Enhancing the Efficiency of CRISPR/Cas9 Precise Gene Editing for Cystic Fibrosis Gene Therapy

by

Kyle E. Seigel

A thesis submitted in conformity with the requirements for the degree of Master of Science

Department of Laboratory Medicine and Pathobiology University of Toronto

© Copyright by Kyle E. Seigel, 2018

ii

Enhancing the Efficiency of CRISPR/Cas9 Precise Gene Editing for

Cystic Fibrosis Gene Therapy

Kyle E. Seigel

Master of Science

Department of Laboratory Medicine and Pathobiology University of Toronto

2018

Abstract

Cystic fibrosis (CF) is the most common cause of chronic obstructive lung disease in children

and young adults, yet there is no cure for this disease. Intimations of curative strategies have

emerged from gene therapy, but these approaches have failed in clinical trials due to insufficient

safety and efficacy profiles. The molecular technology necessary to overcome preeminent

challenges in gene therapy may reside in programmable nucleases – namely CRISPR/Cas9 –

which allow for site-specific genomic integration of therapeutic transgenes. However, the

clinical utility of these technologies is limited by inherently low efficiencies. Here, we use flow

cytometry and PCR assays to demonstrate the efficacy of perturbing DNA repair to overcome

such limitations. We demonstrate that overexpressing CtIP and EXO1-4D can enhance the

efficiency of CRISPR/Cas9-directed transgene integration by ~3- and ~6-fold, respectively.

These discoveries may provide necessary impetus for translating gene therapies into clinical

realities for genetic diseases such as CF.

iii

Acknowledgements

My thesis work and development as a scientist would not have been possible without the

professional wisdom, scientific expertise and personal support of a number of individuals. I would

first like to extend my utmost thanks and gratitude to my supervisor, Dr. Jim Hu. Your expertise

were foundational to the development of this project, and your continual encouragement and

support helped guide me through the adversities of science. I appreciate your persistent attention

to my experiments and challenges in the lab, regardless of how many other things you had going

on. This is a true testament to how much you care for your students; for their personal and

professional development as scientists. I would also like to thank my committee members: Dr.

Martha Brown, Dr. Grant Brown, and Dr. Andras Nagy. The diverse range of scientific expertise

that each of you contributed were instrumental to the progression of my project, and I am grateful

for your many hours of undivided attention. A special thanks to Dr. Martha Brown, who

continually made herself available to discuss my research, as well as to provide technical and

moral support. To Dr. Harry Elsholtz, thank you for your professional guidance and support

throughout the duration of my studies.

My fellow lab members were also extremely helpful in the development of this project. Dr. Huibi

Cao, Linda Li, Cathleen Duan, Fushan Shi, Emily Xia, Amy Zhang, Peter Zhou, Alexandra

Georgiou, Leo Yang, and Randolph Kissoon have all contributed to this work in some capacity.

Thank you for your helpful contributions in lab meetings, technical support at the bench, and

continual friendships. I would also like to extend my gratitude to all the staff in The SickKids-

UHN Flow and Mass Cytometry Facility at PGCRL. In particular, thank you Dr. Sherry Zhao and

Dr. Emily Reddy for tremendous technical support in designing experiments, operating flow

cytometers and analyzing data.

My heartfelt thanks to my family, without whom none of this work would have been possible. To

my Mom and Dad, and my brothers Jordan and Mathew: your unconditional love and support

form the bedrock upon which my career has thus far been built. Although none of you know

precisely what it is that I do, you have continuously supported my endeavours throughout my

academic career without question. Science is a fascinating and tremendously rewarding pursuit,

but it is not without its hardships. You guys have been there through it all and I am forever

iv

grateful. To my parents in particular, the invaluable sacrifices you have both made for my

education, my interests, personal growth and career development are incalculable, without which

none of this is possible. Finally, to my late grandfather Dr. Harold O. Seigel: Papa, your voracity

for understanding nature and insatiable passion for knowledge reverberate in me to this day; and

your contributions to science and technology have been a continual source of inspiration.

