2 international meeting genome editing & gene modulation...

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G E N O M E E D I T I N G & G E N E M O D U L A T I O N 2 0 1 6 | 0 6t h

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A P R I L 2 0 1 6 | O X F O R D | U K Email: [email protected] | Web: http://lpmhealthcare.com/gegm-2016 | Twitter: @LPMHealthcare | Hashtag: #GEGM16

2nd International Meeting

GENOME EDITING & GENE MODULATION CONGRESS | 2016 F u n c t i o n a l G e n o m i c s T e c h n o l o g i e s f o r T r a n s l a t i o n a l R e s e a r c h & T h e r a p e u t i c s

06TH-08TH APRIL 2016 The Edward Boyle Auditorium, St Hilda’s College, Cowley Place Oxford OX4 1DY, United Kingdom

Email: [email protected] Web: http://lpmhealthcare.com/gegm-2016 Twitter: @LPMHealthcare | Hashtag: #GEGM16

TABLE OF CONTENTS

2: General Information 2: Information for Presenters 3: Insurance and Liability 3: Disclaimer 3: Map 4: Podium Agenda 6: Podium Abstracts 18: Poster Abstracts 24: Sponsors and Supporters

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GENERAL INFORMATION Event Dates: 06

th-08

th April 2016

Event Website: http://lpmhealthcare.com/gegm-2016

Venue: The Edward Boyle auditorium, The Jacqueline du Pré Music Building, St Hilda’s College,

Cowley Place, Oxford, OX4 1DY, England, UK

Tel: +44 (0) 1865 276884, Website: http://www.st-hildas.ox.ac.uk

Registration Desk: Located in JdP foyer

Name Badges: The College requests all delegates to wear name badges while on the premises to avoid any

confusion.

Refreshments/Lunch: In JdP foyer

Mobile Phones: As a courtesy to speakers and participants, please switch off your mobile phone during oral

presentations.

Speaker Presentations: We will not be distributing speaker presentations. Therefore, if you interested in

presentation slides of any speakers, please get in touch with them directly.

Internet access: Please use edurom if you can. Otherwise, WiFi Code and instructions for internet access via

your laptop/mobile device can be obtained at the time of registration.

Health and Safety: Please do not leave your belongings unattended or in passageways and familiarise yourself

with emergency exits.

Smoking: In addition to any local venue regulations, UK no-smoking regulations apply on the College

premises.

INFORMATION FOR PRESENTERS

SPEAKERS:

• Presentation standard will be data projection from a central PC. Your presentation is best brought in a memory stick.

• Macintosh will not be available. Therefore, if you are a Macintosh user please bring your own.

• As a courtesy to other speakers and attendees please finish your talk absolutely within your allocated time slot. (Guide: For a 20 minute talk, prepare 12-14 slides maximum ; for a 30 minute talk, prepare 20-22 slides maximum; allow 3-4 minutes for questions). Please check the agenda below for your presentation schedule.

POSTER DISPLAY:

• Please leave your poster at the registration desk when you register.

• There is no specific poster session. Instead the posters will be displayed in the registration/refreshment area for full duration of the meeting.

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INSURANCE AND LIABILITY: Participants are responsible for taking appropriate insurance cover (including health insurance) in connection with their attendance of this event. The event organisers and hosts are not responsible for personal accidents, any travel costs, or the loss of private property, and will not be liable for any claims. Event participants shall be responsible for compensating any loss, should they cause any damage to the host’s property or the venue.

DISCLAIMER: The information specified in oral and poster presentations, written abstracts, biographies and exhibitions come from diverse sources and it is not in the capacity of event organisers to validate it, and is provided on an ‘as-is’ basis. Responsibility for the literary and scientific content of the abstracts and the presentations, both oral and poster, remains with the authors and the presenters. Therefore, the event organisers accept no responsibility for literary or scientific correctness of this information, and shall accept no liability of any kind, should any of the information be incorrect. The event organisers and hosts make no representation or warranty of gain of business or profits as a result of use of services or information provided in connection with the even and shall not be liable for any direct or indirect damages, loss of business, employment, profits or anticipated savings resulting from the use of the services or information provided in connection with the event, in any country or court of law. Furthermore, the materials contained in the event handbook are provided on the understanding that speakers or presenters have the right to their presentation in this manner. Therefore, event organisers and hosts shall not be liable for infringement of third party rights by an event presenter, participant, sponsor, supporter or exhibitor.

This handbook is for use by the Genome Editing & Genome Modulation Congress 2016 Oxford (06th

-

08th

April 2016) participants only. © 2016 Copyright Information: Textual and graphical contents of this handbook are copyright of presenters, sponsors,

instructors and/or LibPubMedia Ltd, unless explicitly stated otherwise. No part of this handbook may be reproduced, distributed

or transmitted in any form or by any means, electronic or mechanical, including but not limited to, photocopy, recording, or any

other information storage or retrieval system, without the prior written permission of the legal copyright owners.

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PODIUM AGENDA (subject to change) Wednesday 06

th April 2016 | The Edward Boyle auditorium, The Jacqueline du Pré Music Building, St Hilda’s College

All talks will be held in the The Edward Boyle auditorium, The Jacqueline du Pré Music Building, St Hilda’s College.

13.00: Registration, welcome coffee, networking and exhibition

13.55: Welcome and housekeeping

Session 1: Chair Dr Gregory Davis

14.00: Opening KEYNOTE: Professor Emmanuelle Charpentier, Department Head, Helmholtz Centre for Infection Research, Germany

14.40: Dr Mark Behlke, Chief Scientific Officer, Integrated DNA Technologies, USA Improved CRISPR editing using chemically-modified crRNA:tracrRNA complexes

15.10: Dr Annaleen Vermeulen, Senior Scientist, Dharmacon/GE Healthcare, Lafayette, CO, USA Utilizing synthetic RNA for CRISPR-Cas9 arrayed screening

15.40: Refreshment break, networking, exhibition and posters

16.10: Dr Eric Paul Bennett, Copenhagen Center for Glycomics (CCG), University of Copenhagen, Denmark

“Hit and seek”: Improved strategies for identifying the breaks in a precisely broken genome

16.40: Dr Stephen Hague, European Droplet Digital PCR Specialist, Bio-Rad Laboratories, UK Ultra-sensitive Quantification of Genome Editing Events by Droplet Digital PCR (ddPCR)

17.10: Dr Kyle Luttgeharm, Application Specialist, Advanced Analytical Technologies Inc. Oak Tree Ct, Ankeny, IA, USA

Development of a high throughput screening protocol for rapid identification of mutated alleles

17.30: Dr Cornelia Hampe, Scientific Support Specialist, Takara Bio Europe, France Gene editing with high efficiency and no additional footprint using novel CRISPR/Cas9 Gesicle technology

17.50: Technology Workshop 1 by Merck KGaA

18.20: Close of Day 1

Thursday 07th

April 2016 | The Edward Boyle auditorium, The Jacqueline du Pré Music Building, St Hilda’s College

08.45: Welcome

Session 2: Chair Dr Mark Behlke

08.50: Dr Ben Davies, Transgenic Core Head and Group Leader, WTCHG, University of Oxford, UK

Efficient CRISPR/Cas9 genome engineering using embryos derived from Cas9 overexpressing transgenic mice

09.20: KEYNOTE: Dr Gregory Davis, Senior R&D Manager, MilliporeSigma, USA Genome and Epigenome Modification with ZFNs and CRISPR/Cas Systems

10.00: Dr Chady Jaber, Thermo Fisher Scientific, USA Improved delivery of Cas9 protein/gRNA complexes using lipofectamine CRISPRMAX


11.00: Dr Philip Webber, Patent Attorney, Dehns Patent and Trademark Attorneys, Oxford, UK CRISPR – the Patent Wars

11.20: Dr Barry Rosen, Senior Principal Scientist, Discovery Sciences, AstraZeneca, UK Precise Genome Editing and Stem Cell Technologies-Novel Tools for Novel Medicines

11.50: Technology Workshop 2 by Merck KGaA

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12.20: Technology Workshop 3 by Dr Barry Rosen

13.00: Lunch, networking, exhibition and posters

Session 3: Chair Dr Barry Rosen

14.00: KEYNOTE: Dr William C Skarnes, Senior Group Leader, The Wellcome Trust Sanger Institute, UK Biallelic genome editing of human iPSCs at scale

14.45: Professor Richard Wade-Martins, The Oxford Parkinson’s Disease Centre, University of Oxford, UK Deciphering the molecular basis of neurodegeneration using human iPSC neurons

15.15: Dr Robin Ketteler, Group Leader MRC LMB and Manager TRRC, University College London, UK Applications of Genome Editing and RNAi to Dissect Autophagy Signaling Pathways

15.45: Dr Christine Seidl, Research Associate, The Kennedy Institute of Rheumatology, University of Oxford, UK Investigating functional microRNA target sites by CRISPR/Cas9 genome editing


17.00: Dr Tony Nolan, Senior Research Fellow, Imperial College London, UK A CRISPR-based gene drive system to suppress populations of malarial mosquitoes

17.30: Dr Michal Minczuk, Group Leader, Mitochondrial Genetics, MRC Mitochondrial Biology Unit, Cambridge, UK Manipulating the human mitochondrial genome with designer nucleases

18.00: Dr Aleksandar Vojta, Associate Professor, University of Zagreb, Faculty of Science, Zagreb, Croatia Targeted CpG methylation using the modified CRISPR-Cas9 system

