the testes of mice, the researchers have identified 30 genes whose deletion makes the male infertile. Next the scientists are planning a novel screening approach to test whether any of about two billion chemicals can disable these genes in a test tube. Promising chemicals could then be fed to male mice to see if they cause infertility.


Female birth control pills use hormones to inhibit a woman’s ovaries from releasing eggs. But hormones have side effects like weight gain, mood changes, and headaches. A trial of one male contraceptive hormone was stopped early in 2011 after one participant committed suicide and others reported depression. Moreover, some drug candidates have made animals permanently sterile which is not the goal of the research. The challenge is to prevent sperm being made without permanently sterilizing the individual.


As a better way to test drugs, Scientists at University of Georgia, USA are investigating yet another high-tech approach. They are turning human skin cells into stem cells that look and act like the spermatogonial cells in the testes. Testing drugs on such cells might provide more accurate leads than tests on mice.


The male pill would also have to start working quickly, a lot sooner than the female pill, which takes about a week to function. Scientists from University of Dundee, U.K. admitted that there are lots of challenges. Because, a women’s ovary usually release one mature egg each month, while a man makes millions of sperm every day. So, the male pill has to be made 100 percent effective and act instantaneously.




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LIVE 9/21 8AM to 10:55 AM Expoloring the Versatility of CRISPR/Cas9 at CHI’s 14th Discovery On Target, 9/19 – 9/22/2016, Westin Boston Waterfront, Boston

Leaders in Pharmaceutical Business Intelligence (LPBI) Group is a

Media Partner of CHI for CHI’s 14th Annual Discovery on Targettaking place September 19 – 22, 2016 in Boston.

In Attendance, streaming LIVE using Social Media

Aviva Lev-Ari, PhD, RN





COMMENTS BY Stephen J Williams, PhD



8:00 Chairperson’s Opening Remarks

TJ Cradick , Ph.D., Head of Genome Editing, CRISPR Therapeutics




8:10 Functional Genomics Using CRISPR-Cas9: Technology and Applications

Neville Sanjana, Ph.D., Core Faculty Member, New York Genome Center and Assistant Professor, Department of Biology & Center for Genomics and Systems Biology, New York University


CRISPR Cas9 is easier to target to multiple genomic loci; RNA specifies DNA targeting; with zinc finger nucleases or TALEEN in the protein specifies DNA targeting


  • This feature of crisper allows you to make a quick big and cheap array of a GENOME SCALE Crisper Knock out (GeCKO) screening library
  • How do you scale up the sgRNA for whole genome?; for all genes in RefSeq, identify consitutive exons using RNA-sequencing data from 16 primary human tissue (alot of genes end with ‘gg’) changing the bases on 3’ side negates crisper system but changing on 5’ then crisper works fine
  • Rank sequences to be specific for target
  • Cloned array into lentiviral and put in selectable markers
  • GeCKO displays high consistency betweens reagents for the same gene versus siRNA; GeCKO has high screening sensitivity
  • 98% of genome is noncoding so what about making a library for intronic regions (miRNA, promoter regions?)
  • So you design the sgRNA library by taking 100kb of gene-adjacent regions
  • They looked at CUL3; (data will soon be published in Science)
  • Do a transcription CHIP to verify the lack of binding of transcription factor of interest
  • Can also target histone marks on promoter and enhancer elements
  • NYU wants to explore this noncoding screens




8:40 Therapeutic Gene Editing With CRISPR/Cas9

TJ Cradick , Ph.D., Head of Genome Editing, CRISPR Therapeutics


NEHJ is down and dirty repair of single nonhomologous end but when have two breaks the NEHJ repair can introduce the inversions or deletions


    • High-throughput screens are fine but can limit your view of genomic context; genome searches pick unique sites so use bioinformatic programs  to design specific guide Rna
    • Bioinformatic directed, genome wide, functional screens
    • Compared COSMID and CCTOP; 320 COSMID off-target sites, 333 CCtop off target
    • Young lab GUIDESeq program genome wide assay useful to design guides
    • If shorten guide may improve specificity; also sometime better sensitivity if lengthen guide


  • Manufacturing of autologous gene corrected product ex vivo gene correction (Vertex, Bayer, are partners in this)



They need to use a clones from multiple microarrays before using the GUidESeq but GUIDEseq is better for REMOVING the off targets than actually producing the sgRNA library you want (seems the methods for library development are not fully advanced to do this)


The score sometimes for the sgRNA design programs do not always give the best result because some sgRNAs are genome context dependent

9:10 Towards Combinatorial Drug Discovery: Mining Heterogeneous Phenotypes from Large Scale RNAi/Drug Perturbations

Arvind Rao, Ph.D., Assistant Professor, Department of Bioinformatics and Computational Biology, The University of Texas MD Anderson Cancer Center


Bioinformatics in CRISPR screens:  they looked at image analysis of light microscopy of breast cancer cells and looked for phenotypic changes


  • Then they modeled in a small pilot and then used the algorithm for 20,000 images (made morphometric measurements)
  • Can formulate training statistical algorithms to make a decision tree how you classify data points
  • Although their algorithms worked well there was also human input from scientists

Aggregate ranking of hits programs available on web like LINKS




10:25 CRISPR in Stem Cell Models of Eye Disease

Alexander Bassuk, M.D., Ph.D., Associate Professor of Pediatrics, Department of Molecular and Cellular Biology, University of Iowa


Blind athlete Michael Stone, biathlete, had eye disease since teenager helped fund and start the clinical trial for Starbardt disease; had one bad copy of ABCA4, heterozygous (inheritable in Ahkenazi Jewish) – a recessive inheritable mutation with juvenile macular degeneration

  • Also had another male in family with disease but he had another mutation in the RPGR gene
  • December 2015 paper Precision Medicine: Genetic Repair of retinitis pigmentosa in patient derived stem cells
  • They were able to correct the iPSCs in the RPGR gene derived from patient however low efficiency of repair, scarless repair, leaves changes in DNA, need clinical grade iPSCs, and need a humanized model of RPGR


10:55 CRISPR in Mouse Models of Eye Disease

Vinit Mahajan, M.D., Ph.D., Assistant Professor of Ophthalmology and Visual Sciences, University of Iowa College of Medicine

  • degeneration of the retina will see brown spots, the macula will often be preserved but retinal cells damaged but with RPGR have problems with peripheral vision, retinitis pigmentosa get tunnel vision with no peripheral vision (a mouse model of PDE6 Knockout recapitulates this phenotype)
  • the PDE6 is linked to the rhodopsin GTP pathway
  • rd1 -/- mouse has something that looks like retinal pigmentosa; has mutant PDE6; is actually a nonsense mutation in rd1 so they tried a crisper to fix in mice
  • with crisper fix of rd1 nonsense mutation the optic nerve looked comparible to normal and the retina structure restored
  • photoreceptors layers- some recovery but not complete
  • sequence results show the DNA is a mosaic so not correcting 100% but only 35% but stil leads to a phenotypic recovery; NHEJ was about 12% to 25% with large deletions
  • histology is restored in crspr repaired mice
  • CRSPR off target effects: WGS and analyze for variants SNV/indels, also looked at on target and off target regions; there were no off target SNVs indels while variants that did not pass quality control screening not a single SNV
  • Rhodopsin mutation accounts for a large % of patients (RhoD190N)
  • injection of gene therapy vectors: AAV vector carrying CRSPR and cas9 repair templates

CAPN mouse models

  • family in Iowa have dominant mutation in CAPN5; retinal degenerates
  • used CRSPR to generate mouse model with mutation in CAPN5 similar to family mutation
  • compared to other transgenic methods CRSPR is faster to produce a mouse model


Meeting #: #BostonDOT16

Meeting @: @BostonDOT


Overall good meeting #s:












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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

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 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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Best in Precision Medicine: RNA May Surpass DNA in Precision Medicine

Curator: Larry H. Bernstein, MD, FCAP


  • “A more complicated application of CRISPR technology is to use it for gene activation,” adds Dr. Tedesco. “Cellecta plans to optimize this application to bring forth highly efficient, inexpensive, high-throughput genetic screens based on their pooled libraries.


RNA May Surpass DNA in Precision Medicine

Scientists based at the Translational Genomics Research Institute have published a review heralding the promise of RNA sequencing (RNA-seq) for precision medicine. The scientists also note that progress will be needed on analytical, bioinformatics, and regulatory fronts, particularly in light of the transcriptome’s variety, dynamism, and wealth of detail. In this image, one aspect of RNA-seq is shown, the alignment with intron-split short reads. It reflects the alignment of mRNA sequence obtained via high-throughput sequencing and the expected behavior of the alignment to the reference genome when the read falls in an exon–exon junction. [Rgocs, Wikipedia]

It’s not an either/or situation. Both DNA sequencing and RNA sequencing hold clinical promise—diagnostically, prognostically, and therapeutically. It must be said, however, that RNA sequencing reflects the dynamic nature of gene expression, shifting with the vagaries of health and disease. Also, RNA sequencing captures more biochemical complexity, in the sense that it allows for the detection of a wide variety of RNA species, including mRNA, noncoding RNA, pathogen RNA, chimeric gene fusions, transcript isoforms, and splice variants, and provides the capability to quantify known, predefined RNA species and rare RNA transcript variants within a sample.

All these potential advantages were cited in a paper that appeared March 21 in Nature Reviews Genetics, in an article entitled, “Translating RNA Sequencing into Clinical Diagnostics: Opportunities and Challenges.” The paper, contributed by scientists based at the Translational Genomics Research Institute (TGen), was definitely optimistic about the clinical utility of RNA sequencing, but it also highlighted the advances that would have to occur if RNA sequencing is to achieve its promise.

In general, the very things that make RNA sequencing so interesting are the same things that make it so challenging. RNA sequencing would take the measure of a world—the transcriptome—that is incredibly rich. To capture all the relevant subtleties of the transcriptome, scientists will have to develop sensitive, precise, and trustworthy analytical techniques. What’s more, scientists will need to find efficient and reliable means of processing and interpreting all of the transcriptome data they will collect. Finally, they will need to continue integrating RNA-based knowledge with DNA-based knowledge. That is, RNA sequencing results can be used to guide the interpretation of DNA sequencing results.

In their Nature Reviews Genetics paper, the TGen scientists review the state of RNA sequencing and offer specific recommendations to enhance its clinical utility. The TGen scientists make a special point about the promise held by extracellular RNA (exRNA). Because exRNA can be monitored by simply taking a blood sample, as opposed to taking a tumor biopsy, it could serve as a noninvasive diagnostic indicator of disease.

“Detection of gene fusions and differential expression of known disease-causing transcripts by RNA-seq represent some of the most immediate opportunities,” wrote the authors. “However, it is the diversity of RNA species detected through RNA-seq that holds new promise for the multi-faceted clinical applicability of RNA-based measures, including the potential of extracellular RNAs as non-invasive diagnostic indicators of disease.”

The first test measuring exRNA was released earlier this year, the paper said, for use measuring specific exRNAs in lung cancer patients. And, the potential for using RNA-seq in cancer is expanding rapidly. Commercial RNA-seq tests are now available, and they provide the opportunity for clinicians to profile cancer more comprehensively and use this information to guide treatment selection for their patients.

In addition, the authors reported on several recent applications for RNA-seq in the diagnosis and management of infectious diseases, such as monitoring for drug-resistant populations during therapy and tracking the origin and spread of the Ebola virus.

Despite these advances, the authors also sounded a few cautionary notes. “There are currently few agreed upon methods for isolation or quantitative measurements and a current lack of quality controls that can be used to test platform accuracy and sample preparation quality,” they wrote. “Analytical, bioinformatics, and regulatory challenges exist, and ongoing efforts toward the establishment of benchmark standards, assay optimization for clinical conditions and demonstration of assay reproducibility are required to expand the clinical utility of RNA-seq.”

Overall, the authors remain hopeful that precision medicine will embrace RNA sequencing. For example, lead author Sara Byron, research assistant professor in TGen’s Center for Translational Innovation, said, “RNA is a dynamic and diverse biomolecule with an essential role in numerous biological processes. From a molecular diagnostic standpoint, RNA-based measurements have the potential for broad application across diverse areas of human health, including disease diagnosis, prognosis, and therapeutic selection.”


Gene Editing Casts a Wide Net   

With CRISPR, Gene Editing Can Trawl the Murk, Catching Elusive Phenotypes amidst the Epigenetic Ebb and Flow

Gene-editing advances will not only open new avenues toward curing genetic diseases but will also rapidly increase the pace of new scientific discoveries about human and other types of genomes. [iStock/adventtr]


  • Genome editing, a much-desired means of accomplishing gene knockout, gene activation, and other tasks, once seemed just beyond the reach of most research scientists and drug developers. But that was before the advent of CRISPR technology, an easy, versatile, and dependable means of implementing genetic modifications. It is in the process of democratizing genome editing.

    CRISPR stands for “clustered, regularly interspaced, short palindromic repeats,” segments of DNA that occur naturally in many types of bacteria. These segments function as part of an ancient immune system. Each segment precedes “spacer DNA,” a short base sequence that is derived from a fragment of foreign DNA. Spacers serve as reminders of past encounters with phages or plasmids.

    The CRISPR-based immune system encompasses several mechanisms, including one in which CRISPR loci are transcribed into small RNAs that may complex with a nuclease called CRISPR-associated protein (Cas). Then the RNA guides Cas, which cleaves invading DNA on the basis of sequence complementarity.

    In the laboratory, CRISPR sequences are combined with a short RNA complementary to a target gene site. The result is a complex in which the RNA guides Cas to a preselected target.

    Cas produces precise site-specific DNA breaks, which, with imperfect repair, cause gene mutagenesis. In more recent applications, Cas can serve as an anchor for other proteins, such as transcriptional factors and epigenetic enzymes. This system, it seems, has almost limitless versatility.