v

Table of Contents

Acknowledgments ....................................................................................................................... iii

Table of Contents ......................................................................................................................... v

List of Tables ............................................................................................................................... vii

List of Figures .............................................................................................................................viii

Abbreviations ............................................................................................................................... ix

CHAPTER 1. INTRODUCTION .............................................................................................. 1

1.1 Cystic fibrosis ........................................................................................................................ 1

1.1.1 CF biochemistry .................................................................................................... 1

1.1.2 CF pathophysiology .............................................................................................. 2

1.1.3 CF therapies .......................................................................................................... 3

1.2 Gene therapy .......................................................................................................................... 4

1.2.1 CF gene therapy .................................................................................................... 6

1.3 Gene editing ........................................................................................................................... 7

1.3.1 Zinc finger nucleases ............................................................................................. 8

1.3.2 TALENs ............................................................................................................... 10

1.3.3 CRISPR ............................................................................................................... 11

1.3.3.1 Discovery and mechanisms ................................................................... 11

1.3.3.2 Genome engineering ............................................................................. 12

1.3.3.3 CRISPR/Cas9 biochemistry and engineered Cas9 variants .................. 13

1.3.3.3.1 dCas9............................................................................. 15

1.3.3.3.2 Base pair editors ............................................................ 16

1.3.3.3.3 sgRNA engineering ....................................................... 17

1.3.3.4 CRISPR/Cas9 in genetics and disease modeling .................................. 17

1.3.3.5 CRISPR/Cas9 in gene and cell therapies .............................................. 18

1.4 DNA double-strand breaks .................................................................................................. 21

1.4.1 Non-homologous end-joining ............................................................................. 22

1.4.2 Homology-directed repair ................................................................................... 23

1.4.3 DSB repair pathway choice ................................................................................. 24

vi

1.5 Enhancing gene targeting efficiency by perturbing DSB repair .......................................... 29

CHAPTER 2. HYPOTHESIS AND OBJECTIVES ............................................................... 31

2.1 Rationale .............................................................................................................................. 31

2.1.1 CDK1 .................................................................................................................. 31

2.1.2 Post-translational modification-mimics .............................................................. 31

2.1.3 CtIP ..................................................................................................................... 32

2.1.4 EXO1 .................................................................................................................. 33

2.2 Hypothesis ........................................................................................................................... 34

2.3 Objectives ............................................................................................................................ 34

CHAPTER 3. MATERIALS AND METHODS ..................................................................... 35

3.1 Plasmids ............................................................................................................................... 35

3.2 Flow cytometry .................................................................................................................... 36

3.3 Junction PCR ....................................................................................................................... 37

3.4 Generating the IB3-1-CFTR-EGFP cell line ....................................................................... 37

3.4.1 Transfection ........................................................................................................ 37

3.4.2 FACS .................................................................................................................. 38

3.4.3 Puromycin selection and limited dilution ........................................................... 38

3.5 ssDNA donors ....................................................................................................................... 39

3.6 T7E1 assay ............................................................................................................................ 39

3.7 Data analysis ......................................................................................................................... 40

CHAPTER 4. RESULTS .......................................................................................................... 41

4.1 Flow cytometry assay for measuring gene targeting efficiency ........................................... 41

4.2 CtIP and EXO1-4D significantly enhance gene targeting efficiency .................................. 44

4.3 IB3-1-CFTR-EGFP cell line ................................................................................................ 51

CHAPTER 5. DISCUSSION.................................................................................................... 57

5.1 Future directions ................................................................................................................... 61

5.2 Conclusion ........................................................................................................................... 66

REFERENCES ......................................................................................................................... 68

vii

List of Tables

Table 1. DSB repair proteins and associated mutations

viii

List of Figures

Figure 1. CRISPR/Cas9-directed genomic DNA cleavage.