18.30: Close of Day 2

19.15: Networking Dinner (by prior booking or invitation only)

Friday 08th

April 2016 | The Edward Boyle auditorium, The Jacqueline du Pré Music Building, St Hilda’s College

08.55: Welcome to Day 3

Session 4: Chair TBA

09.00: Dr Lydia Teboul, Group Leader, MRC Harwell, UK CRISPR-aided mutagenesis of the mouse genome

09.30: Dr Emmanouil Metzakopian, Wellcome Trust Sanger Institute, Hinxton, Cambridge, UK Human and Mouse Genome Wide CRISPR-guide RNA Arrayed Libraries

10.00: Dr Fanny Decarpentrie, James TURNER group, The Francis Crick Institute, London, UK New tool for new discoveries: CRISPR/Cas9 genome editing in marsupials


11.00: Dr Cedric Ghevaert, Senior Lecturer, University of Cambridge/Cambridge Blood Centre, UK Platelet production in vitro for transfusion from human pluripotent stem cells: the added power of genome editing

11.30: Miss Cristina Fimiani, Graduate Researcher, SISSA, Trieste, Italy RNA-therapeutics of gene haploinsufficiencies

11.50: Dr Hamid Dolatshad, Research Associate, RDM-Clinical Laboratory Sciences, John Radcliffe Hospital, Oxford, UK Use of CRISPR/CAS9 to investigate the function of gene mutations found in myeloid malignancies

12.20: Discussion and Close

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PODIUM ABSTRACTS Title currently unavailable

Emmanuelle Charpentier – Keynote

Max-Plank Institute, Germany

Abstract currently unavailable

Improved CRISPR editing using chemically-modified crRNA:tracrRNA complexes

Michael A. Collingwood, Ashley M. Jacobi, Mollie S. Schubert, Garrett R. Rettig, and Mark A. Behlke

Integrated DNA Technologies, Inc. Coralville, IA, 52241 USA

The CRISPR/Cas9 system for gene editing is a powerful tool for manipulating mammalian genomes. The natural system in

S. pyogenes employs two RNA molecules, a 42-nt target-specific CRISPR RNA (crRNA) and an 89-nt universal trans-

activating RNA (tracrRNA). We systematically truncated these RNAs and developed optimized versions that show

improved performance in mammalian cells (36-nt crRNA and 67-nt tracrRNA). Although unmodified RNA oligonucleotides

can be used to direct Cas9 cleavage, they are rapidly degraded by serum or cellular nucleases, limiting their functional

activity. Further, unmodified RNAs can trigger an innate immune response in mammalian cells. Extensive studies of

chemical modification strategies for both the crRNA and the tracrRNA were performed, where over 400 RNA oligos were

compared for functional performance in various settings, systematically testing position-specific tolerance for base

modification. Highly functional modified variants were developed where as high as 78% of the crRNA and 84% of the

tracrRNA residues were substituted with 2’OMe RNA. Use of phosphorothioate modified internucleotide linkages and/or

other end-blocking strategies were also beneficial to prevent 5’- or 3’-exonuclease attack. The new length-optimized

chemically-modified crRNA:trRNA synthetic oligonucleotides can be annealed, complexed with recombinant Cas9 protein

and introduced into mammalian cells using lipofection or electroporation to achieve high editing efficiency with minimal

side effects. Good results have been obtained in a variety of systems including: Zebrafish, C. elegans, mammalian tissue

culture cells, iPSCs, mouse embryos, and primary T-cells isolated from human donors.

Utilizing synthetic RNA for CRISPR-Cas9 arrayed screening

Annaleen Vermeulen, Melissa L. Kelley, Emily M. Anderson, Shawn McClelland, Elena Maksimova, Tyler Reed, Steve

Lenger, Žaklina Strezoska, Hidevaldo Machado, Eldon Chou, John Schiel and Anja van Brabant Smith

Dharmacon, part of GE Healthcare, 2650 Crescent Drive, Suite 100, Lafayette, CO 80026, United States of America

The CRISPR-Cas9 system permits researchers to generate precise sequence changes in mammalian, fish and plant

genomes, among others, and consequently has dramatically transformed biological research. Furthermore, this system

has provided a new avenue for functional genomics by empowering gene knockout screens. Pooled lentiviral sgRNA

libraries have been used for screening thousands of genes in parallel. However, there are limitations to this platform,

including the requirement for a phenotypic assay that allows for enrichment or depletion in a cell population and

deconvolution by next-generation sequencing. Arrayed screening, which enables one-gene-per-well knockout, enables a

wider variety of biological assays, such as endpoint assays and high content imaging. Here we describe the generation of

predesigned synthetic crRNA libraries for screening with an emphasis on algorithm development for functional gene

knockout and reduction of off-targeting. In addition, we will demonstrate the use of synthetic crRNA libraries in several

biological assays.

"Hit and seek”: Improved strategies for identifying the breaks in a precisely broken

genome

Eric P. Bennett

Copenhagen Center for Glycomics, Department of Odontology, Faculty of Health Sciences, University of Copenhagen

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The emerging ZFN, TALEN and CRISPR gene editing tools for precise engineering of higher eukaryote genomes have

revolutionized bioscience. In contrast to the speed by which these editing tools are being optimized and strategies for

high throughput use in whole-genome screens are devised, considerable less focus are being devoted to improving

capabilities for targeting, detection and characterization of the induced insertions and/or deletions (indels) at the specific

breakpoint as well as at potential off-targets. Current indel identification approaches include: i) enzyme mismatch

cleavage (EMC) assays and ii) next generation DNA sequencing, which is costly, time and labor intensive and poorly suited

for high throughput screening. Here, we report a novel strategy that combines use of a simple amplicon labelling strategy

with the high throughput capability of DNA fragment analysis by automated Capillary Electrophoresis for simple detection

and characterization of indels induced by precise gene targeting. The strategy is coined IDAA for Indel Detection by

Amplicon Analysis (Z.Yang et al., NAR, 2015) and we demonstrate that IDAA is suitable for detecting indels in both cell

pools with low efficiency targeting and single sorted cells. Furthermore, we show that IDAA is ideally suited for high

throughput detection of indels down single base events, estimation/validation of gRNA design “cutting” efficiencies,

evaluation of off-target events at candidate loci and thus, is ideally suited for CRISPR/Cas9 genome editing surveillance.

Used in combination with a versatile method for increasing, in particular CRISPR/Cas9 editing efficiencies, by fused co-

expression of nuclease and fluorescent protein in the same cell (K.Duda et al., NAR, 2014), we propose a powerful

workflow that translates into significantly higher genome editing frequencies.

Ultra-Sensitive Quantification of Genome Editing Events by Droplet Digital PCR

(ddPCR)

Stephen Hague

Droplet Digital PCR Specialist, Europe, Bio Rad Laboratories, UK

Droplet Digital PCR (ddPCR) allows with unrivalled precision the direct quantification of target DNA or RNA molecules,

counting target copies per microliter of sample. The technology relies on partitioning nucleic acid into droplets,

subsequent amplification and then directly counting positive vs negative droplets using the QX 200 droplet reader system.

The ratio of positive to negative droplets is used to calculate the concentration of the sample. ddPCR has proved to be

particularly powerful when used to quantify Rare events/mutations in a variety of sample types. Recently, Bio Rad has

developed a novel approach using common hydrolysis probe chemistry to quantify both Homology Directed Repair (HDR)

and Non Homologous end joining events (NHEJ). The advantage of ddPCR over existing techniques for genome editing

validation is that it is both time and cost effective, allowing the rapid screening of up to 96 pooled clones in one 96 well

plate reaction. This allows researchers to rapidly screen clones to identify pools harbouring the correct editing event and

further quantify both HDR and NHEJ events at frequencies less than 0.04% and 0.1 % respectively. Currently, Bio Rad

Laboratories are developing a pipeline for the design and manufacture of assays specifically for Genome editing events.

This pipeline will have the capacity to design custom assays for the detection and quantification of both HDR and NHEJ

events and where possible design them such that both events can be detected and quantified in one PCR. We believe the

application of ddPCR coupled to a specific design algorithm will enable the sensitive and rapid screening of genome

editing events to accelerate genome editing protocols.

Development of a high throughput screening protocol for rapid identification of

mutated alleles

Kyle Luttgeharm, Kit-Sum Wong, Danilo Tait, Steve Siembieda

Advanced Analytical Technologies Inc. 2450 SE Oak Tree Ct, Ankeny, IA, USA, 50021

CRISPR gene editing is rapidly advancing as the technology becomes more economical and efficient. However, protocols

to identify CRISPR mutations and determine the natures of these mutations are time consuming and often involve costly

sequencing steps. To overcome this limitation, many researchers are turning to heteroduplexing and enzymatic

mismatch cleavage assays to rapidly screen for mutated lines. However, few studies have been done to compare the

efficiencies of different cleavage enzymes (T7E1 and Surveyor for example), on different types of mutations and lengths

of PCR products. Using Advanced Analytical Technologies Inc. Fragment Analyzer™ and synthetic genes to mimic different

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CRISPR mutations, we are developing an optimized protocol to identify a wide variety of CRISPR mutations, from PCR to

fragment analysis. Of particular interest is determining if a single enzyme can cleave SNPs, additions, and deletions with

high efficiency. Furthermore, it may be possible to determine whether a mutation is mono- or di-allelic by comparing the

ratio of uncut to cut DNA using statistical models that predict heteroduplexing probability. Having a single high-

throughput protocol that results in cleavage of multiple types of mutations while also determining the number of

mutated alleles would allow for efficient screening of CRISPR mutant populations at a level currently not feasible.