  • Edited Stem Cells

    The Sanger Institute Mouse Genetic Program, along with other academic institutions around the world, provides access to thousands of genetically modified mouse strains. “Genetic engineering of mouse embryonic stem (ES) cells by homologous recombination is a powerful technique that has been around since the 1980s,” says William Skarnes, Ph.D., senior group leader at the Wellcome Trust Sanger Institute.

    “A significant drawback of the ES technology is the time required to achieve a germline transmission of the modified genetic locus,” he continues. “While we have an exhaustive collection of modified ES cells, only about 5,000 knockout mice, or a quarter of mouse genome, were derived on the basis of this methodology.”

    The dominant position of the mouse ES cell engineering is now effectively challenged by the CRISPR technology. Compared with very low rates of homologous recombination in fertilized eggs, CRISPR generates high levels of mutations, and off-target effects may be so few as to be undetectable.

    “We used the whole-genome sequencing to thoroughly assess off-target mutations in the offspring of CRISPR-engineered founder animals,” informs Dr. Skarnes. “A mutated Cas9 nuclease was deployed to increase specificity, resulting in nearly perfect targeting.”

    Dr. Skarnes explains that the major mouse genome centers are now switching to CRISPR to complete the creation of the world-wide repository of mouse knockouts. His own research is now focused on genetically engineered induced pluripotent stem cells (iPSCs). These cells are adult cells that have been reprogrammed to an embryonic stem cell–like state, and are thus devoid of ethical issues associated with research on human embryonic stem cells. The ultimate goal is to establish a world-wide panel of reference iPSCs created by high-throughput genetic editing of every single human gene.

    “We are poised to begin a large-scale phenotypic analysis of human genes,” declares Dr. Skarnes. His lab is releasing the first set of functional data on 100 DNA repair genes. “By knocking out individual proteins involved in DNA repair and sequencing the genomes of mutant cells,” declares Dr. Skarnes, “we hope to better understand the mutational signatures that occur in cancer.”

  • Pooled CRISPR Libraries

    Researchers hope to gain a better understanding of the mutational signatures found in cancers by using CRISPR techniques to knock out individual proteins involved in DNA repair and then sequencing the genomes of mutant cells. [iStock/zmeel]

    Connecting a phenotype to the underlying genomics requires an unbiased screening of multiple genes at once. “Pooled CRISPR libraries provide an opportunity to cast a wide net at a reasonably low cost,” says Donato Tedesco, Ph.D., lead research scientist at Cellecta. “Screening one gene at a time on genome scale is a significant investment of time and money that not everyone can afford, especially when looking for common genetic drivers across many cell models.”

    Building on years of experience with shRNA libraries, Cellecta is uniquely positioned to prepare pooled CRISPR libraries for genome-wide or targeted screens of gene families. While shRNA interferes with gene translation, CRISPR disrupts a gene and the genomic level due to imperfections in the DNA repair mechanism.

    To determine if these different mechanisms for inactivating genes affect the results of genetic screens, the team conducted a side-by-side comparison of Cellecta’s Human Genome-Wide Module 1 shRNA Library, which expresses 50,000 shRNA targeting 6,300 human genes, with the library of 50,000 gRNA targeting the same gene set. The concordance between approaches was very high, suggesting that these technologies may be complementary and used for cross-confirmation of results.

    Also, a recently completed Phase I NIH SBIR Grant was used to create and test guiding strand RNA (sgRNA) structures to drastically improve efficiency of gene targeting. For this work, Cellecta used a pool library strategy to simultaneously test multiple sgRNA structures for their efficiency and specificity. An early customized Cellecta pooled gRNA library was successfully utilized for screening for epigenetic genes. This particular screen is highly dependent on a complete loss of function, and could not have been accomplished by shRNA inhibition.

    Scientists from Epizyme interrogated 600 genes in a panel of 100 cell lines and, in addition to finding many epigenetic genes required for proliferation in nearly all cell lines, were able to identify validate several essential epigenetic genes required only in subsets of cells with specific genetic lesions. In other words, pooled cell line screening was able to distinguish targets that are likely to produce toxic side effects in certain types of cancer cells from gene targets that are essential in most cells.

    “A more complicated application of CRISPR technology is to use it for gene activation,” adds Dr. Tedesco. “Cellecta plans to optimize this application to bring forth highly efficient, inexpensive, high-throughput genetic screens based on their pooled libraries.

  • Chemically Modified sgRNA

    Scientists based at Agilent Research Laboratories and Stanford University worked together to demonstrate that chemically modified single guide RNA can be used to enhance the genome editing of primary hepatopoietic stem cells and T cells. This image, which is from the Stanford laboratory of Matthew Porteus, M.D., Ph.D., shows CD34+ human hematopoietic stem cells that were edited to turn green. Editing involved inserting a construct for green fluorescent protein. About 1,000 cells are pictured here.

    Researchers at Agilent Technologies applied their considerable experience in DNA and RNA synthesis to develop a novel chemical synthesis method that can generate long RNAs of 100 nucleotides or more, such as single guide RNAs (sgRNAs) for CRISPR genome editing. “We have used this capability to design and test numerous chemical modifications at different positions of the RNA molecule,” said Laurakay Bruhn, Ph.D., section manager, biological chemistry, Agilent.

    Agilent Research Laboratories worked closely with the laboratory of Matthew Porteus, M.D., Ph.D., an associate professor of pediatrics and stem cell transplantation at Stanford University. The Agilent and Stanford researchers collaborated to further explore the benefits of chemically modified sgRNAs in genome editing of primary hematopoetic stem cells and T cells.

    Dr. Porteus’ lab chose three key target genes implicated in the development of severe combined immunodeficiency (SCID), sickle cell anemia, and HIV transmission. Editing these genes in the patient-derived cells offers an opportunity for novel precision therapies, as the edited cells can renew, expand, and colonize the donor’s bone marrow.

    Dr. Bruhn emphasized the importance of editing specificity, so that no other cellular function is affected by the change. The collaborators focused on three chemical modifications strategically placed at each end of sgRNAs that Agilent had previously tested to show they maintained sgRNA function. A number of other optimization strategies in cell culturing and transfection were explored to ensure high editing yields.

    “Primary cells are difficult to manipulate and edit in comparison with cell lines already adapted to cell culture,” maintains Dr. Bruhn. Widely varied cellular properties of primary cells may require experimental adaptation of editing techniques for each primary cell type.

    The resulting data showed that chemical modifications can greatly enhance efficiency of gene editing. The next step would translate these findings into animal models. Another advantage of chemical synthesis of RNA is that it can potentially be used to make large enough quantities for therapeutics.

    “We are working with Agilent’s Nucleic Acid Solution Division—a business focused on GMP manufacturing of oligonucleotides for therapeutics—to engage with customers interested in this capability and better understand how we might be able to help them accomplish their goals,” says Dr. Bruhn.

  • Customized Animal Models

    “Given their gene-knockout capabilities, zinc-finger-based technologies and CRISPR-based technologies opened the doors for creation of animal models that more closely resemble human disease than mouse models,” says Myung Shin, Ph.D., senior principal scientist, Merck & Co. Dr. Shin’s team supports Merck’s drug discovery and development program by creating animal models mimicking human genetics.

    For example, Dr. Shin’s team has worked with the Dahl salt-sensitive strain of rats, a widely studied model of hypertension. “We used zinc-finger nucleases to generate a homozygous knockout of a renal outer medullary potassium channel (ROMK) gene,” elaborates Dr. Shin. “The resulting model represents a major advance in elucidating the role of ROMK gene.”

    According to Dr. Shin, the model may also provide a bridge between genetics and physiology, particularly in studies of renal regulation and blood pressure. In one study, the model generated animal data that suggest ROMK plays a key role in kidney development and sodium absorption. Work along these lines may lead to a pharmacological strategy to manage hypertension.

    In another study, the team applied zinc-finger nuclease strategy to knockout the coagulation Factor XII, and thoroughly characterize them in thrombosis and hemostasis studies. Results confirmed and extended previous literature findings suggesting Factor XII as a potential target for antithrombotic therapies that carry minimal bleeding risk. The model can be further utilized to study safety profiles and off-target effects of such novel Factor XII inhibitors.

    “We use one-cell embryos to conduct genome editing with zinc-fingers and CRISPR,” continues Dr. Shin. “The ease of this genetic manipulation speeds up generation of animal models for testing of various hypotheses.”

    A zinc finger–generated knockout of the multidrug resistance protein MDR 1a P-glycoprotein became an invaluable tool for evaluating drug candidates for targets located in the central nervous system. For example, it demonstrated utility in pharmacological analyses.

    Dr. Shin’s future research is directed toward preclinical animal models that would contain specific nucleotide changes corresponding to those of humans. “CRISPR technology,” insists Dr. Shin, “brings an unprecedented power to manipulate genome at the level of a single nucleotide, to create gain- or loss-of-function genetic alterations, and to deeply understand the biology of a disease.”

  • Transcriptionally Active dCas9

    “Epigenome editing is important for several reasons,” says Charles Gersbach, Ph.D., an associate professor of biomedical engineering at Duke University. “It is a tool that helps us answer fundamental questions about biology. It advances disease modeling and drug screening. And it may, in the future, serve as mode of genetic therapy.”

    “One part of our research focuses on studying the function of epigenetic marks,” Dr. Gersback continues. “While many of these marks are catalogued, and some have been associated with the certain gene-expression states, the exact causal link between these marks and their effect on gene expression is not known. CRISPR technology can potentially allow for targeted direct manipulation of each epigenetic mark, one at a time.”

    Dr. Gersback’s team mutated the Cas9 nuclease to create deactivated Cas9 (dCas9), which is devoid of endonuclease activity. Although the dCas9 protein lacks catalytic activity, it may still serve as an anchor for a plethora of other important proteins, such as transcription factors and methyltransferases.

    In an elegant study, Dr. Gersbach and colleagues demonstrated that recruitment of a histone acetyltransferase by dCas9 to a genomic site activates nearby gene expression. Moreover, the activation occurred even when the acetyltransferase domain was targeted to a distal enhancer. Similarly, recruitment of KRAB repressor to a distant site silenced the target gene in a very specific manner. These findings support the important role of three-dimensional chromatin structures in gene activation.

    “Genome regulation by epigenetic markers is not static,” maintains Dr. Gersbach. “It responds to changes in the environment and other stimuli. It also changes during cell differentiation. We designed an inducible system providing us with an ability to execute dynamic control over the target genes.”

    In a light-activated CRISPR-Cas9 effector (LACE) system, blue light may be used to control the recruitment of the transcriptional factor VP64 to target DNA sequences. The system has been used to provide robust activation of four target genes with only minimal background activity. Selective illumination of culture plates created a pattern of gene expression in a population of cells, which could be used to mimic what is observed in natural tissues.

    Together with collaborators at Duke University, Dr. Gersbach intends to carry out the high-throughput analysis of all currently identified regulatory elements in the genome. “Our ultimate goal,” he declares, “is to assign function to all of these elements.”


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Alternative CRISPR discovered

Larry H. Bernstein, MD, FCAP, LPBI


New breakthrough! – A better alternative CRISPR system just identified

Knock-out, Genomic deletions, Gene therapy

CRISPR-Cas9 system has revolutionized the field of genome editing since its first application in human cells was reported in 2012. A recent publication in Cell reported the identification of a different CRISPR system with the potential for even simpler and more precise genome editing. The newly identified CRISPR-Cpf1 system mediates robust DNA interference with features different from Cas9. Cpf1 possesses several advantages over the currently used Cas9 system.

  • The Cpf1 system is simpler than Cas9 system as it requires only a single RNA for its DNA-cutting enzymatic activity.
  • Cpf1 cut has shot overhangs on the exposed ends, allowing more efficient and precise genome engineering; while Cas9 cut produces blunt ends that often undergo mutations when rejoined.
  • Cpf1 is smaller than Cas9, thus easier to deliver into the cells or tissues.
  • Cpf1 cut is far away from the recognition site, leaving space for further editing if mutation occurred at the cutting site.
  • The Cpf1 complex recognize very different PAM sequences than those of Cas9, adding more flexibility in choosing target sites.

These properties of Cpf1 and its potential with more precise gene editing expanded the application scope of CRISPR, from gene knock-out and knock-ins, genomic deletions, to even gene therapy.

Bernd Zetsche, etc. (September 2015) Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR-Case system. Cell


Cpf1 Is a Single RNA-Guided Endonuclease of a Class 2 CRISPR-Cas System  

Bernd Zetsche10, Jonathan S. Gootenberg10, Omar O. Abudayyeh, Ian M. Slaymaker, …, Eugene V. Koonin, Feng Zhang
Cell 2015 22 Oct ;  Volume 163, Issue 3, p759–771.
Figure thumbnail fx1
  • CRISPR-Cpf1 is a class 2 CRISPR 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



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.


Almost all archaea and many bacteria achieve adaptive immunity through a diverse set of CRISPR-Cas (clustered regularly interspaced short palindromicrepeats and CRISPR-associated proteins) systems, each of which consists of a combination of Cas effector proteins and CRISPR RNAs (crRNAs) (Makarova et al., 2011, Makarova et al., 2015). The defense activity of the CRISPR-Cas systems includes three stages: (1) adaptation, when a complex of Cas proteins excises a segment of the target DNA (known as a protospacer) and inserts it into the CRISPR array (where this sequence becomes a spacer); (2) expression and processing of the precursor CRISPR (pre-cr) RNA resulting in the formation of mature crRNAs; and (3) interference, when the effector module—either another Cas protein complex or a single large protein—is guided by a crRNA to recognize and cleave target DNA (or in some cases, RNA) (Horvath and Barrangou, 2010,Sorek et al., 2013, Barrangou and Marraffini, 2014). The adaptation stage is mediated by the complex of the Cas1 and Cas2 proteins, which are shared by all known CRISPR-Cas systems, and sometimes involves additional Cas proteins. Diversity is observed at the level of processing of the pre-crRNA to mature crRNA guides, proceeding via either a Cas6-related ribonuclease or a housekeeping RNaseIII that specifically cleaves double-stranded RNA hybrids of pre-crRNA and tracrRNA. Moreover, the effector modules differ substantially among the CRISPR-Cas systems (Makarova et al., 2011, Makarova et al., 2015,Charpentier et al., 2015). In the latest classification, the diverse CRISPR-Cas systems are divided into two classes according to the configuration of their effector modules: class 1 CRISPR systems utilize several Cas proteins and the crRNA to form an effector complex, whereas class 2 CRISPR systems employ a large single-component Cas protein in conjunction with crRNAs to mediate interference (Makarova et al., 2015).