Figure 2. NHEJ is used to repair DSBs in G1 when CDK activity is low.

Figure 3. HDR is used to repair DSBs in S/G2 when CDK activity is high.

Figure 4. Schematic diagram of mCherry gene integration into the AAVS1 locus.

Figure 5. Enhancement of precise gene targeting efficiency by CDK1, CtIP and EXO1-4D in

HEK293 cells.

Figure 6. Enhancement of precise gene targeting efficiency by CtIP and EXO1-4D in HEK293

cells as measured by flow cytometry.

Figure 7. Enhancement of precise gene targeting efficiency by SCR7 in HEK293 cells.

Figure 8. Junction PCR demonstrating the enhancement of precise gene targeting efficiency by

CtIP and EXO1-4D in HEK293 cells.

Figure 9. Schematic representation of the IB3-1-CFTR-EGFP cell line and the associated

colour-switch assay.

Figure 10. Investigating the identity and CRISPR cleavage efficiency of IB3-1-CFTR-EGFP

cell line clones.

Figure 11. IB3-1 CFTR homology arm sequencing.

Figure 12. Single-stranded mCherry donor production.

ix

Abbreviations

53BP1 p53-binding protein 1

AAV Adeno-associated virus

AAVS1 Adeno-associated virus integration site 1

ABC ATP-binding cassette

AdV Adenovirus

Arg Arginine

ASL Airway surface liquid

Asp Aspartic acid

ATM Ataxia telangiectasia mutated

ATP Adenosine triphosphate

BG Benzylguanine

BIR Break-induced replication

cDNA Complementary DNA

CDK Cyclin-dependent kinase

CF Cystic fibrosis

CFTR Cystic fibrosis transmembrane conductance regulator

CRISPR Clustered regularly interspaced short palindromic repeats

crRNA CRISPR RNA

CtIP Carboxyl-terminal binding protein (CtBP)-interacting protein

dCas9 dead-Cas9

DMSO Dimethyl sulfoxide

DNA-PKcs DNA-dependent protein kinase catalytic subunit

DSB Double-strand break

dsDNA Double-stranded DNA

EDTA Ethylenediaminetetraacetic acid

EGFP Enhanced green fluorescence protein

ENaC Epithelium sodium channel

ER Endoplasmic reticulum

ESC Embryonic stem cell

x

EXO1 Exonuclease 1

FACS Fluorescence-activated cell sorting

FBS Fetal bovine serum

Glu Glutamic acid

GTE Gene targeting efficiency

HD-Ad Helper-dependent adenovirus

HDR Homology-directed repair

HEK293 Human embryonic kidney 293

HPV Human papillomavirus

HR Homologous recombination

HSPC Hematopoietic stem and progenitor cells

Indel Insertion or deletion

iPSC Induced pluripotent stem cell

LDL Low-density lipoprotein

LNP Lipid nanoparticle

MEM Minimal essential media

MMEJ Microhomology-mediated end-joining

MRN MRE11:RAD50:NBS1

MSD Membrane-spanning domain

NBD Nucleotide binding domain

NHEJ Non-homologous end-joining

NPG Non-obese diabetic (NOD)/Prkdcscid/IL-2Rnull

NSC Neural stem cell

NSG Immunodeficient, non-obese diabetic (NOD), severe combined immunodeficient

(SCID) c-/-

ORF Open reading frame

p(A) Polyadenylation

PAM Protospacer-adjacent motif

PCR Polymerase chain reaction

PBS Phosphate-buffered saline

PKA Protein kinase A

PSC Pluripotent stem cell

xi

PTM Post-translational modification

RIPA Radioimmunoprecipitation assay

RNAi RNA interference

RNP Ribonucleoprotein

RPA Replication protein A

RT-qPCR Reverse transcription quantitative real-time polymerase chain reaction

RVD Repeat-variable di-residue

SCID Severe combined immunodeficient

SDSA Synthesis-dependent strand annealing

SEM Standard error of the mean

sgRNA Single-guide RNA

SIN Self-inactivating

siRNA Small interfering RNA

SNCA Synuclein alpha

ssDNA Single-stranded DNA

TALE Transcription activator-like effector

TALEN Transcription activator-like effector nuclease

tracrRNA trans-activating CRISPR RNA

WT Wild type

ZF Zinc finger

ZFN Zinc finger nuclease

-RV Gamma-retrovirus