Gene editing with high efficiency and no additional footprint, using novel

CRISPR/Cas9 Gesicle technology

Cornelia Hampe

Scientific Support Specialist and Product Manager for Genome Engineering Tools, Takara Bio Europe, France

While CRISPR/Cas9 is a powerful technique for gene editing, two significant challenges remain: obtaining efficient delivery

of Cas9 and gene-specific sgRNA to all cell types, and leaving no additional footprint (i.e., persistent and elevated

expression of Cas9 in target cells) that could lead to off-target effects. To address these challenges, we have developed

cell-derived nanovesicles called gesicles. Gesicles contain active Cas9 protein complexed with a target-specific sgRNA.

Gesicles are engineered with glycoproteins on their surface that mediate binding and fusion with the membranes of a

wide range of target cells, allowing delivery of Cas9/sgRNA even to difficult-to-transfect and non-dividing cells. These

features enable gesicles to knock out genes with high efficiency and in a broader range of cell types than plasmid-based

delivery methods. Furthermore, Cas9 protein delivery prevents genomic integration and persistent expression of Cas9,

thus reducing off-target effects.

Efficient CRISPR/Cas9 genome engineering using embryos derived from Cas9

overexpressing transgenic mice

Alberto Cebrian-Serrano, Shijun Zha, Lars Hanssen, Daniel Biggs, Christopher Preece and Ben Davies

Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford OX3 7BN, UK

Genome manipulation in the mouse via microinjection of CRISPR/Cas9 site specific nucleases has allowed the production

time for mouse model development to be significantly reduced and the technique is now being widely adopted as the

method of choice. For laboratories establishing these techniques, the exact mode of delivery of the CRISPR/Cas9

components needs to be optimized. Successful genome manipulation in the mouse has already been reported using Cas9

supplied by microinjection of a DNA construct, purified in vitro transcribed mRNA and recombinant protein. As an

alternative to these options, we have investigated the feasibility of supplying Cas9 genetically and for this purpose have

generated transgenic mice which overexpress Cas9 ubiquitously, through the use of a CAG-Cas9 transgene, targeted to

the ROSA26 locus. Heterozygous and homozygous transgenic mice are indistinguishable from wild-type litter mates and

show normal fertility, suggesting that ubiquitous Cas9 expression in the absence of an activating gRNA is inert.

Microinjection of fertilized zygotes prepared from transgenic Cas9 overexpressing females with guide-RNAs resulted in

high numbers of mutant mice and whole embryo analysis revealed that the level of mutagenesis was found to be

significantly higher when Cas9 was supplied genetically relative to exogenous supply. Genetic supply of Cas9 was able to

mediate loss-of-function alleles by indel mutation, point mutation changes by homology directed repair and small

deletions using two gRNAs.

Genome and Epigenome Modification with ZFNs and CRISPR/Cas Systems

Gregory Davis – Keynote

Senior R&D Manager, Molecular Biotechnology, MilliporeSigma, USA

Zinc finger nucleases and CRISPR/Cas systems have radically changed questions which can be posed in experimental

biology. Mouse genetic models mimicking human disease have been a dominating technology in past decades and can

now be verified by CRISPR modification of genes in human cells or in other organisms (rats, pigs, etc.) which may better

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model certain human disease phenotypes. Despite huge success in creating gene knockouts in a wide array of organisms,

the ability to edit genomes to have user-defined genome modifications is still largely limited by the potential of various

cell types to copy synthetic DNA via homologous recombination (HR). Various cell treatments and donor DNA formats to

address HR limitations will be discussed. The simplicity and low cost of programming CRISPR-Cas systems has created

possibilities for high throughput, genome wide interrogation of gene function. For example, a drug treatment of cells can

be characterized by knocking out tens of thousands of genes in parallel to identify pathways that lead to drug resistance.

While this mode of genome wide genetic interrogation has been available via RNAi and transposons for years, CRISPR

enables are very controlled and effective method of eliminating human gene function. Various new applications of

CRISPR screening will be discussed. Beyond editing the genome at the base pair level, new formats have been developed

which enable editing of the epigenome. In this talk, a new epi-CRISPR technology will be discussed which allows

acetylation of histones to open up chromatin landscapes to allow local gene expression and/or association of enhancers

with distant regulatory elements.

Improved delivery of Cas9 protein/gRNA complexes using lipofectamine CRISPRMAX

Xin Yu, Xiquan Liang, Huimin Xie, Shantanu Kumar, Namritha Ravinder, Jason Potter, Xavier de Mollerat du Jeu, Jonathan

D Chesnut and Chady Jaber

Synthetic Biology Department, Thermo Fisher Scientific, 5781 Van Allen Way, Carlsbad, CA 92008, USA

Objectives: To identify the best lipid nanoparticles for delivery of purified Cas9 protein and gRNA complexes (Cas9 RNPs)

into mammalian cells and to establish the optimal conditions for transfection. Results: Using a systematic approach, we

screened 60 transfection reagents using six commonly-used mammalian cell lines and identified a novel transfection

reagent (named Lipofectamine CRISPRMAX). Based on statistical analysis, the genome modification efficiencies in

Lipofectamine CRISPRMAX-transfected cell lines were 40 or 15 % higher than those in Lipofectamine 3000 or RNAiMAX-

transfected cell lines, respectively. Upon optimization of transfection conditions, we observed 85, 75 or 55 % genome

editing efficiencies in HEK293FT cells, mouse ES cells, or human iPSCs, respectively. Furthermore, we were able to co-

deliver donor DNA with Cas9 RNPs into a disrupted EmGFP stable cell line, resulting in the generation of up to 17 %

EmGFP-positive cells. Conclusion: Lipofectamine CRISPRMAX was characterized as the best lipid nanoparticles for the

delivery of Cas9 RNPs into a variety of mammalian cell lines, including mouse ES cells and iPSCs.

CRISPR – the Patent Wars

Philip Webber

Patent Attorney, Dehns Patent and Trademark Attorneys, Oxford, UK

Patents which cover the basic CRISPR technology have now been granted to Zhang’s group in the US and in Europe, and

these patents are already being licensed by a number of companies and institutions. However, Doudna’s group filed

patent applications earlier than Zhang, but Doudna’s patent applications have not yet been granted. The stage is

therefore set for battles in the US and European Patent Offices as they both fight for the patent rights in this

revolutionary technology. This presentation will first discuss what inventions may be protected in the area of gene

editing/modulation, and then provide an update on the Zhang-Doudna patent wars.

Precise Genome Editing and Stem Cell Technologies-Novel Tools for Novel Medicines

Barry Rosen

Senior Principal Scientist (Director), Discovery Sciences, AstraZeneca Cambridge, UK

Recent advances in Precise Genome Editing(PGE), particularly in the application of CRISPR/Cas9 technology, and

improvements in methods for human stem cell generation and differentiation into disease relevant cell types have the

potential to revolutionize the drug discovery process at all stages from target identification/validation to toxicological

studies, empowering the faster development of drugs which are more likely to be safe and efficacious in humans. The

ability to generate physiologically relevant human cells of defined genetic backgrounds in almost limitless supply and

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engineer them fluently with PGE to create both specific disease models and various endogenous reporter alleles makes

human induced pluripotent stem cells (iPSC) an attractive alternative to the currently used recombinant cell lines or

primary cells in all segments of the drug discovery process. This presentation will describe the use of CRISPR/Cas9 PGE

technology and human iPSC based disease models for target identification/validation at AstraZeneca, highlighting internal

case studies. Furthermore, our external strategic partnerships towards genome-wide target discovery and validation

studies employing whole genome CRISPR libraries will also be described. The potential transformative impact of

CRISPR/Cas9 PGE technologies and human iPSC technology across our drug development pipeline will be stressed.

Biallelic genome editing of human iPSCs at scale

Manousos Koutsourakis, Wendy Bushell and William C. Skarnes

Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, UK

The advent of site-specific nucleases and improved conditions for human iPSC culture now permits efficient engineering

of human stem cells. CRISPR-Cas9 technology, in particular, provides a facile tool for the generation of a range of alleles

in human stem cells with little risk of off-target damage. We established a high-throughput pipeline for the generation of

homozygous knockout human iPSCs. We construct short arm targeting vectors and sgRNA expression plasmids in 96-well

format. Following co-transfection of the targeting vector with Cas9 and sgRNA expression plasmids, we screen for clones

where one allele is targeted by homologous recombination and the second allele is damaged by non-homologous end

joining. Bi-allelic knockout of genes is observed in 10-30% of the colonies screened. Our method lends itself to high-

throughput genotyping: biallelic events are identified by Sanger sequencing of the non-targeted allele. Our aim is to

generate and distribute arrays of human iPS cell knockouts that will be coupled to focused phenotyping screens in

cultured cells. Currently, we are developing a vector-free method using Cas9 ribonucleoprotein (Cas9 RNP) and single

strand oligonucleotides for fluent generation of biallelic point mutations and revertants for disease modelling.