Multiple class 1 CRISPR-Cas systems, which include the type I and type III systems, have been identified and functionally characterized in detail, revealing the complex architecture and dynamics of the effector complexes (Brouns et al., 2008, Marraffini and Sontheimer, 2008, Hale et al., 2009, Sinkunas et al., 2013,Jackson et al., 2014, Mulepati et al., 2014). Several class 2 CRISPR-Cas systems have also been identified and experimentally characterized, but they are all type II and employ homologous RNA-guided endonucleases of the Cas9 family as effectors (Barrangou et al., 2007, Garneau et al., 2010, Deltcheva et al., 2011, Sapranauskas et al., 2011, Jinek et al., 2012, Gasiunas et al., 2012). A second, putative class 2 CRISPR system, tentatively assigned to type V, has been recently identified in several bacterial genomes ( (Schunder et al., 2013, Vestergaard et al., 2014, Makarova et al., 2015). The putative type V CRISPR-Cas systems contain a large, ∼1,300 amino acid protein called Cpf1 (CRISPR from Prevotella and Francisella 1). It remains unknown, however, whether Cpf1-containing CRISPR loci indeed represent functional CRISPR systems. Given the broad applications of Cas9 as a genome-engineering tool (Hsu et al., 2014, Jiang and Marraffini, 2015), we sought to explore the function of Cpf1-based putative CRISPR systems.

Here, we show that Cpf1-containing CRISPR-Cas loci of Francisella novicida U112 encode functional defense systems capable of mediating plasmid interference in bacterial cells guided by the CRISPR spacers. Unlike Cas9 systems, Cpf1-containing CRISPR systems have three features. First, Cpf1-associated CRISPR arrays are processed into mature crRNAs without the requirement of an additional trans-activating crRNA (tracrRNA) (Deltcheva et al., 2011, Chylinski et al., 2013). Second, Cpf1-crRNA complexes efficiently cleave target DNA proceeded by a short T-rich protospacer-adjacent motif (PAM), in contrast to the G-rich PAM following the target DNA for Cas9 systems. Third, Cpf1 introduces a staggered DNA double-stranded break with a 4 or 5-nt 5′ overhang.

To explore the suitability of Cpf1 for genome-editing applications, we characterized the RNA-guided DNA-targeting requirements for 16 Cpf1-family proteins from diverse bacteria, and we identified two Cpf1 enzymes fromAcidaminococcus sp. BV3L6 and Lachnospiraceae bacterium ND2006 that are capable of mediating robust genome editing in human cells. Collectively, these results establish Cpf1 as a class 2 CRISPR-Cas system that includes an effective single RNA-guided endonuclease with distinct properties that has the potential to substantially advance our ability to manipulate eukaryotic genomes.


Large image of Figure 1.

Figure 1

The Francisella novicida U112 Cpf1 CRISPR Locus Provides Immunity against Transformation of Plasmids Containing Protospacers Flanked by a 5′-TTN PAM

(A) Organization of two CRISPR loci found in Francisella novicida U112 (NC_008601). The domain architectures of FnCas9 and FnCpf1 are compared.

(B) Schematic illustrating the plasmid depletion assay for discovering the PAM position and identity. Competent E. coliharboring either the heterologous FnCpf1 locus plasmid (pFnCpf1) or the empty vector control were transformed with a library of plasmids containing the matching protospacer flanked by randomized 5′ or 3′ PAM sequences and selected with antibiotic to deplete plasmids carrying successfully targeted PAM. Plasmids from surviving colonies were extracted and sequenced to determine depleted PAM sequences.

(C and D) Sequence logo for the FnCpf1 PAM as determined by the plasmid depletion assay. Letter height at each position is measured by information content (C) or frequency (D); error bars show 95% Bayesian confidence interval.

(E) E. coli harboring pFnCpf1 provides robust interference against plasmids carrying 5′-TTN PAMs (n = 3; error bars represent mean ± SEM).

See also Figure S1.

Cpf1-Containing CRISPR Loci Are Active Bacterial Immune Systems

The Cpf1-Associated CRISPR Array Is Processed Independent of TracrRNA

Cpf1 Is a Single crRNA-Guided Endonuclease

The RuvC-like Domain of Cpf1 Mediates RNA-Guided DNA Cleavage

Sequence and Structural Requirements for the Cpf1 crRNA

Cpf1-Family Proteins from Diverse Bacteria Share Common crRNA Structures and PAMs

Cpf1 Can Be Harnessed to Facilitate Genome Editing in Human Cells


In this work, we characterize Cpf1-containing class 2 CRISPR systems, classified as type V, and show that its effector protein, Cpf1, is a single RNA-guided endonuclease. Cpf1 substantially differs from Cas9—to date, the only other experimentally characterized class 2 effector—in terms of structure and function and might provide important advantages for genome-editing applications. Specifically, Cpf1 contains a single identified nuclease domain, in contrast to the two nuclease domains present in Cas9. The results presented here show that, in FnCpf1, inactivation of RuvC-like domain abolishes cleavage of both DNA strands. Conceivably, FnCpf1 forms a homodimer (Figure S2B), with the RuvC-like domains of each of the two subunits cleaving one DNA strand. However, we cannot rule out that FnCpf1 contains a second yet-to-be-identified nuclease domain. Structural characterization of Cpf1-RNA-DNA complexes will allow testing of these hypotheses and elucidation of the cleavage mechanism.


See also —-

Regulatory DNA engineered

Larry H. Bernstein, MD, FCAP, Curator



New Type of CRISPR Screen Probes the Regulatory Genome

Aaron Krol

February 8, 2016 | When a geneticist stares down the 3 billion DNA base pairs of the human genome, searching for a clue to what’s gone awry in a single patient, it helps to narrow the field. One of the most popular places to look is the exome, the tiny fraction of our DNA―less than 2%―that actually codes for proteins. For patients with rare genetic diseases, which might be fully explained by one key mutation, many studies sequence the whole exome and leave all the noncoding DNA out. Similarly, personalized cancer tests, which can help bring to light unexpected treatment options, often sequence the tumor exome, or a smaller panel of protein-coding genes.

Unfortunately, we know that’s not the whole picture. “There are a substantial number of noncoding regions that are just as effective at turning off a gene as a mutation in the gene itself,” says Richard Sherwood, a geneticist at Brigham and Women’s Hospital in Boston. “Exome sequencing is not going to be a good proxy for what genes are working.”

Sherwood studies regulatory DNA, the vast segment of the genome that governs which genes are turned on or off in any cell at a given time. It’s a confounding area of genetics; we don’t even know how much of the genome is made up of these regulatory elements. While genes can be recognized by the presence of “start” and “stop” codons―sequences of three DNA letters that tell the cell’s molecular machinery which stretches of DNA to transcribe into RNA, and eventually into protein―there are no definite signs like this for regulatory DNA.

Instead, studies to discover new regulatory elements have been somewhat trial-and-error. If you suspect a gene’s activity might be regulated by a nearby DNA element, you can inhibit that element in a living cell, and see if your gene shuts down with it.

With these painstaking experiments, scientists can slowly work their way through potential regulatory regions―but they can’t sweep across the genome with the kind of high-throughput testing that other areas of genetics thrive on. “Previously, you couldn’t do these sorts of tests in a large form, like 4,000 of them at once,” says David Gifford, a computational biologist at MIT. “You would really need to have a more hypothesis-directed methodology.”

Recently, Gifford and Sherwood collaborated on a paper, published in Nature Biotechnology, which presents a new method for testing thousands of DNA loci for regulatory activity at once. Their assay, called MERA (multiplexed editing regulatory assay), is built on the recent technology boom in CRISPR-Cas9 gene editing, which lets scientists quickly and easily cut specific sequences of DNA out of the genome.

So far, their team, including lead author Nisha Rajagopal from Gifford’s lab, has used MERA to study the regulation of four genes involved in the development of embryonic stem cells. Already, the results have defied the accepted wisdom about regulatory DNA. Many areas of the genome flagged by MERA as important factors in gene expression do not fall into any known categories of regulatory elements, and would likely never have been tested with previous-generation methods.

“Our approach allows you to look away from the lampposts,” says Sherwood. “The more unbiased you can be, the more we’ll actually know.”

A New Kind of CRISPR Screen

In the past three years, CRISPR-Cas9 experiments have taken all areas of molecular biology by storm, and Sherwood and Gifford are far from the first to use the technology to run large numbers of tests in parallel. CRISPR screens are an excellent way to learn which genes are involved in a cellular process, like tumor growth or drug resistance. In these assays, scientists knock out entire genes, one by one, and see what happens to cells without them.

This kind of CRISPR screen, however, operates on too small a scale to study the regulatory genome. For each gene knocked out in a CRISPR screen, you have to engineer a strain of virus to deliver a “guide RNA” into the cellular genome, showing the vicelike Cas9 molecule which DNA region to cut. That works well if you know exactly where a gene lies and only need to cut it once—but in a high-throughput regulatory test, you would want to blanket vast stretches of DNA with cuts, not knowing which areas will turn out to contain regulatory elements. Creating a new virus for each of these cuts is hugely impractical.

The insight behind MERA is that, with the right preparation, most of the genetic engineering can be done in advance. Gifford and Sherwood’s team used a standard viral vector to put a “dummy” guide RNA sequence, one that wouldn’t tell Cas9 to cut anything, into an embryonic stem cell’s genome. Then they grew plenty of cells with this prebuilt CRISPR system inside, and attacked each one with a Cas9 molecule targeted to the dummy sequence, chopping out the fake guide.

Normally, the result would just be a gap in the CRISPR system where the guide once was. But along with Cas9, the researchers also exposed the cells to new, “real” guide RNA sequences. Through a DNA repair mechanism called homologous recombination, the cells dutifully patched over the gaps with new guides, whose sequences were very similar to the missing dummy code. At the end of the process, each cell had a unique guide sequence ready to make cuts at a specific DNA locus—just like in a standard CRISPR screen, but with much less hands-on engineering.

By using a large enough library of guide RNA molecules, a MERA screen can include thousands of cuts that completely tile a broad region of the genome, providing an agnostic look at anywhere regulatory elements might be hiding. “It’s a lot easier [than a typical CRISPR screen],” says Sherwood. “The day the library comes in, you just perform one PCR reaction, and the cells do the rest of the work.”

In the team’s first batch of MERA screens, they created almost 4,000 guide RNAs for each gene they studied, covering roughly 40,000 DNA bases of the “cis-regulatory region,” or the area surrounding the gene where most regulatory elements are thought to lie. It’s unclear just how large any gene’s cis-regulatory region is, but 40,000 bases is a big leap from the highly targeted assays that have come before.

“We’re now starting to do follow-up studies where we increase the number of guide RNAs,” Sherwood adds. “Eventually, what you’d like is to be able to tile an entire chromosome.”

Far From the Lampposts

Sherwood and Gifford tried to focus their assays on regions that would be rich in regulatory elements. To that end, they made sure their guide RNAs covered parts of the genome with well-known signs of regulatory activity, like histone markers and transcription factor binding sites. For many of these areas, Cas9 cuts did, in fact, shut down gene expression in the MERA screens.

But the study also targeted regions around each gene that were empty of any known regulatory features. “We tiled some other regions that we thought might serve as negative controls,” explains Gifford. “But they turned out not to be negative at all.”

The study’s most surprising finding was that several cuts to seemingly random areas of the genome caused genes to become nonfunctional. The authors named these DNA regions “unmarked regulatory elements,” or UREs. They were especially prevalent around the genes Tdgf1 and Zfp42, and in many cases, seemed to be every bit as necessary to gene activity as more predictable hits on the MERA screen.

These results caught the researchers so off guard that it was natural to wonder if MERA screens are prone to false positives. Yet follow-up experiments strongly supported the existence of UREs. Switching the guide RNAs from aTdgf1 MERA screen and aZfp42 screen, for example, produced almost no positive results: the UREs’ regulatory effects were indeed specific to the genes near them.

In a more specific test, the researchers chose a particular URE connected to Tdgf1, and cut it out of a brand new population of cells for a closer look. “We showed that, if we deleted that region from the genome, the cells lost expression of the gene,” says Sherwood. “And then when we put it back in, the gene became expressed again. Which was good proof to us that the URE itself was responsible.”

From these results, it seems likely that follow-up MERA screens will find even more unknown stretches of regulatory DNA. Gifford and Sherwood’s experiments didn’t try to cover as much ground around their target genes as they might have, because the researchers assumed that MERA would mostly confirm what was already known. At best, they hoped MERA would rule out some suspected regulatory regions, and help show which regulatory elements have the biggest effect on gene expression.

“We tended to prioritize regions that had been known before,” Sherwood says. “Unfortunately, in the end, our datasets weren’t ideally suited to discovering these UREs.”

Getting to Basic Principles

MERA could open up huge swaths of the regulatory genome to investigation. Compared to an ordinary CRISPR screen, says Sherwood, “there’s only upside,” as MERA is cheaper, easier, and faster to run.

Still, interpreting the results is not trivial. Like other CRISPR screens, MERA makes cuts at precise points in the genome, but does not tell cells to repair those cuts in any particular way. As a result, a population of cells all carrying the same guide RNA can have a huge variety of different gaps and scars in their genomes, typically deletions in the range of 10 to 100 bases long. Gifford and Sherwood created up to 100 cells for each of their guides, and sometimes found that gene expression was affected in some but not all of them; only sequencing the genomes of their mutated cells could reveal exactly what changes had been made.