- 1 -

Chapter 1 Introduction

1.1 Cystic fibrosis

Cystic fibrosis (CF) is a life-threatening autosomal recessive disease caused by mutations in the

CF transmembrane conductance regulator (CFTR) gene. CF constitutes the most common cause

of severe chronic obstructive lung disease and exocrine pancreatic dysfunction in children and

young adults. The estimated worldwide incidence of CF is 1 in 2,500-3,000 newborns, and the

disease affects more than 70,000 individuals worldwide1,2. Furthermore, although CF is classified

as a rare disease, it is the most common lethal genetic disease in the Caucasian population1,2.

Disease onset occurs within the first year of life, and approximately two thirds of patients are

diagnosed within this time3. CF-related complications can persist throughout the duration of life,

for which life expectancy is approximately 50 years in the developed world3,4.

1.1.1 CF biochemistry

CFTR is a polytopic transmembrane protein that functions as a chloride (Cl−) and bicarbonate

(HCO3−) channel on the apical membranes of secretory epithelia, such as the lungs, sweat glands,

pancreas, and other exocrine glands5. CFTR is a member of the ATP-Binding Cassette (ABC)

transporter family, which utilize ATP hydrolysis to transport substrates such as ions, toxins,

peptides, drugs, and vitamins across biological membranes6. CFTR has a minimal architecture

common to all ABCs. It is made up of two cytosolic nucleotide binding domains (NBDs) that

undergo dimerization upon ATP hydrolysis, and two membrane-spanning domains (MSDs) that

come together to form a single transmembrane ion transport channel7. Most ABC proteins are

active transporters, and therefore use this modular structure in order to couple ATP hydrolysis to

transport substrates against a concentration gradient6. In contrast, ATP hydrolysis powers the

opening and closing of the CFTR ion channel, which in turn mediates passive electrochemical

diffusion of Cl- and HCO3- across cell membranes8,9. CFTR is also unique for its intrinsically

- 2 -

disordered regulatory domain (R domain), which controls channel activity in response to

phosphorylation by cAMP-dependent protein kinases, such as protein kinase A (PKA)8,9.

CF disease pathology is a consequence of structural, functional and/or regulatory defects in CFTR

due to genetic mutation. More than 2000 CFTR sequence variants have been identified to date,

including missense and nonsense mutations, frameshifts, and indels. There are various

mechanisms by which these mutations can result in CFTR dysfunction, the most notable of which

is defective protein processing. This is exemplified by the most common disease-causing CFTR

mutation, the F508 deletion in the NBD1 domain10–12. Almost 50% of CF patients are

homozygous for this mutation, and ~85% carry one F508 allele4. F508 disrupts normal

topography on the NBD1 surface, such that stabilizing interactions at the interface of NBD1 and

MSDs are prevented13. This local structural defect therefore interferes with domain-domain

interactions and prevents F508 CFTR from attaining a native conformation. As a consequence,

F508 CFTR forms aggregate-prone structures that are recognized by quality control mechanisms

in the ER during CFTR maturation, leading to ubiquitination and subsequent proteasome-

mediated degradation14,15. This ultimately prevents functional CFTR trafficking to the plasma

membrane. Other mutations are rare, but can cause CFTR dysfunction via defective channel

regulation or conduction, insufficient protein production, or reduced protein stability16.