Deciphering the molecular basis of neurodegeneration using human iPSC neurons

Richard Wade-Martins

Oxford Parkinson's Disease Centre (OPDC), Department of Physiology, Anatomy and Genetics, University of Oxford, UK

Parkinson's disease (PD) is the second most common neurodegenerative disease and a major unmet clinical need in our

ageing population. The focus of the Oxford Parkinson's Disease Centre (OPDC; www.opdc.ox.ac.uk) is to exploit the

interdisciplinary research environment within Oxford to establish a leading centre focused on translational research

understanding the earliest pathological pathways in PD. We have built a new multi-disciplinary research program across

the translational space comprising the OPDC Discovery Cohort as one of the best-characterised clinical Parkinson’s cohorts

in the world, a core expertise in molecular genetics and the largest induced pluripotent stem-cell (iPSC) research program

in Parkinson’s in Europe. We have developed a range of genetic models with the aim of studying disease processes in

human iPSC-derived dopamine neurons for phenotypic screens and target discovery in Parkinson’s. The OPDC has

generated ~150 iPSC lines from ~50 individuals comprising sporadic PD patients, patients carrying mutations in the leucine

rich repeat kinase 2 (LRRK2), glucocerebrosidase (GBA) and alpha-synuclein (SNCA) genes and controls. Mature

dopaminergic neurons with correct morphology express essential protein markers, exhibit key neurophysiological features

and reveal robust neurobiological deficits in PD lines. In depth phenotypic analysis comparing iPSC-derived dopamine

neurons from patients carrying the GBA-N370S mutation with controls revealed elevated ER stress, autophagic

perturbations and increased release extracellular α-synuclein into the extracellular medium. The ability to study in depth

such previously inaccessible neurons will allow detailed dissection of genetic mechanisms and identify molecular

therapeutic targets.

Applications of Genome Editing and RNAi to Dissect Autophagy Signaling Pathways

Joana Costa, Alexander Agrotis, Niccolo Pengo, Jacob Heintze and Robin Ketteler

MRC Laboratory for Molecular Cell Biology, University College London, London, WC1E 6BT, UK

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The recent development of clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9 for experimental

purposes has added a new dimension to functional genomics. While traditional loss-of-function approaches rely on

siRNA-mediated reduction in transcript levels, it is now possible to screen for knockout phentoypes. Libraries with whole-

genome target coverage have been generated that can be used in pooled screening approaches using positive or negative

selection readouts. For high-content screening assays, the use of arrayed shRNA libraries would be advantageous as

phenotypes are often qualitative and not linked to survival of a cell population. Multiple companies and academic labs are

now developing arrayed gRNA libraries for the use in arrayed library screening. We have used two approaches for arrayed

library screening utilizing the CRISPR/Cas9 technology. One uses viral stocks of lentiviral gRNA expression vectors

targeting human kinases. These constructs have been transduced into target cells and assessed for their effect on cell

viability and autophagy. We have compared these results with the use of a siRNA library targeting the human kinome and

find significant differences in phenotypes. This may be related to differences in off-target effects and the efficiency of

target gene modulation. In addition, we have taken another approach using genome modified knockout cell panels

generated by CRISPR/Cas9. These cells are homogenous in phenotype and sequence-verified. Using these cell panels, we

find that some previously observed siRNA phenotypes can be recapitulated in genome edited knockout cell panels.

Overall, we provide evidence that arrayed library screening is feasible and can complement current siRNA approaches.

Investigating functional microRNA target sites by CRISPR/Cas9 genome editing

Christine I. Seid, and Chris L. Murphy

The Kennedy Institute of Rheumatology, University of Oxford, Roosevelt Drive, Oxford OX3 7FY, UK

During the last decade, the importance of non-coding RNAs (ncRNAs) in regulation of the cellular transcriptome has

drawn the attention of scientific research towards elucidation of these mechanisms. A very important class of short

ncRNA, microRNAs (miRs), have been shown to profoundly influence cellular mRNA levels by targeting mRNAs via

sequence-complementarity over a short, 7-8 nt long sequence, predominantly found in the 3’UTR of their targets and

resulting in downregulation of the mRNA and subsequently reduction of the corresponding protein. The power of miRs in

transcriptome regulation has raised interest to use these ncRNAs in therapeutic approaches. However, due to the

property of a miR having multiple mRNA targets, clinical approaches have been hampered due to the possible detrimental

side effects that a global change of a certain miR may have for the individual. Frequently, a single mRNA displays more

than one potential target site for a given miR but often only one site is functionally relevant. Thus, if therapeutic

approaches in the context of miRs are to be fruitfully developed, the possibility of changing the specifically relevant target

site on the affected mRNA, rather than perturbing the miR level itself, would be of paramount importance. In the current

presentation, I will discuss an approach to determine the effect of single-nucleotide target-site mutations by genome

editing using CRISPR/Cas9 using the example of primary human articular chondrocytes (HACs). HACs display severe

limitations for traditional transfection protocols and different means of generating editing events will be explained. The

ability to determine and edit the most dominant site that results in a change of mRNA expression will allow us to once

again pursue a therapeutic miR-based approach.

A CRISPR-based gene drive system to suppress populations of malarial mosquitoes

Tony Nolan

Senior Research Fellow, Infection and Immunity, Department of Life Sciences, Imperial College London, London, UK

Synthetic gene drive systems using site-specific endonucleases to spread traits into a population, at rates much faster

than simple Mendelian genetics, were first proposed more than a decade ago. Such gene drive systems have huge

potential for transforming populations of insect pests and disease vectors in ways that are beneficial for human health

and over meaningful timeframes. The advent of CRISPR-Cas9 has brought this possibility closer to fruition due to the

adaptability and activity of this enzyme in being engineered to recognise virtually any sequence in a wide range of

organisms. We have engineered a gene drive system designed to cause population suppression in the principle mosquito

vector of malaria by using CRISPR-based driving constructs to target and disrupt genes with confirmed roles in female

fertility. We see close to 100% transmission rates of the CRISPR drive alleles in each generation (instead of the 50%

expected from Mendelian inheritance) and a rapid increase in frequency over subsequent mosquito generations in a

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caged experiment leading to a drastic decrease in the reproductive output. These results provide the basis for the

development of a gene drive system that has the potential to substantially reduce mosquito populations to levels that

would not support malaria transmission. Moreover, our approach is broadly applicable to a range of invasive pests and

vectors of disease.

Manipulating the human mitochondrial genome with designer nucleases

Michal Minczuk

Mitochondrial Genetics, MRC Mitochondrial Biology Unit, Cambridge, UK

Human mitochondria contain a small, multi-copy DNA genome (mtDNA), from which thirteen proteins and all RNAs

necessary for their expression are produced. All proteins encoded by human mtDNA are essential subunits of the

oxidative phosphorylation system. Mutations in protein or RNA coding genes and regulatory elements of mtDNA often

have severe biochemical consequences, resulting in mitochondrial disease. In most cases, mutant and wild-type mtDNAs

coexist within a single cell, resulting in heteroplasmy. The selective elimination of mutant mtDNA, and consequent

enrichment of wild-type mtDNA, can rescue pathological phenotypes in patent-derived heteroplasmic cells. In our work

we have developed mitochondrially targeted zinc finger-nucleases (mtZFNs) for degradation of mutant mtDNA through

site-specific DNA cleavage. We have successfully used mtZFNs to target and cleave mtDNA harbouring disease-associated

point mutations or large-scale deletions. More recently, we achieved a near-complete elimination of mutant mtDNA by

dynamic dose-controlled treatment with mtZFNs, which limits off-target catalysis and undesired mtDNA copy number

depletion. Our current efforts focus on the use mtZFNs in order to manipulate mtDNA heteroplasmy in animal models of

mtDNA disease. To conclude, our research provides proof-of-principle that mtZFN-based approaches offer means for

mtDNA heteroplasmy manipulation in basic research, and may provide a strategy for therapeutic intervention in selected

mitochondrial diseases.

Targeted CpG methylation using the modified CRISPR-Cas9 system

Aleksandar Vojta, Paula Dobrinić, Vanja Tadić, Luka Bočkor, Marija Klasić, Petra Korać and Vlatka Zoldoš

Department of Biology, Division of Molecular Biology, University of Zagreb, Faculty of Science, Zagreb, Croatia

We have repurposed the CRISPR-Cas9 system for DNA methylation at targeted CpG sites. This novel tool enables

epigenetic silencing of gene expression by methylation of CpG sites in regulatory regions of targeted genes. The reversible

nature of epigenetic modifications, including DNA methylation, has been already exploited in cancer therapy for

remodelling the aberrant epigenetic landscape. However, the classical approach uses epigenetic inhibitors non-

selectively. In contrast to that, epigenetic editing at specific sites in the genome could selectively alter the gene

expression pattern. The novel tool for specific DNA methylation that we developed consists of deactivated Cas9 (dCas9)

nuclease fused to the catalytic domain of the DNA methyltransferase DNMT3A. It is targeted to any 20 bp sequence

followed by the NGG trinucleotide by co-expression of a guide RNA. We demonstrated CpG methylation by the fusion

protein in a ~35 bp wide region. We also showed that multiple guide RNAs could target the dCas9-DNMT3A construct to

multiple sites, which enabled methylation of a wider genome region. Alternatively, multiple targeting could be used to

silence several genes simultaneously. DNA methylation activity was highly specific for the targeted region and heritable

across mitotic divisions. Finally, we demonstrated that directed DNA methylation of the wider region in the promoter of

the target locus IL6ST decreased its expression, which served as a proof of the concept of artificial epigenetic silencing by

targeted CpG methylation in vivo.