By repeating these experiments many times, and learning which mutations affect gene expression, it will eventually be possible to pin down the exact DNA bases that make up each regulatory element. Future studies might even be able to distinguish between regulatory elements with small and large effects on gene expression. In Gifford and Sherwood’s MERA screens, the target genes were altered to produce a green fluorescent protein, so the results were read in terms of whether cells gave off fluorescent light. But a more precise, though expensive, approach would be to perform RNA sequencing, to learn which cuts reduced the cell’s ability to transcribe a gene into RNA, and by how much.

A MERA screen offers a rich volume of data on the behavior of the regulatory genome. Yet, as with so much else in genetics, there are few robust principles to let scientists know where they should be focusing their efforts. Histone markers provide only a very rough sketch of regulatory elements, often proving to be red herrings on closer examination. And the existence of UREs, if confirmed by future experiments, shows that we don’t yet even know which areas of the genome to rule out in the hunt for regulatory regions.

“Every dataset we get comes closer and closer to computational principles that let us predict these regions,” says Sherwood. As more studies are conducted, patterns may emerge in the DNA sequences of regulatory elements that link UREs together, or reveal which histone markers truly point toward regulatory effects. There might also be functional clues hidden in these sequences, hinting at what is happening on a molecular level as regulatory elements turn genes on and off in the course of a cell’s development.

For now, however, the data is still rough and disorganized. For better and for worse, high-throughput tools like MERA are becoming the foundation for most discoveries in genetics—and that means there is a lot more work to do before the regulatory genome begins to come into focus.

CORRECTED 2/9/16: Originally, this story incorrectly stated that only certain cell types could be assayed with MERA for reasons related to homologous recombination. In fact, the authors see no reason MERA could not be applied to any in vitro cell line, and hope to perform screens in a wide range of cell types. The text has been edited to correct the error.


Gene Editing for Exon 51: Why CRISPR Snipping might be better than Exon Skipping for DMD

Reporter: Aviva Lev-Ari, PhD, RN

Why CRISPR might be better than exon skipping for DMD: Snipping vs. skipping for DMD

By Lauren Martz, Senior Writer
Published on Thursday, January 21, 2016

As if to preempt the regulatory setbacks in Duchenne muscular dystrophy (DMD) that last week disappointed the field, a trio of preclinical studies emerged two weeks earlier showing that cutting out DMD mutations with gene editing might offer a viable alternative to the exon-skipping strategies that have dominated the pipeline. Now, the question is whether there’s reason to believe the mouse studies will translate any better to the clinic.

The studies, published Dec. 31 in Science, provide in vivo proof of concept for the first time that CRISPR-Cas9 used postnatally can have a disease-modifying effect. Despite the hype around its therapeutic promise, the technology has so far proved itself primarily in research applications, for example, in modifying cells for in vitro screening or creating animal models of disease.


Other related articles published in this Open Access Online Scientific Journal include the following:

Gene Editing: The Role of Oligonucleotide Chips

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

Gene Editing by creation of a complement without transcription error

Curator: Larry H. Bernstein, MD, FCAP

UPDATED – Medical Interpretation of the Genomics Frontier – CRISPR – Cas9:  Gene Editing Technology for New Therapeutics

Authors and Curators: Larry H Bernstein, MD, FCAP and Stephen J Williams, PhD and Curator: Aviva Lev-Ari, PhD, RN

DNA Structure and Oligonucleotides

Curator: Larry H Bernstein, MD, FCAP


Genome Engineering: Genome Editing with CRISPR-Cas9

Reporter: Aviva Lev-Ari, PhD, RN

Two videos

DIY CRISPR Genome Engineering



2:24 / 4:12

Genome Editing with CRISPR-Cas9




RNAi, CRISPR and Gene Expression

Larry H. Bernstein, MD, FCAP, Curator


Down and Out with RNAi and CRISPR

Gene-Silencing and Gene-Disabling Techniques Are Moving To the Heart of Drug Discovery

  • Click Image To Enlarge +
    RNA interference (RNAi) silences, or knocks down, the translation of a gene by inducing degradation of a gene target’s transcript. To advance RNAi applications, Thermo Fisher Scientific has developed two types of small RNA molecules: short interfering RNAs and microRNAs. The company also offers products for RNAi analysis in vitro and in vivo, including libraries for high-throughput applications.

    Genes can be knocked down with RNA interference (RNAi) or knocked out with CRISPR-Cas9. RNAi, the screening workhorse, knocks down the translation of genes by inducing rapid degradation of a gene target’s transcript.

    CRISPR-Cas9, the new but already celebrated genome-editing technology, cleaves specific DNA sequences to render genes inoperative. Although mechanistically different, the two techniques complement one another, and when used together facilitate discovery and validation of scientific findings.

    RNAi technologies along with other developments in functional genomics screening were discussed by industry leaders at the recent Discovery on Target conference. The conference, which emphasized the identification and validation of novel drug targets and the exploration of unknown cellular pathways, included a symposium on the development of CRISPR-based therapies.

    RNAi screening can be performed in either pooled or arrayed formats. Pooled screening provides an affordable benchtop option, but requires back-end deconvolution and deep sequencing to identify the shRNA causing the specific phenotype. Targets are much easier to identify using the arrayed format since each shRNA clone is in an individual well.

    “CRISPR complements RNAi screens,” commented Ryan Raver, Ph.D., global product manager of functional genomics at Sigma-Aldrich. “You can do a whole genome screen with either small interfering RNA (siRNA) or small hairpin RNA (shRNA), then validate with individual CRISPRs to ensure it is a true result.”

    A powerful and useful validation method for knockdown or knockout studies is to use lentiviral open reading frames (ORFs) for gene re-expression, for rescue experiments, or to detect whether the wild-type phenotype is restored. However, the ORF randomly integrates into the genome. Also, with this validation technique, gene expression is not acting under the endogenous promoter. Accordingly, physiologically relevant levels of the gene may not be expressed unless controlled for via an inducible system.

    In the future, CRISPR activators may provide more efficient ways to express not only wild-type but also mutant forms of genes under the endogenous promoter.

    Choice of screening technique depends on the researcher and the research question. Whole gene knockout may be necessary to observe a phenotype, while partial knockdown might be required to investigate functions of essential or lethal genes. Use of both techniques is recommended to identify all potential targets.

    For example, recently, a whole genome siRNA screen on a human glioblastoma cell line identified a gene, known as FAT1, as a negative regulator of apoptosis. A CRISPR-mediated knockout of the gene also conferred sensitivity to death receptor–induced apoptosis with an even stronger phenotype, thereby validating FAT1’s new role and link to extrinsic apoptosis, a new model system.

    Dr. Raver indicated that next-generation RNAi libraries that are microRNA-adapted might have a more robust knockdown of gene expression, up to 90–95% in some cases. Ultracomplex shRNA libraries help to minimize both false-negative and false-positive rates by targeting each gene with ~25 independent shRNAs and by including thousands of negative-control shRNAs.

    Recently, a relevant paper emerged from the laboratory of Jonathan Weissman, Ph.D., a professor of cellular and molecular pharmacology at the University of California, San Francisco. The paper described how a new ultracomplex pooled shRNA library was optimized by means of a microRNA-adapted system. This system, which was able to achieve high specificity in the detection of hit genes, produced robust results. In fact, they were comparable to results obtained with a CRISPR pooled screen. Members of the Weissman group systematically optimized the promoter and microRNA contexts for shRNA expression along with a selection of guide strands.

    Using a sublibrary of proteostasis genes (targeting 2,933 genes), the investigators compared CRISPR and RNAi pooled screens. Data showed 48 hits unique to RNAi, 40 unique to CRISPR, and an overlap of 21 hits (with a 5% false discovery rate cut-off). Together, the technologies provided a more complete research story.

    Arrayed CRISPR Screens

  • Click Image To Enlarge +
    Synthetic crRNA:tracrRNA reagents can be used in a similar manner to siRNA reagents for assessment of phenotypes in a cell population. Top row: A reporter cell line stably expressing Cas9 nuclease was transfected with GE Dharmacon’s Edit-R synthetic crRNA:tracrRNA system, which was used to target three positive control genes (PSMD7, PSMD14, and VCP) and a negative control gene (PPIB). Green cells indicate EGFP signaling occuring as a result of proteasome pathway disruption. Bottom row: A siGENOME siRNA pool targeting the same genes was used in the same reporter cell line.

    “RNA screens are well accepted and will continue to be used, but it is important biologically that researchers step away from the RNA mechanism to further study and validate their hits to eliminate potential bias,” explained Louise Baskin, senior product manager, Dharmacon, part of GE Healthcare. “The natural progression is to adopt CRISPR in the later stages.”

    RNAi uses the cell’s endogenous mechanism. All of the components required for gene knockdown are already within the cell, and the delivery of the siRNA starts the process. With the CRISPR gene-editing system, which is derived from a bacterial immune defense system, delivery of both the guide RNA and the Cas9 nuclease, often the rate limiter in terms of knockout efficiency, are required.

    In pooled approaches, the cell has to either drop out or overexpress so that it is sortable, limiting the types of addressable biological questions. A CRISPR-arrayed approach opens up the door for use of other analytical tools, such as high-content imaging, to identify hits of interest.

    To facilitate use of CRISPR, GE recently introduced Dharmacon Edit-R synthetic CRISPR RNA (crRNA) libraries that can be used to carry out high-throughput arrayed analysis of multiple genes. Rather than a vector- or plasmid-based gRNA to guide the targeting of the Cas9 cleavage, a synthetic crRNA and tracrRNA are used. These algorithm-designed crRNA reagents can be delivered into the cells very much like siRNA, opening up the capability to screen multiple target regions for many different genes simultaneously.

    GE showed a very strong overlap between CRISPR and RNAi using this arrayed approach to validate RNAi screen hits with synthetic crRNA. The data concluded that CRISPR can be used for medium- or high-throughput validation of knockdown studies.

    “We will continue to see a lot of cooperation between RNAi and gene editing,” declared Baskin. “Using the CRISPR mechanism to knockin could introduce mutations to help understand gene function at a much deeper level, including a more thorough functional analysis of noncoding genes.

    “These regulatory RNAs often act in the nucleus to control translation and transcription, so to knockdown these genes with RNAi would require export to the cytoplasm. Precision gene editing, which takes place in the nucleus, will help us understand the noncoding transcriptome and dive deeper into how those genes regulate disease progression, cellular development and other aspects of human health and biology.”

    Tool Selection

    Click Image To Enlarge +
    Schematic of a pooled shRNA screening workflow developed by Transomic Technologies. Cells are transduced, and positive or negative selection screens are performed. PCR amplification and sequencing of the shRNA integrated into the target cell genome allows the determination of shRNA representation in the population.

    The functional genomics tool should fit the specific biology; the biology should not be forced to fit the tool. Points to consider include the regulation of expression, the cell line or model system, as well as assay scale and design. For example, there may be a need for regulatable expression. There may be limitations around the cell line or model system. And assay scale and design may include delivery conditions and timing to optimally complete perturbation and reporting.

    “Both RNAi- and CRISPR-based gene modulation strategies have pros and cons that should be considered based on the biology of the genes being studied,” commented Gwen Fewell, Ph.D., chief commercial officer, Transomic Technologies.

    RNAi reagents, which can produce hypomorphic or transient gene-suppression states, are well known for their use in probing drug targets. In addition, these reagents are enriching studies of gene function. CRISPR-Cas9 reagents, which produce clean heterozygous and null mutations, are important for studying tumor suppressors and other genes where complete loss of function is desired.

    Timing to readout the effects of gene perturbation must be considered to ensure that the biological assay is feasible. RNAi gene knockdown effects can be seen in as little as 24–72 hours, and inducible and reversible gene knockdown can be realized. CRISPR-based gene knockout effects may become complete and permanent only after 10 days.

    Both RNAi and CRISPR reagents work well for pooled positive selection screens; however, for negative selection screens, RNAi is the more mature tool. Current versions of CRISPR pooled reagents can produce mixed populations containing a fraction of non-null mutations, which can reduce the overall accuracy of the readout.

    To meet the needs of varied and complex biological questions, Transomic Technologies has developed both RNAi and CRISPR tools with options for optimal expression, selection, and assay scale. For example, the company’s shERWOOD-UltramiR shRNA reagents incorporate advances in design and small RNA processing to produce increased potency and specificity of knockdown, particularly important for pooled screens.

    Sequence-verified pooled shRNA screening libraries provide flexibility in promoter choice, in vitro formats, in vivo formats, and a choice of viral vectors for optimal delivery and expression in biologically relevant cell lines, primary cells or in vivo.

    The company’s line of lentiviral-based CRISPR-Cas9 reagents has variable selectable markers such that guide RNA- and Cas9-expressing vectors, including inducible Cas9, can be co-delivered and selected for in the same cell to increase editing efficiency. Promoter options are available to ensure expression across a range of cell types.

    “Researchers are using RNAi and CRISPR reagents individually and in combination as cross-validation tools, or to engineer CRISPR-based models to perform RNAi-based assays,” informs Dr. Fewell. “Most exciting are parallel CRISPR and RNAi screens that have tremendous potential to uncover novel biology.”

    Converging Technologies

    The convergence of RNAi technology with genome-editing tools, such as CRISPR-Cas9 and transcription activator-like effector nucleases, combined with next-generation sequencing will allow researchers to dissect biological systems in a way not previously possible.

    “From a purely technical standpoint, the challenges for traditional RNAi screens come down to efficient delivery of the RNAi reagents and having those reagents provide significant, consistent, and lasting knockdown of the target mRNAs,” states Ross Whittaker, Ph.D., a product manager for genome editing products at Thermo Fisher Scientific. “We have approached these challenges with a series of reagents and siRNA libraries designed to increase the success of RNAi screens.”