1.1.2 CF pathophysiology

In the disease state, CFTR dysfunction results in Cl- and HCO3- impermeability across epithelial

apical membranes5. Such defective ion conductivity alters the quantity and composition of

epithelial fluids, in turn precipitating organ dysfunction. The widespread nature of this

abnormality in various tissues is concordant with the diverse clinical manifestations of CF disease

pathology, namely respiratory disease, intestinal obstruction, pancreatic dysfunction, salty sweat

and male infertility5. However, the main cause of morbidity and mortality in CF is progressive

lung disease17.

CF respiratory pathology is a consequence of defective CFTR-regulated ion homeostasis between

lung epithelia and the airway surface liquid (ASL), resulting in epithelial dehydration and

- 3 -

thickened mucus. The ASL is a region of fluid on the apical surface of airway epithelia, which is

composed of a periciliary layer adjacent to the epithelium and a superficial mucus layer. The

mucus layer contains gel-forming mucins that trap inhaled particles, which are subsequently

removed from the lung via mucociliary clearance18. Furthermore, antimicrobial factors within the

ASL are involved in innate and adaptive host immunity to protect the airways from inhaled

pathogens. The ASL requires a tightly regulated volume, pH, ionic and nutrient content for

sufficient antimicrobial activity, proper ciliary function and mucociliary clearance18.

CFTR directly controls anion flow between the airway epithelia and the ASL by mediating HCO3-

and Cl- movement, and indirectly by controlling Na+ diffusion via the epithelium sodium channel

(ENaC)19,20. CFTR normally inhibits ENaC, but its loss of function in CF results in excessive Na+

flow into epithelia. These electrochemical changes draw water into the epithelium, depleting ASL

depth and immersing cilia in the mucus layer, an environment in which their movement is severely

impaired. As a consequence, mucus builds up and airways become obstructed due to insufficient

mucociliary clearance21. Viscous and nutrient-rich mucus in turn provide a favourable

environment for bacterial colonization. Furthermore, the functions of many antimicrobial factors

are inhibited in the abnormally acidified ASL, a consequence of defective CFTR-mediated HCO3-

transport18. Consequently, CF patients suffer from chronic bacterial infections that facilitate

characteristic features of CF lung pathology, such as inflammation and progressive lung disease

involving tissue remodelling.

1.1.3 CF therapies

CF survival has dramatically improved from 5 years in the 1960s to now up to 50 years due to

clinical advances in treating symptoms and slowing disease progression3,4. The significant

progress made in extending life expectancy is largely a result of standardized multi-organ

therapies. Such treatments include antibiotics for bacterial lung infections – especially

Pseudomonas aeruginosa, a hallmark of CF – in addition to mucolytics that release thick

pulmonary mucus, chest physiotherapy, and high-calorie nutrition22. Lung transplantation

programs have also been beneficial23, and collectively these treatments have resulted in half of

CF patients being represented by adults in numerous countries24.

- 4 -

Novel mutation-specific therapies have recently been introduced into the clinic and many others

are in development or clinical trials. High-throughput drug screening has led to the identification

and development of two categories of mutation-specific therapies: ‘correctors’ that target the

CFTR processing defect, and ‘potentiators’ that improve mutant CFTR channel activity. Ivacaftor,

a CFTR potentiator, has been clinically successful in improving CFTR ion transport and

pulmonary function in patients with the G551D CFTR genotype25. However, these patients still

require additional therapies including pancreatic enzyme replacement, inhaled mucolytics, and

antibiotics16. Furthermore, this class of CFTR mutations accounts for only 4-5% of CF patients4.

As previously mentioned, ~85% of CF patients have the 'F508 CFTR defect. The drug

lumacaftor, a ‘corrector’, initially showed promise in rescuing the 'F508 CFTR folding defect in

primary cells26. However, this drug proved ineffective in improving lung function in 'F508

homozygous patients in clinical trials27. Lumacaftor-ivacaftor combinatorial therapy has also been

explored (Orkambi) for the 'F508 defect. Although this therapy has been approved due to success

in clinical trials, the overall efficacy was modest at best28. Accordingly, no effective treatments

are available for 'F508 CFTR patients, the most common CF-causing mutation.