Designing for success: the right CRISPR design strategies for the right experiment

Victor Dillard, Leigh Brody, Neil Humphryes, Riley Doyle

Desktop Genetics, 3P1 Cooper House, 2 Michael Road, London SW6 2AD, UK

The importance of selecting high activity sgRNAs is most emphasized when designing genome-scale CRISPR libraries

targeting thousands of genes, as selecting guides with high, and comparable, activity, can minimize bias and improve

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screen readouts. However, CRISPR design is a complex multi-variate problem. The weight of each variable differs

depending on the experiment under consideration. Current methods of designing CRISPR experiments are based on a

single-variable, resulting in poor experimental outcomes. We have developed an integrated bioinformatics software

platform to provide a single tool for designing sgRNA guides, which considers a wide range of design variables, including

focusing on minimizing off-target activity while achieving high activity at the target locus. Our current scoring methods

were tested and validated using an sgRNA library of over 3,500 sgRNAs in essential genes with a variety of predicted

sgRNA activities and specificities. From this experiment, we also investigated current design ‘best-practices’ and were

unable to corroborate some currently accepted sgRNA selection criteria, which demonstrates the need to constantly

challenge and improve sgRNA design rules. By using the right design rules for the right experiment, common modes of

failure can avoided. These rules have been integrated into the free DESKGEN web platform, now used by over 2,000

researchers worldwide. The platform is continuously maintained up to date with the latest CRISPR developments,

ensuring the CRISPR research community benefits from the latest science, always. From the numerous experiments

enabled by the DESKGEN platform to date, we can demonstrate key design improvements to maximise experimental

outcome.

CRISPR-aided mutagenesis of the mouse genome

Joffrey Mianné, Adam Caulder, Gemma Codner, Ruairidh King, Rachel Fell, Wendy Gardiner, Martin Fray, Sara Wells and

Lydia Teboul

The Mary Lyon Centre, MRC Harwell, HSIC, Oxon, OX11 0RD, UK

Mouse models are valuable tools to understand genes functions, genetic diseases and to develop and test new

therapeutic treatments in vivo. The ability to introduce tailored modifications within the mouse genome is essential to

generate models for the study of human diseases. The recently developed CRISPR/Cas system as genome engineering tool

has brought new perspectives for the generation of mouse models in a more efficient and precise fashion, at reduced

price, all within a shorter time scale. We will present the use of the CRISPR/Cas9 technology at the Mary Lyon Centre,

MRC Harwell to introduce a wide range of modifications into the mouse genome through different methods. We will first

present our high throughput mouse production pipeline allowing us to generate alleles containing indels, tailored

deletions or point mutations through direct injection into zygotes. We will detail the parameters we tested and how we

optimised the process to increase the efficiency and the diversity of allele generated via this pipeline, as well as to ensure

their quality. We will then present our data obtained for enhancing the homologous recombination rate in mouse

embryonic stem cell through co-electroporation of IKMC targeting vector and CRISPR reagents. Finally, we will introduce a

new pilot consisting of a new CRISPR reagents delivery strategy through in vivo testis electroporation. Developing these

methods for genome engineering will enable the generation of a wide range of increasingly complex alleles in mice, both

in a custom and high throughput context.

Human and Mouse Genome Wide CRISPR-guide RNA Arrayed Libraries

Emmanouil Metzakopian

Career Development Fellow, Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, UK

CRISPR technology has rapidly improved biological research, such that accurate genome editing has now become

common practice for many labs within a few years of its initial development. CRISPR-Cas9 has also proven to be a useful

tool with great potential in genome wide genetic screens. Here we present the first genome wide, arrayed guide RNA

screening libraries for CRISPR-Cas9. These libraries have been designed to cover 17166 human and 20430 mouse genes.

Two gRNAs are designed per gene giving a total of 34332 gRNAs for human and 40860 gRNAs for mouse genomes,

respectively. These gRNAs have been cloned in a lentivirus backbone containing piggyBac transposase recognition

elements for additional flexibility in generating stable cell lines. To test the efficiency of these libraries, we selected gRNAs

targeting genes from the GPI anchor protein pathway. We observed that over 95% of these gRNAs successfully induced

DNA cleavage and consequently gene loss of function. These libraries offer the prospect for performing high-throughput

screens for genes of considerable scientific and therapeutic value in mouse and human cells.

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New tool for new discoveries: CRISPR/Cas9 genome editing in marsupials

Fanny DECARPENTRIE

James Turner laboratory, The Francis Crick Institute, Mill Hill laboratory, London, UK

Marsupial mammals (also called Metatherians, e.g. kangaroos, opossums…) diverged from Eutherian mammals (e.g.

mouse, humans…) 180 million years ago. Marsupials have the remarkable distinction of giving birth to underdeveloped

pups that will finish their development outside the body of the mother. Their place in the mammalian evolutionary tree

and their incredible biology makes them extremely interesting to study evolution, reproduction and development.

Unfortunately, the use of this great model was very limited due to the lack of genetic tools available… until the era of

CRISPR/cas9 technology. We are developing the first in vivo/ex vitro genome engineered marsupial model, using the grey

short-tailed opossum. Inspired by the well-defined mouse protocols, we are now able to collect, inject, mutate and

culture opossum pre-implantation embryos. Our next challenge is the transfer of mutated embryos into surrogate

females to create in-vivo opossum mutants. This new CRISPR/Cas9 engineered opossum model can already be used to

study the biology of marsupial pre-implantation embryos and will hopefully be available soon for in-vivo studies, making it

possible to answer burning questions like ‘what is the sex determination system in marsupials?’

Platelet production in vitro for transfusion from human pluripotent stem cells: the

added power of genome editing

Thomas Moreau, Amanda Dalby, Annette Mueller, Guenaelle Bouet, Maria Colzani, Daniel Howard and Cedric Ghevaert

Department of Haematology, University of Cambridge and NHS Blood and Transplant, Cambridge, UK, and, Wellcome

Trust – Medical Research Council Cambridge Stem Cell Institute, Cambridge, UK

Platelet transfusions to thrombocytopenic patients are increasing by 7-10% per year. We are currently entirely reliant on

donor-derived platelets, which have limitations: short shelf-life and precarious supply chain, risk of donor-derived

transmitted infections and issues of HLA mismatch in chronic recipients. We are aiming to develop protocols to produce

platelets in vitro from a renewable source of stem cells - human pluripotent stem cells (hPSCs) - using a methodology and

reagents compatible with the production of a clinical grade commercially viable product. First we developed a chemically

defined forward programming (FoP) approach to produce megakaryocytes (MKs) from hPSCs based on the

overexpression of 3 key transcription factors (TFs; GATA1, FLI1 and TAL1) driven by lentiviral vectors. This FoP protocol

generates pure MK cultures (>80% CD41+ CD42+ cells) which expanded in vitro for several months culminating on

average to 2x105 MKs per starting hPSC with minimum cytokine requirement and cell handling. Second we addressed the

challenge of the low platelet number produced per MKs in vitro (at best 10 platelets per MK whilst in vivo it is estimated

that MKs produce >1000 platelets per cell). To improve platelet release and harvest, we recreated the charateristics of

the bone marrow vascular niche in vitro. First we use collagen-based 3-dimensional porous scaffolds to recreate the

physical 3-dimensional space. Second we screened a library of 50 recombinant ectodomain proteins in order to identify

cell-to-cell contact signal proteins that promote proplatelet formation. We identified two, which can be further

immobilised to collagen 3D-scaffolds leading to a 5-fold increase in platelet yield. Finally we integrated the scaffolds in a

bespoke parallel-flow two-chamber bioreactor to a platelet yield of >100 platelets per MK. Platelet are enucleated cells

and can therefore be irradiated prior to administration to patients. This minimises the potential tumorigenic risk

nucleated cells within the platelet harvest and allow the genetic modifications of the stem cells from which platelets are

eventually produced.

Genome editing can be applied in 3 key areas of this venture.

(a) We are creating cell lines where all 3 transcription factors affecting the forward programming will be under the

control of an inducible cassette inserted in a safe harbour of the genome. This will allow the production of cells

without lentiviral transduction.

(b) One of the main benefits from the production of platelets in vitro would be to provide platelets that match HLA

Class I expression for multitransfused or multiparous patients who are alloimmunised against HLA class I

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epitopes. We have now used CRIPR technology to knock-down HLA class I expression in stem cells lines from

which “universal” platelets can be generated.

(c) Platelets contain alpha-granules that harbour over 300 different proteins, including clotting and growth factors.

We can genome edit stem cells in order to “package” proteins of interest in the granules thereby generating

platelets that can act as delivery vehicles for specific growth or clotting factors.