    Thermo Fisher’ provides lipid-transfection RNAiMax reagents, which effectively deliver siRNA. In addition, the company’s Silencer and Silencer Select siRNA libraries provide consistent and significant knockdown of the target mRNAs. These siRNA libraries utilize highly stringent bioinformatic designs that ensure accurate and potent targeting for gene-silencing studies. The Silencer Select technology adds a higher level of efficacy and specificity due to chemical modifications with locked nucleic acid (LNA) chemistry.

    The libraries alleviate concerns for false-positive or false-negative data. The high potency allows less reagent use; thus, more screens or validations can be conducted per library.

    Dr. Whittaker believes that researchers will migrate regularly between RNAi and CRISPR-Cas9 technology in the future. CRISPR-Cas9 will be used to create engineered cell lines not only to validate RNAi hits but also to follow up on the underlying mechanisms. Cell lines engineered with CRISPR-Cas9 will be utilized in RNAi screens. In the long term, CRISPR-Cas9 screening will likely replace RNAi screening in many cases, especially with the introduction of arrayed CRISPR libraries.

    Validating Antibodies with RNAi

    Unreliable antibody specificity is a widespread problem for researchers, but RNAi is assuaging scientists’ concerns as a validation method.

    The procedure introduces short hairpin RNAs (shRNAs) to reduce expression levels of a targeted protein. The associated antibody follows. With its protein knocked down, a truly specific antibody shows dramatically reduced or no signal on a Western blot. Short of knockout animal models, RNAi is arguably the most effective method of validating research antibodies.

    The method is not common among antibody suppliers—time and cost being the chief barriers to its adoption, although some companies are beginning to embrace RNAi validation.

    “In the interest of fostering better science, Proteintech felt it was necessary to implement this practice,” said Jason Li, Ph.D., founder and CEO of Proteintech Group, which made RNAi standard protocol in February 2015. “When researchers can depend on reproducibility, they execute more thorough experiments and advance the treatment of human diseases and conditions.”


Junk DNA Kept in Good Repair by Nuclear Membrane  

Heterochromatin has the dubious distinction of being called the “dark matter” of DNA, and it has even suffered the indignity of being dismissed as “junk DNA.” But it seems to get more respectful treatment inside the nucleus, where it has the benefit of a special repair mechanism. This mechanism, discovered by scientists based at the University of Southern California (USC), transports broken heterochromatin sequences from the hurly-burly of the heterochromatin domain so that they can be repaired in the relative peace and quiet of the nuclear periphery.

This finding suggests that the nuclear membrane is more versatile than is generally appreciated. Yes, it serves as a protective container for nuclear material, and it uses its pores to manage the transport of molecules in and out of the nucleus. But it may also play a special role in maintaining the integrity of heterochromatin, which tends to be overlooked because it consists largely of noncoding DNA, including repetitive stretches of no apparent function.

“Scientists are now starting to pay a lot of attention to this mysterious component of the genome,” said Irene E. Chiolo, Ph.D., an assistant professor at USC. “Heterochromatin is not only essential for chromosome maintenance during cell division; it also poses specific threats to genome stability. Heterochromatin is potentially one of the most powerful driving forces for cancer formation, but it is the ‘dark matter’ of the genome. We are just beginning to unravel how repair works here.”

Dr. Chilo led an effort to understand how heterochromatin stays in good repair, even though it is particularly vulnerable to a kind of repair error called ectopic recombination. This kind of error is apt to occur when flaws in repeated sequences undergo homologous recombination (HR) by means of double-strand break (DSB) repair. Specifically, repeated sequences tend to recombine with each other during DNA repair.

Working with the fruit fly Drosophila melanogaster, Dr. Chilo’s team observed that breaks in heterochromatin are repaired after damaged sequences move away from the rest of the chromosome to the inner wall of the nuclear membrane. There, a trio of proteins mends the break in a safe environment, where it cannot accidentally get tangled up with incorrect chromosomes.

The details appeared October 26 in Nature Cell Biology, in an article entitled, “Heterochromatic breaks move to the nuclear periphery to continue recombinational repair.”

“[Heterochromatic] DSBs move to the nuclear periphery to continue HR repair,” the authors wrote. “Relocalization depends on nuclear pores and inner nuclear membrane proteins (INMPs) that anchor repair sites to the nuclear periphery through the Smc5/6-interacting proteins STUbL/RENi. Both the initial block to HR progression inside the heterochromatin domain, and the targeting of repair sites to the nuclear periphery, rely on SUMO and SUMO E3 ligases.”

“We knew that nuclear membrane dysfunctions are common in cancer cells,” Dr. Chiolo said. “Our studies now suggest how these dysfunctions can affect heterochromatin repair and have a causative role in cancer progression.”

This study may help reveal how and why organisms become more predisposed to cancer as they age—the nuclear membrane progressively deteriorates as an organism ages, removing this bulwark against genome instability.

Next, Dr. Chiolo and her team will explore how the movement of broken sequences is accomplished and regulated, and what happens in cells and organisms when this membrane-based repair mechanism fails. Their ultimate goal is to understand how this mechanism functions in human cells and identify new strategies to prevent their catastrophic failure and cancer formation.

Gene Found that Regulates Stem Cell Number Production

Gene Found that Regulates Stem Cell Number Production

The gene Prkci promotes the generation of differentiated cells (red). However if Prkci activity is reduced or absent, neural stem cells (green) are promoted. [In Kyoung Mah]

A scientific team from the University of Southern California (USC) and the University of California, San Diego have described an important gene that maintains a critical balance between producing too many
and too few stem cells. Called Prkci, the gene influences whether stem cells self-renew to produce more stem cells, or differentiate into more specialized cell types, such as blood or nerves.

When it comes to stem cells, too much of a good thing isn’t necessarily a benefit: producing too many new stem cells may lead to cancer; making too few inhibits the repair and maintenance of the body.

In their experiments, the researchers grew mouse embryonic stem cells, which lacked Prkci, into embryo-like structures in the laboratory. Without Prkci, the stem cells favored self-renewal, generating large numbers of stem cells and, subsequently, an abundance of secondary structures.

Upon closer inspection, the stem cells lacking Prkci had many activated genes typical of stem cells, and some activated genes typical of neural, cardiac, and blood-forming cells. Therefore, the loss of Prkci can also encourage stem cells to differentiate into the progenitor cells that form neurons, heart muscle, and blood.

Prkci achieves these effects by activating or deactivating a well-known group of interacting genes that are part of the Notch signaling pathway. In the absence of Prkci, the Notch pathway produces a protein that signals to stem cells to make more stem cells. In the presence of Prkci, the Notch pathway remains silent, and stem cells differentiate into specific cell types.

These findings have implications for developing patient therapies. Even though Prkci can be active in certain skin cancers, inhibiting it might lead to unintended consequences, such as tumor overgrowth. However, for patients with certain injuries or diseases, it could be therapeutic to use small molecule inhibitors to block the activity of Prkci, thus boosting stem cell production.

“We expect that our findings will be applicable in diverse contexts and make it possible to easily generate stem cells that have typically been difficult to generate,” said Francesca Mariani, Ph.D., principal investigator at the Eli and Edythe Broad Center for Regenerative Medicine and Stem Cell Research at USC.

Their study (“Atypical PKC-iota Controls Stem Cell Expansion via Regulation of the Notch Pathway”) was published in a Stem Cell Reports.

Atypical PKC-iota Controls Stem Cell Expansion via Regulation of the Notch Pathway

In Kyoung Mah,1 Rachel Soloff,2,3 Stephen M. Hedrick,2 and Francesca V. Mariani1, *

Stem Cell Reports (2015),

The number of stem/progenitor cells available can profoundly impact tissue homeostasis and the response to injury or disease. Here, we propose that an atypical PKC, Prkci, is a key player in regulating the switch from an expansion to a differentiation/maintenance phase via regulation of Notch, thus linking the polarity pathway with the control of stem cell self-renewal. Prkci is known to influence symmetric cell division in invertebrates; however a definitive role in mammals has not yet emerged. Using a genetic approach, we find that loss of Prkci results in a marked increase in the number of various stem/progenitor cells. The mechanism used likely involves inactivation and symmetric localization of NUMB, leading to the activation of NOTCH1 and its downstream effectors. Inhibition of atypical PKCs may be useful for boosting the production of pluripotent stem cells, multipotent stem cells, or possibly even primordial germ cells by promoting the stem cell/progenitor fate.

The control of asymmetric versus symmetric cell division in stem and progenitor cells balances self-renewal and differentiation to mediate tissue homeostasis and repair and involves key proteins that control cell polarity. In the case of excess symmetric division, too many stem-cell-like daughter cells are generated that can lead to tumor initiation and growth. Conversely, excess asymmetric cell division can severely limit the number of cells available for homeostasis and repair (Go´mez-Lo´pez et al., 2014; Inaba and Yamashita, 2012). The Notch pathway has been implicated in controlling stem cell self-renewal in a number of different contexts (Hori et al., 2013). However, how cell polarity, asymmetric cell division, and the activation of determinants ultimately impinges upon the control of stem cell expansion and maintenance is not fully understood. In this study, we examine the role of an atypical protein kinase C (aPKC), PRKCi, in stem cell self-renewal and, in particular, determine whether PRKCi acts via the Notch pathway. PKCs are serine-threonine kinases that control many basic cellular processes and are typically classified into three subgroups—conventional, novel, and the aPKCs iota and zeta, which, in contrast to the others, are not activated by diacylglyceride or calcium. The aPKC proteins are best known for being central components of an evolutionarily conserved Par3-Par6-aPKC trimeric complex that controls cell polarity in C. elegans, Drosophila, Xenopus, zebrafish, and mammalian cells (Suzuki and Ohno, 2006).

Before Notch influences stem cell self-renewal, the regulation of cell polarity, asymmetric versus symmetric cell division, and the segregation of cell fate determinants such as NUMB may first be required (Knoblich, 2008). For example, mutational analysis in Drosophila has demonstrated that the aPKC-containing trimeric complex is required for maintaining polarity and for mediating asymmetric cell division during neurogenesis via activation and segregation of NUMB (Wirtz-Peitz et al., 2008). NUMB then functions as a cell fate determinant by inhibiting Notch signaling and preventing self-renewal (Wang et al., 2006). In mammals, the PAR3-PAR6-aPKC complex also can bind and phosphorylate NUMB in epithelial cells and can regulate the unequal distribution of Numb during asymmetric cell division (Smith et al., 2007). During mammalian neurogenesis, asymmetric division is also thought to involve the PAR3-PAR6-aPKC complex, NUMB segregation, and NOTCH activation (Bultje et al., 2009).

Mice deficient in Prkcz are grossly normal, with mild defects in secondary lymphoid organs (Leitges et al., 2001). In contrast, deficiency of the Prkci isozyme results in early embryonic lethality at embryonic day (E)9.5 (Seidl et al., 2013; Soloff et al., 2004). A few studies have investigated the conditional inactivation of Prkci; however, no dramatic changes in progenitor generation were detected in hematopoietic stem cells (HSCs) or the brain (Imai et al., 2006; Sengupta et al., 2011), although one study found evidence of a role for Prkci in controlling asymmetric cell division in the skin (Niessen et al., 2013). Analysis may be complicated by functional redundancy between the iota and zeta isoforms and/or because further studies perturbing aPKCs in specific cell lineages and/or at specific developmental stages are needed.

Here, we investigate the requirement of Prkci in mouse cells using an in vitro system that bypasses early embryonic lethality. Embryonic stem (ES) cells are used to make embryoid bodies (EBs) that develop like the early post-implantation embryo in terms of lineage specification and morphology and can also be maintained in culture long enough to observe advanced stages of cellular differentiation (Desbaillets et al., 2000). Using this approach, we provide genetic evidence that inactivation of Prkci signaling leads to enhanced generation of pluripotent cells and some types of multipotent stem cells, including cells with primordial germ cell (PGC) characteristics. In addition, we provide evidence that aPKCs ultimately regulate stem cell fate via the Notch pathway.

Figure 1. Prkci/ EBs Contain Cells with Pluripotency Characteristics (A and A0 ) Day (d) 12 heterozygous EBs have few OCT4/E-CAD+ cells, while null EBs contain many in clusters at the EB periphery. Inset: OCT4 (nucleus)/E-CAD (cytoplasm) double-positive cells. (B and B0 ) Adjacent sections in a null EB show that OCT4+ cells are likely also SSEA1+. (C) Dissociated day-12 Prkci/ EBs contain five to six times more OCT4+ and approximately three times more SSEA1+ cells than heterozygous EBs (three independent experiments). (D and D0 ) After 2 days in ES cell culture, no colonies are visible in null SSEA1 cultures while present in null SSEA1+ cultures (red arrows). (E–E00) SSEA1+ sorted cells can be maintained for many passages, 27+. (E) Prkci+/ sorted cells make colonies with differentiated cells at the outer edges (n = 27/35). (E0 ) Null cells form colonies with distinct edges (n = 39/45). (E00) The percentage of undifferentiated colonies is shown. ***p < 0.001. (F) Sorted null cells express stem cell and differentiation markers at similar levels to normal ES cells (versus heterozygous EBs) (three independent experiments). (G) EBs made from null SSEA1+ sorted cells express germ layer marker genes at the indicated days. Error bars indicate mean ± SEM, three independent experiments. Scale bars, 100 mm in (A, D, and E); 25 mm in (B). See also Figure S1.