Overall, despite clinical advances in treating symptoms and slowing disease progression, there is

still no cure for CF. As small molecules have proven inadequate for dramatically improving CF

pathology, distinct efforts have been made to correct the underlying source of the disorder, i.e.,

the CFTR genetic defect. For these purposes, investigators have long been interested in the

prospect of gene therapy for CF ever since the CFTR gene was discovered29.

1.2 Gene therapy

Scientists, clinicians and the general public have long been captivated by gene therapy, which

involves the correction or replacement of malfunctioning genes that give rise to disease

pathologies. Considerable interest in this approach is warranted due to the potential to treat

diseases at their genetic origins. Classically, gene therapy has been based on non-targeted,

retroviral-mediated insertion of therapeutic transgenes into the host genome. The approach was

initially based on gamma-retroviruses (J-RVs), but was evidently accompanied by numerous

limitations and salient risk factors, namely, poor gene transfer efficiencies in progenitor cells and

- 5 -

tumorigenesis due to vector insertion near oncogenes30–34. Accordingly, J-RVs were largely

replaced by safer and more efficacious vectors, such as self-inactivating-J-RVs (SIN-J-RVs) and

lentiviruses, which have found success in a number of phase 1/2 clinical trials in the past few

years. Notably, ex vivo gene transfer and subsequent transplantation of autologous hematopoietic

stem and progenitor cells (HSPCs) has been clinically useful in treating the genetic immune

diseases Wiskott-Aldrich syndrome (WAS)35,36 and X-linked severe combined immunodeficiency

(SCID-X1)37; as well as the blood disease E-thalassaemia38; and the neurodegenerative storage

diseases adrenoleukodystrophy39 and metachromatic leukodystrophy40.

Despite clinical success using improved delivery vectors, integrating viral vectors carry risks of

transcriptional silencing, insertional mutagenesis, and oncogenic transformation41–43.

Accordingly, lipid nanoparticles (LNPs) have emerged as an alternative to viral-mediated gene

delivery. LNPs similarly have the capacity to protect nucleic acid contents from degradation while

allowing cellular uptake of their cargo. However, these delivery methods are relatively generic

and cell type-nonspecific. Nonetheless, there are a number of clinical trials currently investigating

the utility of LNP-delivered siRNAs for the treatment of viral infections, cardiovascular diseases

as well as cancer44. In the realm of genetic diseases, LNPs have thus far shown preclinical utility

for the treatment of ocular45,46, auditory47, and hepatic disorders48.

Non-integrating viral vectors have also been successfully used for gene therapy applications.

These viral delivery platforms are maintained as episomal vectors expressing therapeutic

transgenes. Therefore, while avoiding potential insertional mutagenesis, the lack of permanent

genetic modifications restricts the utility of these approaches to targeting low-turnover or non-

proliferating cells. Various serotypes of adeno-associated viruses (AAVs) have been employed

for these purposes. For example, AAVs are able to stably express coagulation factor IX (FIX) – a

protein deficient in hemophilia B patients – from non-proliferating hepatocytes in animal

models49. Improvements in AAV vector design have since resulted in successful clinical

treatments of hemophilia B using this gene therapy approach50. Furthermore, AAV-based gene

therapies for type 2 Leber congenital amaurosis (LCA) and retinitis pigmentosa (both inherited

retinal diseases (IRDs)), recently became the first FDA-approved in vivo gene therapies for

clinical commercial use in the United States under the brand name LUXTURNATM (voretigene

neparvovec, AAV2-hRPE65v2) by Spark Therapeutics (NCT00999609)51.