RNA-therapeutics of gene haploinsufficiencies

Cristina Fimiani1, Elisa Goina

2 and Antonello Mallamaci

1

1SISSA, via Bonomea 265, 34136 Trieste, Italy

2present address: ICGEB, Area Science Park, Padriciano 99, 34149 Trieste, Italy

Artificial transactivation of endogenous genes ad libitum is a desirable goal for a number of basic and applied research

purposes. This goal has been recently achieved by three types of artificial enzymes: Zinc finger- (ZF-), TransActivator Like

Element- (TALE-), CRISPR- type transactivators. We recently developed a smaller and fully synthetic ribonucleoproteic

transactivator prototype. Kept together via an MS2 coat protein/RNA interface, it includes a fixed, polypeptidic

transactivating domain and a variable RNA domain that recognizes the desired gene. Thanks to this device, we specifically

upregulated five genes, in cell lines and primary cultures of murine pallial precursors. Even if gene upregulation was small,

however it was sufficient to inhibit neuronal differentiation. Our transactivator activity was restricted to cells in which the

target gene is normally transcribed. These features make our prototype a promising tool for clean rescue of gene

haploinsufficiencies, since it could lead to a specific overstimulation of the spared gene allele in its physiological

expression domain, even upon generalized delivery. On the other hand, we are interested in stimulating transcription of

endogenous genes by small activating RNAs (saRNAs). These are miRNA/siRNA-like molecules, supposed to destabilize

transcription-inhibiting ncRNAs or facilitate the recruitment of transcriptional complexes to chromatin. The transcription

gain they elicit is small; however it may be sufficient to influence the behaviour of cells in a robust way. Moreover, silent

genes generally do not respond to them. As such, also saRNAs are a promising tool for therapeutic stimulation of gene

transcription. We selected a number of saRNAs able to stimulate haploinsufficient genes involved in CNS morphogenesis

and physiology. We are studying their mechanisms of action as well as working at their exploitation for in vivo correction

of gene haploinsufficiency.

Use of CRISPR/CAS9 to investigate the function of gene mutations found in myeloid

malignancies

Hamid Dolatshad1, Simona Valletta

1, Matthias Bartenstein

2, Bon Ham Yip

1, Erica Bello

1, Shanisha Gordon

2, Yiting Yu

2,

Jacqueline Shaw1, Swagata Roy

1, Laura Scifo

1, Anna Schuh

3, Andrea Pellagatti

1, Tudor A. Fulga

4, Amit Verma

2, Jacqueline

Boultwood1

1Bloodwise Molecular Haematology Unit, Radcliffe Dept of Medicine, University of Oxford, JR Hospital, Oxford, UK

2Albert Einstein College of Medicine, Bronx, NY, USA

3NIHR Biomedical Research Centre, University of Oxford, Oxford, UK

4Weatherall Institute of Molecular Medicine, Radcliffe Dept of Medicine, University of Oxford, JR Hospital, Oxford, UK

The myelodysplastic syndromes (MDS) represent a heterogeneous group of myeloid malignancies. Recent studies have

illuminated the molecular landscape of MDS. The most common mutations found in MDS occur in genes that are

epigenetic modifiers (e.g. ASXL1) or regulators of RNA splicing (e.g. SF3B1). Approximately 78-89% of MDS patients

harbour at least one known gene mutation. Although it is clear that the common gene mutations impact both the

pathophysiology and prognosis in MDS, their role in MDS disease initiation and progression still unclear. We have used

the CRISPR/Cas9 mediated homology directed repair to correct the ASXL1 homozygous nonsense mutation present in the

KBM5 myeloid leukaemia cell line, which lacks ASXL1 protein expression. CRISPR/Cas9-mediated ASXL1 homozygous

correction resulted in protein re-expression with restored normal function, including down-regulation of polycomb

repressive complex 2 target genes. Significantly reduced cell growth and increased myeloid differentiation, providing new

insights into the role of ASXL1 in human myeloid cell differentiation. Mice xenografted with mutation-corrected KBM5

cells showed significantly longer survival than uncorrected xenografts. Given its successful application in the KBM5 cells,

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the CRISPR/Cas9 system is being used to correct recurrent gene mutations found in MDS haematopoietic stem and

progenitor cells (HSPC). We electroporate a complex composed of the recombinant Cas9 protein and gRNA with the DNA

template into the MDS progenitor cells. In this study we have generated a cell line with corrected mutation in epigenetic

modifier and believe that this is a powerful technique that can illuminate the effect of the common MDS associated

mutations on normal and malignant cellular function, and secondly to provide proof-of-concept for gene correction in

primary adult HSC derived from patients with a myeloid malignancy, with potential therapeutic application in the future.

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WORKSHOPS Workshop 1

Title: Beyond simple knockouts: considerations for successfully engineering disease SNPs and reporter transgene

integrations.

Abstract: Deriving knockout cell lines and animals has become relatively straightforward compared with projects focused

on modeling disease SNPs. A common roadblock is encountered when CRISPR-based SNP conversion works in one cell

type, but not in another cell type which is critical for generating biologically relevant information. Furthermore, it has

been observed that some donor DNA formats work at different rates among different cell types. This workshop will cover

strategies to help you make decisions about experimental design in genome editing which maintain project momentum

and derive meaningful biological data.

Workshop 2

Title: How to successfully perform exploratory primary screens using CRISPR/Cas9 and RNAi technologies

Abstract: The simplicity and low cost of programming CRISPR-Cas systems has created possibilities for high throughput,

genome wide interrogation of gene function. For example, a drug treatment of cells can be characterized by knocking out

tens of thousands of genes in parallel to identify pathways that lead to drug resistance. In this workshop, the key

considerations for executing screens will be discussed covering the entire experimental flow from gRNA design, to cell

treatments, to deep sequencing and statistical analysis.

Workshop 3

Title: Q & A session on CRISPR project design with Dr Barry Rosen

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POSTER ABSTRACTS Silencing Aspergillus terreus yap1 gene to clarify its role in the regulation of

lovastatin biosynthetic genes

Ailed Pérez-Sánchez, Roxana Miranda and Javier Barrios-González

Departamento de Biotecnología, Universidad Autónoma Metropolitana Unidad Iztapalapa UAM-I, México

Lovastatin (LOV) is a secondary metabolite, produced by Aspergillus terreus that lowers cholesterol levels in blood. Earlier

studies of our group showed a link between reactive oxygen species (ROS) and LOV biosynthesis, since a ROS build up

during production phase (idiophase) was detected. In a subsequent work we showed that ROS regulate LOV biosynthesis

at a transcriptional level, although the mechanism is presently unknown. It is considered that stress response

transcription factor(s) could be this link between ROS and the biosynthetic genes. A search for transcription factors

binding sites in the promoter regions of the LOV genes: lovE and lovF, revealed putative sites for several oxidative stress

response TFs including Yap1. In this work expression profiles of A. terreus yap1 and other relevant genes were performed.

In addition, yap1 was silenced and transformants characterized. Vector pGdpPkiRNAi-yap1 was constructed and

transformed into A. terreus TUB F-514. yap1 expression profile was very similar to that of sod1 during lovastatin

fermentation. That is, highly expressed during trophophase but down regulated during idiophase. In the transformants,

the ROS build up began before schedule, reaching higher levels. This brought about earlier and stronger expression of

LOV genes, and manifested in an early start of LOV biosynthesis and higher LOV production than the parental strain.

These results suggested that Yap1 could be acting as a negative regulator of LOV genes. Nevertheless, it could also be an

indirect effect due to earlier and higher ROS accumulation. A precise amount of antioxidant was added, to the LOV

fermentation with the siYAp strain, to generate a ROS accumulation profile similar to the one with the parental strain.

Under these conditions siYap and the parental strain showed similar LOV production kinetics. It is concluded that Yap1 is

not directly regulating LOV genes. It only directs antioxidant enzymes expression, thus regulating ROS accumulation.

Development of tools for modulation of genes involved in protein glycosylation by

genome editing

Luka Bočkor, Vanja Tadić, Paula Dobrinić, Dora Markulin, Aleksandar Vojta, Vlatka Zoldoš

Department of Molecular Biology, Faculty of Science, University of Zagreb, Horvatovac 102a, 10000 Zagreb, Croatia

Glycosylation is one of the most important post-translational modifications of proteins contributing to a number of

functions in protein biology, from protein folding and quality control to recognition events such as cell signalling and

immune surveillance. Glycans have come into focus recently as contributors or cause of different human complex

diseases. A recent GWAS study of the IgG glycosylation identified several genes with the IL6ST locus being among the top

hits. IgG represents an excellent model glycoprotein because it has one glycosylation site at each heavy chain. Many of

the structures present on IgG have been well defined with many important functional effects of alternative IgG

glycosylation described. On the other hand, our previous studies identified significant associations between HNF1A

methylation and plasma protein fucosylation. In order to carry out functional studies of the role of the IL6ST and HNF1A

genes in protein glycosylation, we constructed TALENs and Cas9-based nucleases targeting those genes. To test the

activity of the designed nucleases and select the most efficient ones, we measured their ability to induce double-strand

breaks at targeted sites in the HEK293 cell line by a recombination-based luciferase assay. The assay is based on a

reporter vector containing a duplicated region of the firefly luciferase cDNA. The repeats are separated by a sequence

that is targeted by the nucleases. Double strand breaks generated by the nucleases in the sequence, flanked by the

repeats, will favour the recombination within the luciferase gene and restore the correct primary sequence in the

reporter vector. Efficacy of different nucleases is then determined as luciferase activity. Using this assay we were able to

identify the most efficient nucleases for both genes in order to create knockouts in lymphoblastoma and HepG2 cell lines,

as surrogates for B cells and hepatocytes where IL6ST and HNF1A genes have a putative role in protein glycosylation.