Prkci/ Cultures Have More Pluripotent Cells Even under Differentiation Conditions First, we compared Prkci null EB development to that of Prkci/ embryos. Consistent with another null allele (Seidl et al., 2013), both null embryos and EBs fail to properly cavitate (Figures S1A and S1B). The failure to cavitate is unlikely to be due to the inability to form one of the three germ layers, as null EBs express germ-layer-specific genes (Figure S1E). A failure of cavitation could alternatively be caused by an accumulation of pluripotent cells. For example, EBs generated from Timeless knockdown cells do not cavitate and contain large numbers of OCT4-expressing cells (O’Reilly et al., 2011). In addition, EBs generated with Prkcz isoform knockdown cells contain OCT4+ cells under differentiation conditions (Dutta et al., 2011; Rajendran et al., 2013). Thus, we first evaluated ES colony differentiation by alkaline phosphatase (AP) staining. After 4 days without leukemia inhibitory factor (LIF), Prkci/ ES cell colonies retained crisp boundaries and strong AP staining. In contrast, Prkci+/ colonies had uneven colony boundaries with diffuse AP staining (Figures S1F–S1F00). To definitively detect pluripotent cells, day-12 EBs were assayed for OCT4 and E-CADHERIN (E-CAD) protein expression. Prkci+/ EBs had very few OCT4/E-CAD double-positive cells (Figure 1A); however, null EBs contained large clusters of OCT4/E-CAD double-positive cells, concentrated in a peripheral zone (Figure 1A0 ). By examining adjacent sections, we found that OCT4+ cells could also be positive for stage-specific embryonic antigen 1 (SSEA1) (Figures 1B and 1B0 ). Quantification by fluorescence-activated cell sorting (FACS) analysis showed that day-12 Prkci/ EBs had more OCT4+ and SSEA1+ cells than Prkci+/ EBs (Figure 1C). We did not find any difference between heterozygous and wild-type cells with respect to the number of OCT4+ or SSEA1+ cells or in their levels of expression for Oct4, Nanog, and Sox2 (Figures S1I, S1I0 and S1J). However, we did find that Oct4, Nanog, and Sox2 were highly upregulated in OCT4+ null cells (Figure S1G). Thus, together, these data indicate that Prkci/ EBs contain large numbers of pluripotent stem cells, despite being cultured under differentiation conditions.

Functional Pluripotency Tests If primary EBs have a pluripotent population with the capacity to undergo self-renewal, they can easily form secondary EBs (O’Reilly et al., 2011). Using this assay, we found that more secondary EBs could be generated from Prkci/ versus Prkci+/ EBs, especially at days 6, 10, and 16; even when plated at a low density to control for aggregation (Figure S1H). To test whether SSEA1+ cells could maintain pluripotency long term, FACS-sorted Prkci/ SSEA1+ and SSEA1 cells were plated at a low density and maintained under ES cell culture conditions. SSEA1 cells were never able to form identifiable colonies and could not be maintained in culture (Figure 1D). SSEA1+ cells, however, formed many distinct colonies after 2 days of culture, and these cells could be maintained for over 27 passages (Figures 1D0 , 1E0 , and 1E00). Prkci+/ SSEA1+ cells formed colonies that easily differentiated at the outer edge, even in the presence of LIF (Figure 1E). In contrast Prkci/ SSEA1+ cells maintained distinct round colonies (Figure 1E0 ). Next, we determined whether null SSEA1+ cells expressed pluripotency and differentiation markers similarly to normal ES cells. Indeed, we found that Oct4, Nanog, and Sox2 were upregulated in both null SSEA1+ EB cells and heterozygous ES cells. In addition, differentiated markers (Fgf5, T, Wnt3, and Afp) and tissue stem/progenitor cell markers (neural: Nestin, Sox1, and NeuroD; cardiac: Nkx2-5 and Isl1; and hematopoietic: Gata1 and Hba-x) were downregulated in both SSEA1+ cells and heterozygous ES cells (Figure 1F). SSEA1+ cells likely have a wide range of potential, since EBs generated from these cells expressed markers for all three germ layers (Figure 1G).

Figure 2. Prkci and Pluripotency Pathways (A) ERK1/2 phosphorylation (Y202/Y204) is reduced in null ES cells and early day (d)-6 null EBs compared to heterozygous EBs and strongly increased at later stages. The first lane shows ES cells activated (A) by serum treatment 1 day after serum depletion. (B) Quantification of pERK1/2 normalized to non-phosphorylated ERK1/2 (three independent experiments; mean ± SEM; **p < 0.01). (C) pERK1/2 Y202/Y204 is strongly expressed in the columnar epithelium of heterozygous EBs that have just cavitated. Null EBs have lower expression. OCT4 and pERK1/2 expression do not co-localize. Scale bar, 100 mm. (D) pERK1/2Y202/Y204 levels are lower in null SSEA1+ sorted cells than in heterozygous or in null day-12 EBs that have undergone further differentiation. pSTAT3 and STAT levels are unchanged. See also Figure S2.

ERK1/2 Signaling during EB Development Stem cell self-renewal has been shown to require the activation of the JAK/STAT3 and PI3K/AKT pathways and the inhibition of ERK1/2 and GSK3 pathways (Kunath et al., 2007; Niwa et al., 1998; Sato et al., 2004; Watanabe et al., 2006). We found that both STAT3 and phosphorylated STAT3 levels were not grossly altered and that the p-STAT3/STAT3 ratio was similar between heterozygous and null ES cells and EBs (Figures S2A and S2B). In addition we did not see any difference in AKT, pAKT, or b-CATENIN levels when comparing heterozygous to null ES cells or EBs (Figures S2A and S2C). Thus, the effects observed by the loss of Prkci are unlikely to be due to a significant alteration in the JAK/STAT3, PI3K/AKT, or GSK3 pathways.

Next, we investigated ERK1/2 expression and activation. Consistent with other studies showing ERK1/2 activation to be downstream of Prkci in some mammalian cell types (Boeckeler et al., 2010; Litherland et al., 2010), pERK1/2 was markedly inactivated in Prkci null versus heterozygous ES cells. In addition, during differentiation, null EBs displayed strong pERK1/2 inhibition early (until day 6). Later, pERK1/2 was activated strongly, as the EB began differentiating (Figures 2A and 2B). By immunofluorescence, pERK1/2 was strongly enriched in the columnar epithelium of control EBs, while overall levels were much lower in Prkci/ EBs (Figure 2C). In addition, high OCT4 expression correlated with a marked inactivation of pERK1/2 (Figure 2C). Next, we examined Prkci/ SSEA1+ cells by western blot. We found that SSEA1+ cells isolated from day-12 null EBs had pSTAT3 expression levels similar to whole EBs, while pERK1/2 levels were low (Figure 2D). Thus, these experiments indicate that the higher numbers of pluripotent cells in null EBs correlate with a strong inactivation of ERK1/2.

Neural Stem Cell Fate Is Favored in Prkci/ EBs It is well known that ERK/MEK inhibition is not sufficient for pluripotent stem cell maintenance (Ying et al., 2008); thus, other pathways are likely involved. Therefore, we used a TaqMan Mouse Stem Cell Pluripotency Panel (#4385363) on an OpenArray platform to investigate the mechanism of Prkci action. Day 13 and day 20 Prkci/ EBs expressed high levels of pluripotency and stemness markers versus heterozygous EBs, including Oct4, Utf1, Nodal, Xist, Fgf4, Gal, Lefty1, and Lefty2. However, interestingly, EBs also expressed markers for differentiated cell types and tissue stem cells, including Sst, Syp, and Sycp3 (neural-related genes), Isl1 (cardiac progenitor marker), Hba-x, and Cd34 (hematopoietic markers). Based on this first-pass test, we sought to determine whether loss of Prkci might favor the generation of neural, cardiac, and hematopoietic cell types and/or their progenitors.

Figure 3. Neural Stem Cell Populations Are Increased in Null EBs (A–C0 ) Prkci/ EBs (B) have more NESTINpositive cells than Prkci+/ EBs (A). (C and C0 ) MAP2 and TUJ1 are expressed in null EBs, similarly to heterozygous EBs (data not shown). (D) EBs were assessed for PAX6 expression, and the images were used for quantification (Figures S3A and S3B). The pixel count ratio of PAX6+ cells in null EBs (green) is substantially higher than that found in heterozygous EBs (black) (three independent experiments; mean ± SEM; *p < 0.05). (E–F000) Day 4 after RA treatment, Prkci/ EBs have more NESTIN- than TUJ1-positive neurons (E and F). However, null cells can still terminally differentiate into NEUROD-, NEUN-, and MAP2-positive cells (F0 –F000). Scale bars, 25 mm in (A and C) and 50 mm in (E). See also Figure S3. Ste

The Generation of Cardiomyocyte and Erythrocyte Progenitors Is Also Favored Next, we examined ISL1 expression (a cardiac stem cell marker) by immunofluorescence and found that Prkci/ EBs contained larger ISL1 clusters compared with Prkci+/ EBs; this was confirmed using an image quantification assay (Figures 4A, 4A0 , and 4C). Differentiated cardiac cells and ventral spinal neurons can also express ISL1 (Ericson et al., 1992); therefore, we also examined Nkx2-5 expression, a better stem cell marker and regulator of cardiac progenitor determination (Brown et al., 2004), by RT-PCR and immunofluorescence. In null EBs, Nkx2-5 was upregulated (Figure 4D). In addition, in response to RA, which can promote cardiac fates in vitro (Niebruegge et al., 2008), cells expressing NKX2-5 were more prevalent in null versus heterozygous EBs (Figures 4B and 4B0 ).The abundant cardiac progenitors found in null EBs were still capable of undergoing differentiation (Figures 4E–4F0 ).

Figure 4. Cardiomyocyte and Erythrocyte Progenitors Are Increased in Prkci/ EBs (A–F0 ) In (A, A0 , E, and E0 ), Prkci/ EBs cultured without LIF have more ISL1 (cardiac progenitor marker) and a-ACTININ-positive cells compared to heterozygous EBs. (C) At day (d) 9, the pixel count ratio for ISL1 expression indicates that null EBs (green) have larger ISL1 populations than heterozygous EBs (black) (three independent experiments, n = 20 heterozygous EBs, 21 null EBs total; mean ± SEM; *p < 0.05). In (B, B0 , D, F, and F0 ), RA treatment induces more NKX2-5 (both nuclear and cytoplasmic) and a-ACTININ expression in null EBs. Arrows point to fibers in (F0 ). (G) Null EBs (green) generate more beating EBs with RA treatment compared to heterozygous EBs (black) (four independent experiments; mean ± SEM; *p < 0.05, ***p < 0.001). (H) Dissociated null EBs of different stages (green) generate more erythrocytes in a colony-forming assay (CFU-E) (four independent experiments; mean ± SEM; **p < 0.01). (I) Examples of red colonies. (J) Gene expression for primitive HSC markers is upregulated in null EBs (relative to heterozygous EBs) (three independent experiments; mean ± SEM). Scale bars, 50 mm in (A, B, and E); 100 mm in (F), and 25 mm in (I). See also Figure S4. 6

Hba-x expression is restricted to yolk sac blood islands and primitive erythrocyte populations (Lux et al., 2008; Trimborn et al., 1999). Cd34 is also a primitive HSC marker (Sutherland et al., 1992). Next, we determined whether the elevated expression of these markers observed with OpenArray might represent higher numbers of primitive hematopoietic progenitors. Using a colony-forming assay (Baum et al., 1992), we found that red colonies (indicative of erythrocyte differentiation; examples in Figure 4I) were produced significantly earlier and more readily from cells isolated from null versus heterozygous EBs (Figure 4H). By quantitative real-time PCR, upregulation of Hba-x and Cd34 genes confirmed the OpenArray results (Figure 4J). In addition, we found Gata1, an erythropoiesis-specific factor, and Epor, an erythropoietin receptor that mediates erythroid cell proliferation and differentiation (Chiba et al., 1991), to be highly upregulated in null versus heterozygous EBs (Figure 4J). These data suggest that the loss of Prkci promotes the generation of primitive erythroid progenitors that can differentiate into erythrocytes.

To determine whether the aforementioned tissue stem cells identified were represented in the OCT4+ population that we described earlier, we examined the expression of PAX6, ISL1, and OCT4 in adjacent EB sections. We found that cells expressing OCT4 appeared to represent a distinct population from those expressing PAX6 and ISL1 (although some cells were PAX6 and ISL1 double-positive) (Figures S4A–S4C).

Prkci/ Cells Are More Likely to Inherit NUMB/aNOTCH1 Symmetrically The enhanced production of both pluripotent and tissue stem cells suggests that the mechanism underlying the action of Prkci in these different contexts is fundamentally similar. Because the Notch pathway controls stem cell self-renewal in many contexts (Hori et al., 2013), and because previous studies implicated a connection between PRKCi function and the Notch pathway (Bultje et al., 2009; Smith et al., 2007), we examined the localization and activation of a key player in the Notch pathway, NUMB, (Inaba and Yamashita, 2012). Differences in NUMB expression were first evident in whole EBs, where polarized expression was evident in the ectodermal and endodermal epithelia of heterozygous EBs, while Prkci/ EBs exhibited a more even distribution (Figures 5A–5B0 ). To more definitively determine the inheritance of NUMB during cell division, doublets undergoing telophase or cytokinesis were scored for symmetric (evenly distributed in both cells) or asymmetric (unequally distributed) NUMB localization (examples: Figures 5C and 5C0 ).

Because NUMB can be directly phosphorylated by aPKCs (both PRKCi and PRKCz) (Smith et al., 2007; Zhou et al., 2011), loss of Prkci might be expected to lead to decreased NUMB phosphorylation. Three NUMB phosphorylation sites—Ser7, Ser276, and Ser295—could be aPKC mediated (Smith et al., 2007). By immunofluorescence, we found that one of the most well-characterized sites (Ser276), was strongly inactivated in null versus heterozygous EBs, especially in the core (Figures 5F and 5G). Western analysis also confirmed that the levels of pNUMB (Ser276) were decreased in null versus heterozygous EBs (Figure S5F). Thus, genetic inactivation of Prkci leads to a marked decrease in the phosphorylation status of NUMB.