- 6 -

1.2.1 CF gene therapy

Although CF affects multiple organs of the body, the primary cause of morbidity and mortality is

progressive lung disease. Furthermore, because of the monogenic nature of CF and the

accessibility of airway epithelia to gene therapy vectors, corrective gene therapy for CF lung

disease has been a major effort for the last few decades. Three years after the discovery of the

CFTR gene in 198929, Rosenfeld et al. successfully delivered replication-deficient first generation

adenovirus (AdV) carrying CFTR cDNA to the airway epithelium of rats, from which transgenic

CFTR mRNA and protein could be expressed52. AdVs are double-stranded, non-integrating

episomal viruses with a natural tropism for airway epithelial cells53. Furthermore, AdVs possess

high transduction efficiencies and relatively low virulence, and were therefore an obvious first

choice for CF gene therapy. Alternative non-viral liposomal CFTR delivery had also partially

corrected Cl- transport defects in the tracheal epithelium of CFTR knockout mice54. These

promising preclinical studies precipitated the first CF gene therapy trial. Although the study was

small and not placebo-controlled, Zabner et al. demonstrated correction of Cl- transport defects in

the nasal epithelium of three CF patients using first generation AdV carrying CFTR cDNA55.

Similar results were also obtained using liposome-mediated gene transfer56. Since then, nine

additional AdV CF gene therapy trials have been conducted57–65. However, this approach has

ultimately failed due to inefficient gene transfer, unsustainable gene expression, and the induction

of immune responses that preclude repeated administrations.

AAVs have also been extensively investigated as a delivery vector for CF gene therapy. Similar

to AdVs, AAVs are non-pathogenic and remain episomal in transduced cells in vivo66. A number

of CF gene therapy trials using AAV vectors have been conducted since 199867–72. Although

initial trials demonstrated sufficient safety profiles, they were ultimately discontinued due to lack

of efficacy in improving lung function. More recent studies utilizing pseudotyped lentiviral

vectors have demonstrated some efficacy in delivering genes to the airway of mice and pigs73–77,

however these have yet to be investigated in clinical trials.

Due to the evident limitations associated with the aforementioned vector delivery systems, our

group and others have been investigating the utility of helper-dependent adenoviral (HD-Ad)

vectors. HD-Ad vectors are based on AdV serotype 5 with all viral coding sequences deleted78.

Accordingly, these vectors exhibit stable, long-term transgene expression due to attenuated

- 7 -

immunogenicity, while maintaining adenoviral tropism for epithelial cells79. Our group has

demonstrated that CFTR knockout mice are protected from acute lung infection following HD-

Ad delivery of the CFTR gene to mouse airways80. Furthermore, this delivery elicited stable

transgene expression for up to 15 weeks in the absence of measurable pulmonary inflammation.

HD-Ad vectors have also been used to deliver transgenes to human primary epithelial cells81,82,

as well as airway epithelia and submucosal glands of pigs in vivo83. However, a major challenge

here is that the airway epithelium is a dynamic tissue that undergoes slow yet constant turnover,

and therefore therapeutic transgene expression will be limited accordingly84. This limitation

illuminates a more broadly diffuse challenge in gene therapy applications. That is, the efficacy of

safer, non-integrating vectors is limited by the half-life and finitude of their cognate target cells.

Furthermore, although stable transgene expression can be attained using integrating viral vectors,

untargeted integration of synthetic genetic elements into the host genome carries risks of

transcriptional silencing, insertional mutagenesis, and oncogenic transformation41–43.

The requisite technology for ultimately surmounting these barriers have recently come to light

with the dawn of programmable nucleases. Such technologies endow the capacity to make

targeted and permanent site-specific genomic edits, such as knocking down endogenous genes, or

precisely knocking in exogenous transgenes. These technologies can therefore mediate permanent

transgene expression while avoiding insertional mutagenesis associated with integrating vectors.

Furthermore, our group has recently demonstrated the capacity for targeting porcine airway

stem/progenitor cells in vivo85. Our modern approach to CF gene therapy is thus to correct CFTR

mutations into the genome of airway stem/progenitor cells using programmable nucleases, in

order to permanently correct CF genetic defects and consequent pathophysiology.