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Antisense Oligonucleotide Mediated Rescue of a Deep Intronic Point Mutation in

OPA1

Tobias Bonifert1, Irene Gonzalez Menendez

1, Matthis Synofzik

2,3, Ludger Schöls

2,3, Bernd Wissinger

1

1 Molecular Genetics Laboratory, Institute for Ophthalmic Res, Centre for Ophthalmology, University of Tübingen, Germany

2Department of Neurodegenerative Diseases, Hertie-Institute for Clinical Brain Research, University of Tübingen, Germany

3Center for Neurodegenerative Diseases (DZNE), Helmholtz Association of German Research Centers, Tübingen, Germany

Inherited Optic Neuropathies (ION) represent an important cause of blindness in the European working-age population.

Despite progress in mutation screenings and disease gene identification, a large number of all cases remain unsolved –

partly due to exon-centered screening approaches. Recently we reported the discovery of four independent families with

deep intronic point mutations (DIM) in OPA1 that cause mis-splicing of the pre-mRNA transcript by creating a cryptic 3’

splice site (ss). In accordance with the haploinsufficiency model, heterozygosity for a DIM leads to reduced levels of OPA1

protein which in turn drives disorganization of the patients’ mitochondria and provokes ION. As a rescue strategy we

thought to prevent missplicing of the mutant pre-mRNA by applying 2’O-Methyl-Antisense Oligonucleotides (AONs) with

a full-length phosphorothioate backbone that target the cryptic 3’ss and the predicted novel branch point (BP) created by

the DIM, respectively. Transfection of patient-derived primary fibroblasts with these AONs could rescue correct splicing of

the mutant pre-mRNA in a time and concentration dependent mode of action. The treatment showed strong rescue

effects (~66%) using the 3’ss targeting AON and moderate rescue (~16%) using the BP targeting AON. Application of

mixtures of both AONs did not further enhance the effect. A peak efficiency could be observed two days after treatment,

however significant splice correction was still seen 14 days post transfection. Applying quantitative Western Blot analysis,

we could demonstrate increased amounts of OPA1 protein with a peak at ~3 days post treatment. Slight concentration-

dependent cytotoxic effects could be observed with the 3’ss targeting AON. These might be caused by off -target binding

sites in other transcripts that harbor ALU elements similar to the intron investigated here. In summary we provide the

first mutation-specific in vitro rescue strategy for OPA1 deficiency using synthetic AONs.

The utility of SmartFlare probes to detect heme oxygenase-1 transcript

level during selection of CRISPR/Cas-generated HO-1 knockout cells.

Maria Czarnek, and Joanna Bereta

Department of Cell Biochemistry, Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University in Kraków,

Gronostajowa 7, 30-387 Kraków, Poland

Knockout cells are invaluable research model in modern biological sciences. The ability to modify genomes in a precise

and efficient fashion is needed to understand how genotype affects different physiological and pathological processes.

Genome editing tools, including CRISPR/Cas9, enable researchers to modify virtually any locus across mammalian

genome. However, modification efficiency may be the primary limitation. Cas9 can induce the formation of targeted

double strand breaks in mammalian chromosomes. Subsequent indel formation via the nonhomologous end-joining

(NHEJ) repair mechanism often introduces nonsense or frameshift mutations, which leads to nonsense-mediated

degradation of mRNA. The aim of our project was to test whether SmartFlare probes may be utilized to sort out the cells

with a low level of a transcript of interest. Heme oxygenase-1-deficient HEK293 cells were generated by CRISPR/Cas9

genome engineering. Fluorescence intensity of the cells after their incubation with HO-1-specific SmartFlare probe was

evaluated with flow cytometry. The results were compared with qRT-PCR-based HO-1 relative expression level

measurements. Additionally, the levels of HO-1 transcript after HEK293 stimulation with iron-containing porphyrin

(hemin) were quantified using both qRT-PCR and SmartFlare fluorescence measurements.

Evaluation of CRISPR/Cas9 efficiency for muscle-specific Mir-378a targeting in vivo

Szymon Czauderna1, Maria Czarnek

2, Mateusz Tomczyk

1, Alicja Jozkowicz

1 and Jozef Dulak

1

1Department of Medical Biotechnology, Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University,

Gronostajowa 7, Krakow, Poland

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2Department of Cell Biochemistry, Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University,

Gronostajowa 7, Krakow, Poland

CRISPR/Cas system was adopted to target eukaryotic DNA making it the most widely used method for genome editing due

to its efficiency and simplicity. Our work aims to evaluate CRISPR/Cas efficiency of Mir-378a locus targeting in vivo using

adeno-associated viruses as vectors for CRISPR/Cas delivery. sgRNA flanking Mir-378a were manually designed and cloned

into pX601 plasmid (Addgene). sgRNA efficiency and plasmid performance were evaluated in Neuro-2a or C2C12 by Cel-I

digestion, diagnostic PCR or western blot. Off-target prediction was performed using rgenome software. For dual sgRNA

expression custom synthesized cassette was cloned into pX600 vector by Gibson assembly. For muscle-specific Cas9

expression short Cas9 version from Staphylococcus aureus was expressed from desmin promoter. Designed sgRNAs

show high cleavage efficiency ranging from 40 to 70%. Off-target analysis revealed substantial number of potential

unspecific sgRNA target sites. Thus, for further study only those with lowest potential off-target activity were selected.

Verification of custom plasmid showed that expression of two sgRNAs from single vector results in efficient excision of

DNA fragment which size corresponds to whole Mir-378a. Next, we evaluated desmin promoter for Cas9 expression. Cas9

expression driven from desmin promoter was undetected on western blot. However, even very low level of Cas9 (below

western blot detection limit) was sufficient to induce DNA cleavage as revealed by Cel-I digestion, but with lower

efficiency than from CMV promoter. Our results obtained so far indicate high efficiency of designed sgRNA for Mir-378a

targeting and full functionality of our custom prepared vectors which are suitable for AAV production. What is more, even

low level of Cas9 is able to induce DNA cleavage which might have important practical implication for genome editing in

vivo in terms of tissue specificity and safety.

Development of epigenetic CRISPR-Cas9 system for targeted methylation at specific

CpG sites

Paula Dobrinić, Aleksandar Vojta, Vanja Tadić, Luka Bočkor, Marija Klasić, Petra Korać, Vlatka Zoldoš

Division of Molecular Biology, Department of Biology, Faculty of Science, University of Zagreb, Horvatovac 102a, 10000

Zagreb, Croatia

DNA methylation is an important epigenetic mechanism involved in gene regulation. In mammals, DNA methylation

mostly occurs at symmetrical CpG dinucleotides. Cytosine methylation of gene regulatory regions has usually been

associated with gene silencing, but functional studies have been limited due to the lack of methods for targeted

manipulation of methylation marks. To that end, we developed a flexible, easily programmable tool for targeted

methylation at specific CpG sites in mammalian cells, based on the CRISPR-Cas9 system. We completely abolished the

nuclease activity of Cas9 protein and added the DNA methyltransferase domain of human DNMT3A, using a short peptide

linker. We validated this newly developed dCas9-DNMT3A fusion construct in HEK293 cells, where we targeted

unmethylated promoter regions of two different genes, BACH2 and IL6ST. At both loci, we tested a number of targeting

single guide RNAs (sgRNAs) binding in different orientations and at different positions relative to the CpG sites analysed

by bisulphite pyrosequencing. In most cases, dCas9-DNMT3A fusion induced significant increase in methylation level at

cytosines adjacent to sgRNA-binding site, reaching up to 30-60% at particular CpG sites. Transfection of pooled sgRNAs

targeting the same locus induced even higher methylation increase and decrease in the gene transcript level. The dCas9-

DNMT3A fusion protein can be used to efficiently introduce CpG methylation at specific genomic locus and to probe the

function of specific CpG methylation events in gene regulation.

Identifying somatosensory subtypes using CRISPR/Cas9-directed transgene

integration

Donald P Julien, Alex W Chan and Alvaro Sagasti

Department of Molecular, Cell and Developmental Biology, University of California, Los Angeles USA

The skin is a multimodal sensory organ, capable of detecting multiple types of touch stimuli (e.g. mechanical, thermal,

and chemical). Underlying this sensory diversity are multiple populations of somatosensory axons that terminate as

specialized sensory endings in the skin. The diversity of sensory termini in the skin and the how their organization is

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established during development, however, remains poorly understood. The embryonic zebrafish is an ideal model for

investigating the development of somatosensory neurons because they can be imaged in live animals. However,

differentiating individual sensory subtypes remains a challenge due to a paucity of transgenic reporters. We hypothesize

that sensory subtypes can be distinguished by the expression of specific molecules that regulate their development or

sensory function. To test this hypothesis and generate new transgenic tools for analyzing somatosensory subtypes, we are

using CRISPR/Cas9-directed mutagenesis to target integration of Gal4-containing reporter transgenes at candidate

genomic loci. To identify candidate genes that label somatosensory subtypes, we are targeting transgenes to the

upstream enhancer regions of each candidate. To interrogate the function of promising candidate genes, we are also

generating loss-of-function gene traps by targeting transgene integration to the coding region of each locus. Consistent

with previous studies, this approach results in highly efficient transgene integration at specific genomic loci. In addition to

providing information about the function of somatosensory subtypes and the molecular mechanisms that organize their

innervation territories in the skin, this study will generate valuable tools for future studies of somatosensory

development.