Notch pathway inhibition by NUMB has been observed in flies and mammals (Berdnik et al., 2002; French et al., 2002). Therefore, we investigated whether reduced Numb activity in Prkci/ EBs might lead to enhanced NOTCH1 activity and the upregulation of the downstream transcriptional readouts (Meier-Stiegen et al., 2010). An overall increase in NOTCH1 activation was supported by western blot analysis showing that the level of activated NOTCH1 (aNOTCH1) was strongly increased in day 6 and day 10 null versus heterozygous EBs (Figure S5G). This was supported by immunofluorescence in EBs, where widespread strong expression of aNOTCH1 was seen in most null cells (Figures 5I and 5I0 ), while in heterozygous EBs, this pattern was observed only in the OCT4+ cells (Figures 5H and 5H0 ).

Figure 5. Prkci/ Cells Preferentially Inherit Symmetric Localization of NUMB and aNOTCH1 and Notch Signaling Is Required for Stem Cell Self-Renewal in Null Cells (A–B0 ) In (A and B), day (d)-7 heterozygous EBs have polarized NUMB localization within epithelia and strong expression in the endoderm, while null EBs have a more even distribution. (A0 and B0 ) Enlarged views. (C and C0 ) Asymmetric and symmetric NUMB expression examples. (D) Doublets from day-10 null EBs have more symmetric inheritance when compared to day-10 heterozygous doublets (three independent experiments; mean ± SEM; **p < 0.01). A red line indicates a ratio of 1 (equal percent symmetric and asymmetric). (E) CD24high null doublets exhibited more symmetric NUMB inheritance than CD24high heterozygous doublets (three independent experiments; mean ± SEM; *p < 0.05). A red line indicates where the ratio is 1. (F and G) Decreased pNUMB (Ser276) is evident in the core of null versus heterozygous EBs (n = 10 of each genotype). (H–I0 ) In (H and I), aNOTCH1 is strongly expressed in heterozygous EBs, including both OCT4+ and OCT4 cells, while strong aNOTCH1 expression is predominant in OCT4+ cells of null EBs (n = 10 of each genotype)). (H0 and I0 ) Enlarged views of boxed regions. OCT4+ cells are demarcated with dotted lines. (J and J0 ) OCT4+ cells express HES5 strongly in the nucleus (three independent experiments). (K) Null doublets from dissociated EBs have more symmetric aNOTCH1 inheritance compared to heterozygous doublets (three independent experiments; mean ± SEM; **p < 0.01). A red line indicates where the ratio is 1. (L) CD24high Prkci/ doublets exhibit more symmetric aNOTCH1 than CD24high heterozygous doublets (three independent experiments; mean ± SEM; *p < 0.05). A red line indicates where the ratio is 1. (M and M0 ) Examples of asymmetric and symmetric aNOTCH1 localization. (N and O) Day-3 DMSO-treated null ES colonies show strong AP staining all the way to the colony edge in (N). Treatment with 3 mM DAPT led to more differentiation in (O). (P–R) OCT4 is strongly expressed in day-4 DMSO-treated null ES cultures (P). With DAPT (Q,R), OCT4 expression is decreased. (S) Working model: In daughter cells that undergo differentiation, PRKCi can associate with PAR3 and PAR6. NUMB is recruited and directly phosphorylated. The activation of NUMB then leads to an inhibition in NOTCH1 activation and stimulation of a differentiation/maintenance program. In the absence of Prkci, the PAR3/PAR6 complex cannot assemble (although it may do so minimally with Prkcz). NUMB asymmetric localization and phosphorylation is reduced. Low levels of pNUMB are not sufficient to block NOTCH1 activation, and activated NOTCH1 preserves the stem cell self-renewal program. We suggest that PRKCi functions to drive differentiation by pushing the switch from an expansion phase that is symmetric to a differentiation and/or maintenance phase that is predominantly asymmetric. In situations of low or absent PRKCi, we propose that the expansion phase is prolonged. Scale bars, 50 mm in (A, B, F, G, H, I, J, J0 , P–R); 200 mm in (A0 and B0 ); 25 mm in (C, C0 , M, and M0 ); and 100 mm in (H0 , I0 , N, and O). See also Figure S5.

Figure 6. Additional Inhibition of PRKCz Results in an Even Higher Percentage of OCT4-, SSEA1-, and STELLA-Positive Cells (A and A0 ) After day 4 without LIF, heterozygous ES cells undergo differentiation in the presence of Go¨6983, while null ES cells stay as distinct colonies in (A0 ). (B and B0 ) Go¨6983 stimulates an increase in OCT4+ populations in heterozygous EBs and an even larger OCT4+ population in null EBs in (B0 , insets: green and red channels separately). (C–D0 ) An even higher percentage of cells are OCT4+ (C and C0 ) and SSEA1+ (D and D0 ) with Go¨6983 treatment (day 12, three independent experiments). (E and F) More STELLA+ clusters containing a larger number of cells are present in drugtreated heterozygous EBs. (G and H) Null EBs also have more STELLA+ clusters and cells. Drug-treated null EBs exhibit a dramatic increase in the number of STELLA+ cells. (I–K) Some cells are double positive for STELLA and VASA in drug-treated null EBs (yellow arrows). There are also VASAonly (green arrows) and STELLA-only cells (red arrows) (three independent experiments). (L–P) Treatment with ZIP results in an increase in OCT4+ and STELLA+ cells. ZIP treatment also results in more cells that are VASA+ (three independent experiments); n = 11 for Prkci+/, and n = 13 for Prkci+/ + ZIP; n = 14 for Prkci/, and n = 20 for Prkci/ + ZIP; eight EBs assayed for both STELLA and VASA expression). Scale bars, 100 mm in (A and A0 ); 50 mm in (B and B0 ); and 25 mm in (E, I, and L).

DISCUSSION In this report, we suggest that Prkci controls the balance between stem cell expansion and differentiation/maintenance by regulating the activation of NUMB, NOTCH1, and Hes /Hey downstream effector genes. In the absence of Prkci, the pluripotent cell fate is favored, even without LIF, yet cells still retain a broad capacity to differentiate. In addition, loss of Prkci results in enhanced generation of tissue progenitors such as neural stem cells and cardiomyocyte and erythrocyte progenitors. In contrast to recent findings on Prkcz (Dutta et al., 2011), loss of Prkci does not appear to influence STAT3, AKT, or GSK3 signaling but results in decreased ERK1/2 activation. We hypothesize that, in the absence of Prkci, although ERK1/2 inhibition may be involved, it is the decreased NUMB phosphorylation and increased NOTCH1 activation that promotes stem and progenitor cell fate. Thus, we conclude that PRKCi, a protein known to be required for cell polarity, also plays an essential role in controlling stem cell fate and generation via regulating NOTCH1 activation.

Notch Activation Drives the Decision to Self-Renew versus Differentiate Notch plays an important role in balancing stem cell selfrenewal and differentiation in a variety of stem cell types and may be one of the key downstream effectors of Prkci signaling. Sustained Notch1 activity in embryonic neural progenitors has been shown to maintain their undifferentiated state (Jadhav et al., 2006). Similarly, sustained constitutive activation of NOTCH1 stimulates the proliferation of immature cardiomyocytes in the rat myocardium (Collesi et al., 2008). In HSCs, overexpression of constitutively active NOTCH1 in hematopoietic progenitors and stem cells supports both primitive and definitive HSC selfrenewal (Stier et al., 2002). Together, these studies suggest that activation and/or sustained Notch signaling can lead to an increase in certain tissue stem cell populations. Thus, a working model for how tissue stem cell populations are favored in the absence of Prkci involves a sequence of events that ultimately leads to Notch activation. Recent studies have shown that aPKCs can be found in a complex with NUMB in both Drosophila and mammalian cells (Smith et al., 2007; Zhou et al., 2011); hence, in our working model (Figure 5S), we propose that the localization and phosphorylation of NUMB is highly dependent on the activity of PRKCi. When Prkci is downregulated or absent (as shown here), cell polarity is not promoted, leading to diffuse distribution and decreased phosphorylation of NUMB. Without active NUMB, NOTCH1 activation is enhanced, Hes/Hey genes are upregulated, and stem/progenitor fate generation is favored. To initiate differentiation, polarization could be stochastically determined but could also be dependent on external cues such as the presentation of certain ligands or extracellular matrix (ECM) proteins (Habib et al., 2013). When PRKCi is active and the cell becomes polarized, a trimeric complex is formed with PRKCi, PAR3, and PAR6. Numb is then recruited and phosphorylated, leading to Notch inactivation, the repression of downstream Hes/Hey genes, and differentiation is favored (see Figure 5S). Support for this working model comes from studies in Drosophila showing that the aPKC complex is essential for Numb activation and asymmetric localization (Knoblich, 2008; Smith et al., 2007; Wang et al., 2006). Additional studies on mouse neural progenitors show that regulating Numb localization and Notch activation is critical for maintaining the proper number of stem/progenitor cells in balance with differentiation (Bultje et al., 2009). Thus, an important function for PRKCi may be to regulate the switch between symmetric expansion of stem/progenitor cells to an asymmetric differentiation/maintenance phase. In situations of low or absent PRKCi, we propose that the expansion phase is favored. Thus, temporarily blocking either, or both, of the aPKC isozymes may be a powerful approach for expanding specific stem/progenitor populations for use in basic research or for therapeutic applications.

Although we do not see changes in the activation status of the STAT3, AKT, or GSK3 pathway, loss of Prkci results in an inhibition of ERK1/2 (Figures 2A and 2B). This result is consistent with the findings that ERK1/2 inhibition is both correlated with and directly increases ES cell selfrenewal (Burdon et al., 1999). Modulation of ERK1/2 activity by Prkci has been observed in cancer cells and chondrocytes (Litherland et al., 2010; Murray et al., 2011). Although it is not clear whether a direct interaction exists between Prkci and ERK1/2, Prkcz directly interacts with ERK1/2 in the mouse liver and in hypoxia-exposed cells (Das et al., 2008; Peng et al., 2008). The Prkcz isozyme is still expressed in Prkci null cells but evidently cannot suf- ficiently compensate and activate the pathway normally. Furthermore, knocking down Prkcz function in ES cells does not result in ERK1/2 inhibition, suggesting that this isozyme does not impact ERK1/2 signaling in ES cells (Dutta et al., 2011). Therefore, although PRKCi may interact with ERK1/2 and be directly required for its activation, ERK1/2 inhibition could also be a readout for cells that are more stem-like. Further studies will be needed to address this question.

Utility of Inhibiting aPKC Function Loss of Prkci resulted in EBs that contained slightly more STELLA+ cells than EBs made from +/ cells. Furthermore, inhibition of both aPKC isozymes by treating Prkci null cells with the PKC inhibitor Go¨6983 or the more specific inhibitor, ZIP, strongly promoted the generation of large clusters of STELLA+ and VASA+ cells, suggesting that inhibition of both isozymes is important for PGC progenitor expansion (Figure 6). It is unclear what the mechanism for this might be; however, one possibility is that blocking both aPKCs is necessary to promote NOTCH1 activation in PGCs or in PGC progenitor cells that may ordinarily have strong inhibitions to expansion (Feng et al., 2014). Regardless of mechanism, the ability to generate PGC-like cells in culture is notoriously challenging, and our results provide a method for future studies on PGC specification and differentiation. Expansion of stem/progenitor pools may not be desirable in the context of cancer. Prkci has been characterized as a human oncogene, a useful prognostic cancer marker, and a therapeutic target for cancer treatment. Overexpression of Prkci is found in epithelial cancers (Fields and Regala, 2007), and Prkci inhibitors are being evaluated as candidate cancer therapies (Atwood et al., 2013; Mansfield et al., 2013). However, because our results show that Prkci inhibition leads to enhanced stem cell production in vitro, Prkci inhibitor treatment as a cancer therapy might lead to unintended consequences (tumor overgrowth), depending on the context and treatment regimen. Thus, extending our findings to human stem and cancer stem cells is needed.

In summary, here, we demonstrate that loss of Prkci leads to the generation of abundant pluripotent cells, even under differentiation conditions. In addition, we show that tissue stem cells such as neural stem cells, primitive erythrocytes, and cardiomyocyte progenitors can also be abundantly produced in the absence of Prkci. These increases in stem cell production correlate with decreased NUMB activation and symmetric NUMB localization and require Notch signaling. Further inhibition of Prkcz may have an additive effect and can enhance the production of PGC-like cells. Thus, Prkci (along with Prkcz) may play key roles in stem cell self-renewal and differentiation by regulating the Notch pathway. Furthermore, inhibition of Prkci and or Prkcz activity with specific small-molecule inhibitors might be a powerful method to boost stem cell production in the context of injury or disease.


CRISPR Gene Editing Trial

Larry H Bernstein, MD, FCAP, Curator



CRISPR Gene Editing to Be Tested on People by 2017


A biotechnology company says it will test advanced gene-engineering methods to treat blindness.

By Antonio Regalado on November 5, 2015

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Malaria Resistant Mosquitos by Design

Larry H. Bernstein, MD, FCAP, Curator



CRISPR-Powered Malaria Mosquito Gene Drive

Using the precision gene-editing tool, researchers demonstrate an ability to create large populations of malaria parasite–resistant mosquitoes.

By Tracy Vence | November 24, 2015

Using CRISPR, investigators at the Universities of California (UC) in San Diego and Irvine have engineered transgenic Anopheles stephensimosquitoes carrying an anti-malaria parasite effector gene “capable of introgressing the genes throughout wild vector populations,” they wrote in a PNAS paper published this week (November 23). The resulting gene-drive system could help wipe out the malaria pathogen (Plasmodium falciparum) within a targeted population of A. stephensi vectors, Anthony James of UC Irvine and his colleagues wrote.

“We know the gene works,” James said in a statement. “The mosquitoes we created are not the final brand, but we know this technology allows us to efficiently create large populations.”

As Nature noted, this study is not the first to report engineered Anopheles that stifle the malaria parasite but, until now, “researchers lacked a way to ensure that the resistance genes would spread rapidly through a wild population.” CRISPR/Cas9 gene-editing enabled this feat. “Males and females derived from transgenic females . . . produce progeny with a high frequency of mutations in the targeted genome sequence, resulting in near-Mendelian inheritance ratios of the transgene,” James and his colleagues wrote in their paper. (See “Reining in Gene Drives,” The Scientist, November 2015.)

“This work suggests that we’re a hop, skip, and jump away from actual gene-drive candidates for eventual release,” Kevin Esvelt of the Wyss Institute who was not involved in the work told Nature. “This is a major advance because it shows that gene drives will likely be effective in mosquitoes,” Esvelt told MIT Technology Review. “Technology is no longer the limitation.”

In the UC Irvine statement, study coauthor Ethan Bier of UC San Diego added that “the ability of this system to carry large genetic payloads should have broad applications to the future use of related CRISPR-based ‘active genetic’ systems.”

Reining in Gene Drives

Researchers have developed two methods to avoid the unchecked spread of engineered genes through wild populations.

By Karen Zusi | November 18, 2015

“Gene drive” is a phenomenon that causes a gene to be inherited at a rate faster than Mendelian principles would dictate. It relies on genes that can copy themselves onto a corresponding location in a paired chromosome, thereby overriding typical allele inheritance patterns. In conjunction with CRISPR/Cas9, gene drives can be created with almost any DNA sequence, raising questions about the risk of engineered genes spreading quickly through a population. But a team of researchers from Harvard University published a study this week (November 16) in Nature Biology that offers some safety constraints on the system.

A description of the first CRISPR/Cas9 gene drive system was published in March by a team at the University of California, San Diego, and showed rapid spreading of a normally recessive phenotype inDrosophila. Other labs are researching the system’s potential to wipe out insect-borne diseases such as malaria by spreading mutated genes throughout a mosquito population. But the strategy carries the risk of accidental contamination of wild populations.

“We have a responsibility to keep our experiments confined to the laboratory,” Kevin Esvelt, an evolutionary engineer and coauthor on the paper, told Nature. “The basic lesson is: if you don’t have to build a gene drive that can spread through a wild population, then don’t.”


‘Gene drive’ mosquitoes engineered to fight malaria

Mutant mozzies could rapidly spread through wild populations.

Heidi Ledford & Ewen Callaway!/image/1.18858%20.jpg_gen/derivatives/landscape_630/1.18858%20.jpg

The Anopheles stephensi mosquito can spread the malaria parasite to humans.

Mutant mosquitoes engineered to resist the parasite that causes malaria could wipe out the disease in some regions — for good.

Humans contract malaria from mosquitoes that are infected by parasites from the genusPlasmodium. Previous work had shown that mosquitoes could be engineered to rebuff the parasiteP. falciparum1, but researchers lacked a way to ensure that the resistance genes would spread rapidly through a wild population.

In work published on 23 November in the Proceedings of the National Academy of Sciences, researchers used a controversial method called ‘gene drive’ to ensure that an engineered mosquito would pass on its new resistance genes to nearly all of its offspring2 — not just half, as would normally be the case.

The result: a gene that could spread through a wild population like wildfire.

“This work suggests that we’re a hop, skip and jump away from actual gene-drive candidates for eventual release,” says Kevin Esvelt, an evolutionary engineer at Harvard University in Cambridge, Massachusetts, who studies gene drive in yeast and nematodes.

For Anthony James, a molecular biologist at the University of California, Irvine, and an author of the paper, such a release would spell the end of a 30-year quest to use mozzie genetics to squash malaria.

James and his laboratory have painstakingly built up the molecular tools to reach this goal. They have worked out techniques for creating transgenic mosquitoes — a notoriously challenging endeavour — and isolated genes that could confer resistance to P. falciparum. But James lacked a way to ensure that those genes would take hold in a wild population.

Fast forward

The concept of engineering a gene drive has been around for about a decade, and James’s laboratory had tried to produce them in the past. The process was agonizingly slow.

Then, in January, developmental biologists Ethan Bier and Valentino Gantz at the University of California, San Diego, contacted James with a stunning finding: they had engineered a gene drive in fruit flies, and wondered whether the same system might work in mosquitoes. James jumped at the opportunity to find out.

Bier and Gantz had used a gene-editing system called CRISPR–Cas9 to engineer a gene drive. They inserted genes encoding the components of the system that were designed to insert a specific mutation in their fruit flies. The CRISPR–Cas9 system then copied that mutation from one chromosome to the other3. James used that system in mosquitoes to introduce two genes that his past work showed would cause resistance to the malaria pathogen.

The resulting mosquitoes passed on the modified genes to more than 99% of their offspring. Although the researchers stopped short of confirming that all the insects were resistant to the parasite, they did show that the offspring expressed the genes.

“It’s a very significant development,” says Kenneth Oye, a political scientist who studies emerging technologies at the Massachusetts Institute of Technology in Cambridge. “Things are moving rapidly in this field.”

Other teams are developing gene drives that could control malaria. A team at Imperial College London has developed a CRISPR-based gene drive in Anopheles gambiae, the mosquito species that transmits malaria in sub-Saharan Africa. The group’s gene drive inactivates genes involved in egg production in female mosquitoes, which could be used to reduce mosquito populations, according to team member Austin Burt, an evolutionary geneticist. Their results will be published inNature Biotechnology next month, Burt says.

Oye notes that such technological advances are outpacing the regulatory and policy discussionsthat surround the use of gene drive to engineer wild populations. Gene drives are controversial because of the potential that they hold for altering entire ecosystems.

Before testing gene drive in the field, Oye hopes that researchers will study the long-term consequences of the changes, such as their stability and potential to spread to other species, as well as methods to control them. “I’m less worried about malevolence than getting something wrong,” he says.

Esvelt says that the US-based researchers made a wise decision in selecting a non-native mosquito species for their experiments. (The team worked with Anopheles stephensi, which is native to the Indian subcontinent.) “Even if they escaped the lab, there’d be no one to mate with and spread the drive,” Esvelt says.

James predicts that it will take his team less than a year to prepare mosquitoes that would be suitable for field tests, but he is in no rush to release them. “It’s not going to go anywhere until the social science advances to the point where we can handle it,” he says. “We’re not about to do anything foolish.”



  1. Isaacs, A. T. et al. PLoS Pathog. 7, e1002017 (2011).
  2. Gantz, V. M. et al. Proc. Natl Acad. Sci. USA




With This Genetic Engineering Technology, There’s No Turning Back

Designers of a “selfish” gene able to spread among mosquitoes say it could wipe out malaria, but the scientific community is at odds over whether or not we should do it.

The students in Anthony James’s basement insectary at the University of California, Irvine, knew they’d broken the laws of evolution when they looked at the mosquitoes’ eyes.

By rights, the bugs, born from fathers with fluorescent red eyes and mothers with normal ones, should have come out only about half red. Instead, as they counted them, first a few and then by the hundreds, they found 99 percent had glowing eyes.

More important than the eye color is that James’s mosquitoes also carry genes that stop the malaria parasite from growing. If these insects were ever released in the wild, their “selfish” genetic cargo would spread inexorably through mosquito populations, and potentially stop the transmission of malaria.

The technology, called a “gene drive,” was built using the gene-editing technology known as CRISPR and is being reported by James, a specialist in mosquito biology, and a half dozen colleagues today in the Proceedings of the National Academy of Sciences.

A functioning gene drive in mosquitoes has been anticipated for more than a decade by public health organizations as a revolutionary novel way to fight malaria. Now that it’s a reality, however, the work raises questions over whether the technology is safe enough to ever be released into the wild.

“This is a major advance because it shows that gene drives will likely be effective in mosquitoes,” says Kevin Esvelt, a gene drive researcher at Harvard University’s Wyss Institute. “Technology is no longer the limitation.”


Highly efficient Cas9-mediated gene drive for population modification of the malaria vector mosquito Anopheles stephensi


Malaria continues to impose enormous health and economic burdens on the developing world. Novel technologies proposed to reduce the impact of the disease include the introgression of parasite-resistance genes into mosquito populations, thereby modifying the ability of the vector to transmit the pathogens. Such genes have been developed for the human malaria parasite Plasmodium falciparum. Here we provide evidence for a highly efficient gene-drive system that can spread these antimalarial genes into a target vector population. This system exploits the nuclease activity and target-site specificity of the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) system, which, when restricted to the germ line, copies a genetic element from one chromosome to its homolog with ≥98% efficiency while maintaining the transcriptional activity of the genes being introgressed.

Genetic engineering technologies can be used both to create transgenic mosquitoes carrying antipathogen effector genes targeting human malaria parasites and to generate gene-drive systems capable of introgressing the genes throughout wild vector populations. We developed a highly effective autonomous Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-associated protein 9 (Cas9)-mediated gene-drive system in the Asian malaria vector Anopheles stephensi, adapted from the mutagenic chain reaction (MCR). This specific system results in progeny of males and females derived from transgenic males exhibiting a high frequency of germ-line gene conversion consistent with homology-directed repair (HDR). This system copies an ∼17-kb construct from its site of insertion to its homologous chromosome in a faithful, site-specific manner. Dual anti-Plasmodium falciparum effector genes, a marker gene, and the autonomous gene-drive components are introgressed into ∼99.5% of the progeny following outcrosses of transgenic lines to wild-type mosquitoes. The effector genes remain transcriptionally inducible upon blood feeding. In contrast to the efficient conversion in individuals expressing Cas9 only in the germ line, males and females derived from transgenic females, which are expected to have drive component molecules in the egg, produce progeny with a high frequency of mutations in the targeted genome sequence, resulting in near-Mendelian inheritance ratios of the transgene. Such mutant alleles result presumably from nonhomologous end-joining (NHEJ) events before the segregation of somatic and germ-line lineages early in development. These data support the design of this system to be active strictly within the germ line. Strains based on this technology could sustain control and elimination as part of the malaria eradication agenda.

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CRISPR Gene Editing Trial

Larry H Bernstein, MD, FCAP, Curator



CRISPR Gene Editing to Be Tested on People by 2017


A biotechnology company says it will test advanced gene-engineering methods to treat blindness.

By Antonio Regalado on November 5, 2015


Editas CEO Katrine Bosley

The biotechnology startup Editas Medicine intends to begin tests of a powerful new form of gene repair in humans within two years.

Speaking this week at the EmTech conference in Cambridge, Massachusetts, Editas CEO Katrine Bosley said the company hopes to start a clinical trial in 2017 to treat a rare form of blindness using CRISPR, a groundbreaking gene-editing technology.

If Editas’s plans move forward, the study would likely be the first to use CRISPR to edit the DNA of a person.

CRISPR technology was invented just three years ago but is so precise and cheap to use it has quickly spread through biology laboratories. Already, scientists have used it to generate genetically engineered monkeys, and the technique has stirred debate over whether modified humans are next (see “Engineering the Perfect Baby”).

Editas is one of several startups, including Intellia Therapeutics and CRISPR Therapeutics, that have plans to use the technique to correct DNA disorders that affect children and adults. Bosley said that because CRISPR can “repair broken genes” it holds promise for treating several thousand inherited disorders caused by gene mistakes, most of which, like Huntington’s disease and cystic fibrosis, have no cure.

Editas, which had not previously given a timeline for an initial human test of CRISPR, will try to treat one form of a rare eye disease called Leber congenital amaurosis, which affects the light-receiving cells of the retina.

The condition Editas is targeting affects only about 600 people in the U.S., says Jean Bennet, director of advanced retinal and ocular therapeutics at the University of Pennsylvania’s medical school. “The target that they have selected is fantastic; it has all the right characteristics in terms of making a correction easily,” says Bennett, who isn’t involved in the Editas study.

Children with LCA are born seeing only large, bright shapes, and infants are diagnosed when they don’t look into their mother’s eyes or react to colorful balloons. Their poor vision can progress to “stone cold blindness where everything is black,” says Bennett.

Editas picked the disease in part because it is relatively easy to address with CRISPR, Bosley said. The exact gene error is known, and the eye is easy to reach with genetic treatments. “It feels fast, but we are going at the pace science allows,” she said. There are still questions about how well gene-editing will work in the retina and whether side effects could be caused by unintentional changes to DNA.

Editas plans to deliver the CRISPR technology as a gene therapy. The treatment will involve injecting into the retina a soup of viruses loaded with the DNA instructions needed to manufacture the components of CRISPR, including a protein that can cut a gene at a precise location. Bosley said in order to treat LCA, the company intends to delete about 1,000 DNA letters from a gene called CEP290 in a patient’s photoreceptor cells.

After the edit, preliminary lab experiments show, the gene should function correctly again. Bosley said Editas still needs to test the approach further in the lab and in animals before a human study starts.

Editas was created by venture capital funds including Third Rock Ventures in 2013 and was cofounded by scientists including Feng Zhang of the MIT/Harvard Broad Institute. It has raised more than $160 million in capital, allowing it to pursue ideas for several treatments simultaneously, Bosley said.

Although the Editas study could be the first for CRISPR in humans, it wouldn’t be the first clinical study of gene editing. An older method called zinc fingers is already in testing to treat HIV infection by the biotechnology company Sangamo Biosciences. But the versatility and ease with which CRISPR can change DNA means it may outpace earlier approaches.

Theoretically, gene editing could be used to repair broken genes in any part of the body. But in practice it is difficult to make DNA repairs in most cell types, such as brain cells. The eye is an exception because doctors can inject treatment directly under the retina.

There is already a gene-therapy treatment for one form of LCA in late-stage clinical testing by Philadelphia biotech Spark Therapeutics, says Bennett, who helped develop that treatment. In that case, an entire, healthy version of a gene is being added to eye cells. Typically, gene therapy can only add genes, not edit them.

LCA has several different genetic causes, and standard gene therapy won’t work for the form of the disease Editas is looking at. That is because the required gene, CEP290, is too big to fit inside a virus, says Bennett, and so there is no easy way to add it.

By targeting an exceptionally rare illness, Editas may have an easier time getting a treatment tested and approved. However, the eventual cost of such a treatment could be extraordinarily high, given the small number of people who would need it. Bennett says only around 3,000 Americans have LCA, and about 20 percent of those have the form being targeted by Editas.

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