Direct delivery of Cas9-sgRNA protein complex via cell-derived nanovesicles

Montse Morell, Tatiana Garachtchenko, Lily Lee, Thomas P. Quinn, Michael Haugwitz, and Andrew Farmer (Presenter by

Cornelia Hampe)

Takara Bio Europe, France

While CRISPR/Cas9 is a powerful technique for genome manipulation, two significant challenges remain: obtaining

efficient delivery of Cas9 to all cell types and achieving fewer off-target effects. Recently, it has been demonstrated that

genome editing via delivery of Cas9 protein is as effective as plasmid-based delivery, but with the added benefit of less

off-target effects due to the short lifespan of the Cas9 protein in the cell (1). Here we report delivery of Cas9 protein using

cell-derived nanovesicles called gesicles. Gesicles are produced by a mammalian packaging cell via co-overexpression of

three components: a nanovesicle-inducing glycoprotein, Cas9, and the sgRNA specific to the target gene. Gesicles display

a protein on their surface that mediates binding and fusion with the cellular membrane of target cells. Based on this

principle, we have developed a method for actively packaging sgRNA-loaded Cas9 into gesicles via ligand-dependent

dimerization (iDimerize™ technology). This dimerization approach allowed us to efficiently package active Cas9 containing

a nuclear localization signal (NLS) into these nanovesicles. Gesicle-based protein delivery does not rely on recombinant

protein from a bacterial source or the use of a transfection reagent. We show that gesicles carrying a Cas9- sgRNA protein

complex (Cas9 Gesicles) can mediate target-gene knockout in a variety of cell types. The observed knockout efficiency is

often considerably higher than what is observed with plasmid-based delivery of sgRNA and Cas9. This nanoparticle-based

method allows for tight control of the dose and duration of the Cas9-sgRNA complex in the cell; this level of control

decreases off-target effects. Close analysis of the particles shows that they are stable over several freeze/thaw cycles and

are consistent in size (~150–170 nm). Overall, these particles can be considered a novel and effective tool for Cas9 and

sgRNA delivery to any target cells.

Engineering the mouse genome using CRISPR/Cas9 technology

Ruairidh King, Joffrey Mianné, Rachel Fell, Adam Caulder, Gemma Codner, Marina Maritati, Martin Fray, Wendy Gardiner,

MLC Microinjection team, Sara Wells, and Lydia Teboul

The Mary Lyon Centre, MRC Harwell, HSIC, Oxon, OX11 0RD, UK

Mouse models are valuable tools to understand gene functions, genetic diseases, and to develop and test new

therapeutic treatments in vivo. The ability to introduce tailored modifications within the mouse genome is essential to

generate these models of human diseases. The recently developed CRISPR/Cas9 system as genome engineering tool has

brought new perspectives for the generation of mouse models in a precise fashion, at reduced price, and within a shorter

time scale. Here we will report the use of the CRISPR/Cas9 technology at the Mary Lyon Centre, MRC Harwell, to engineer

the mouse genome through different methods. We will first present our optimised high throughput mouse production

pipeline allowing us to generate alleles containing indels, tailored deletions, or point mutations through direct injection

into zygotes. We will then report the use of the CRISPR/Cas9 technology to engineer and enhance the genetic background

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of the C57BL/6N mouse strain by correcting a mutation in the Cdh23 gene. We will present our data obtained for

enhancing the homologous recombination rate in mouse embryonic stem cells through co-electroporation of the IKMC

targeting vector and CRISPR reagents. Finally, we will introduce our pilot of a new delivery strategy of CRISPR reagents

through in vivo testis electroporation. Developing these methods for genome engineering will enable the generation of a

wide range of increasingly complex alleles in mice, both in a bespoke and high-throughput context.

Candidate testing of axonal sprouting genes in a cortical circuit induced by limb

overuse after stroke

Esther H. Nie1, Giovanni Coppola

2, Riki Kawaguchi

2, S. Thomas Carmichael

1

1Dept. Neurology, UCLA David Geffen School of Medicine, Los Angeles, CA

2Depts of Psychiatry and Biobehavioral Sciences, UCLA David Geffen School of Medicine, Los Angeles, CA

Stroke survivors worldwide are left with permanent sensorimotor and cognitive disabilities for which there are no medical

treatments. However, a behavioral paradigm of limb overuse after stroke, known as constraint-induced movement

therapy (CIMT), has been shown in clinical trials to result in significant motor improvements. The goal of the current

project is to understand how specific gene systems drive this important neural repair process. We have previously

mapped a unique connection to premotor cortex upon post-stroke limb overuse. We find that CIMT-like limb overuse

drives the activity-dependent formation of connections between premotor cortex (PMC) and retrosplenial cortex (RSC), a

brain area involved in spatial learning. The finding was significant and replicated across two independent cohorts (p<0.05,

Hotelling’s T test). PMC connected RSC neurons were FACS purified for RNA-Seq transcriptome analysis. Bioinformatics

reveal that the RSC-PMC circuit is characterized by circuit-specific expression of activity-induced and growth-related

genes. Pathway analyses indicate that limb overuse induces canonical signaling in calcium signaling, cell cycle, and Ephrin

A pathways. From the relatively small group of 162 genes that are regulated by limb overuse, top candidates have been

statistically prioritized (FDR<0.1, p<0.005) for in-vitro neuronal outgrowth screening using primary cortical neurons.

Recent knockout experiments in-vitro show that gRNAs directed against these genes can facilitate CRISPR/cas9 editing of

candidate genes. Cel-I mutation detection has confirmed knockout of selected genes. Ongoing design of an AAV-mediated

cas9 for in-vivo delivery aims to target candidates that increase neuronal outgrowth from the in-vitro screening dataset.

Through single and multiplexed genetic gain and loss-of-function studies, we have promising molecular candidates to

enhance convergent injury and activity-dependent molecular pathways during brain repair after stroke.

Functional Multiparameter Analysis of Double Strand Breaks and Associated

Biomarkers during Genome Editing

Lysann Sauer1, Stefan Rödiger

1, Jens Schneider

1, Peter Schierack

1, Dirk Roggenbuck

1,2, Christian Schröder

1

1Faculty of Natural Sciences, Brandenburg University of Technology Cottbus – Senftenberg, Germany

2Medipan GmbH, Dahlewitz/Berlin, Germany

A powerful and highly precise technology, adapted from the microbial adaptive immune system, is enabling the systemic

analysis of the mammalian genome. The CRISPR-associated RNA-guided endonuclease Cas9 induces target-specific

double-strand break (DSB) thereby activating DNA repair pathways such as nonhomologous end joining (NHEJ) or

homology-directed repair (HDR). The delivery of the CRISPR/Cas9 components and potential off-target mutagenesis are of

concern and require further elucidation. This is of particular importance for clinical applications and personalized ;

therefore, we used our automatized imaging platform AKLIDES® to functionally analyze cellular responses upon genome

editing via CRISPR/Cas9. Firstly, we established a number of sgRNAs targeting the DNA loci of: Splicing Factor

Proline/Glutamine-Rich (SFPQ), Centromere protein B (CENP-B) and Sjögren syndrome type B antigen (SS-B). SFPQ is

involved in DNA repair response, while the latter two have implications in autoimmune disease and cancer. We

transfected HEK293T cells and later included CaCo-2 and HepG2 cells. After we verified the functionality of the sgRNA, we

did immunofluorescence staining to analyze the location of the Cas9 protein as well as the DSB marker 53BP1 and

phosphorylated histone H2AX (γH2AX). Digital image analysis was done using the AKLIDES® platform. Etoposide

treatment was used as a control to induce DSBs, γH2AX foci were successfully co-localized with 53BP1 and further

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biomarkers independently of the cells type and transfection efficiency. Our data indicate correlations between

sgRNA/Cas9 treatment and γH2AX and 53BP1; however, further investigations are needed using Fluorescence in situ

hybridization.

Involvement of lncRNAs in muscle differentiation

Rossella Tita1, Monica Ballarino

1, Andrea Cipriano

1, Fabio Desideri

1 and Irene Bozzoni

1,2,3

1Dept of Biology and Biotechnology Charles Darwin, Sapienza University of Rome, Rome, Italy

2Center for Life Nano Science@Sapienza, Istituto Italiano di Tecnologia, Rome, Italy

3Institute Pasteur Fondazione Cenci-Bolognetti and IBPM, Sapienza University of Rome, Rome, Italy

In recent years, it has been discovered that genomes of multicellular organism are characterized by the pervasive

expression of different types of non-coding RNAs (ncRNA) and among these, long non-coding RNAs (IncRNAs) are

included. A transcriptome analysis performed during in vitro murine muscle differentiation allowed us to identify new

lncRNAs differentially expressed during myogenesis. These transcripts were classified on the basis of their expression in

proliferating versus differentiated conditions, muscle-restricted activation and subcellular localization. We are now

focusing on the characterization of a nuclear lncRNA, lnc-405, which is up-regulated in myotubes and conserved in

human. To dissect its role in vivo we are using the CRISPR Cas9 strategy to generate transgenic mice which phenotypes

need to be deeply characterized. This strategy consists in the co-injection of Cas9 mRNAs, a targeting sequence (sgRNA)

and a single-stranded oligodeoxynucleotide (ssODN) as a template for homology-directed repair (HDR) system, into the

zigote. With this strategy we successfully obtained heterozygous mice and now, we are crossing them to generate the

homozygous mouse. A similar strategy will be also performed in human myoblasts to study whether lnc-405 function is

conserved during the evolution.

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