1.3 Gene editing

Classical genetics in the time of Mendel and Morgan was empirically limited to spontaneous

mutations as the objects of study. The first hints of deliberate genetic manipulation came from

Muller (1927)86, and Auerbach & Robson (1947)87, who demonstrated that radiation and chemical

treatment, respectively, could potentiate the rate of mutagenesis. More advanced molecular

technologies subsequently harnessed inducible transposable elements to transfer genetic

- 8 -

information. However, these procedures were all limited to inducing undirected and stochastic

mutations. It wasn’t until the late 1970’s that the first targeted genomic manipulations were

produced in yeast and mammalian cells by homologous recombination (HR)83–87. HR can be

utilized to transfer genetic information between two homologous genetic elements, and will be

discussed in more detail in subsequent sections. Utilizing spontaneous HR for genomic

manipulation was targeted and precise, yet suffered great practical inefficiencies requiring

powerful selection and laborious characterization83–87.

The key to high-efficiency, precise genome editing was discovered serendipitously through the

study of DNA damage and repair, specifically DNA double-strand breaks (DSBs). It was found

that the generation of DSBs by ionizing radiation led to sister chromatid crossover events92, and

that DSBs are intentionally induced by meiotic cells to facilitate genetic recombination between

homologous sequences93. These findings precipitated experiments using specific endonucleases

to facilitate HR and targeted gene transfer, which ultimately nucleated modern genome

engineering94–96. Artificial nucleases can now be deliberately programmed, endowing researchers

with the technical capacity to introduce DSBs at predetermined genomic loci. Following a

targeted genomic DSB, the requisite DNA repair mechanisms can then be hijacked to introduce

specific genetic modifications. Briefly, gene knock-outs can be generated if the break is repaired

by error-prone non-homologous end-joining (NHEJ). In contrast, homology-directed repair

(HDR) – which encompasses a number of distinct mechanisms including HR – is required for

precise gene repair or site-specific integration of transgenes.

1.3.1 Zinc finger nucleases

Zinc fingers (ZFs) are the most common DNA-binding domains found in the human genome. ZFs

are comprised of many subclasses, of which Cys2His2 has been repurposed for genome

engineering. Cys2His2 ZF domains are comprised of tandemly repeating zinc-binding motifs of

30 amino acids, in which positions -1, 3, and 6 mediate direct contact with three contiguous base

pairs of DNA and contribute to the binding specificity of the motif97,98. Of the natural Cys2His2

domains, Zif-268 has proven the most stable and versatile framework upon to which to engineer

- 9 -

ZFs with novel binding specificities. Modular zinc finger domains with unique specificities can

then be linked in tandem in order to recognize novel DNA targets 9-18 bp in length99.

Fusion of ZF tandem repeats to an unbiased restriction endonuclease effectively generates a

programmable, site-specific endonuclease. This was first successfully done by Kim et al., who

fused the cleavage domain of the FokI endonuclease to a ZF array, in turn creating the first zinc

finger nuclease (ZFN)100. Because FokI is an obligate dimer, effective DSB generation requires

two ZFNs concomitantly targeting adjacent DNA loci. Despite the evident utility of ZFNs, target

site overlap and crosstalk between ZF domains significantly complicate the prediction of binding

specificities, and therefore the production of ZFNs targeting user-defined sequences101–103.

Therefore, as there is no simple universal code for matching ZFN amino acid sequence to target

DNA sequence, selection strategies are often required for the production of ZFNs104,105. ZFNs

also appear to have a limited targeting range106. Nonetheless, ZFNs were essentially the first

generation of programmable site-specific endonucleases, and accordingly were the first with

prospective biotechnological and clinical utility.

ZFNs have been successfully utilized for genome engineering in swine107, cattle108, zebrafish109,

mice110, as well as human embryonic stem cells (ESCs) and induced pluripotent stem cells

(iPSCs)111. Prospective clinical utility has been demonstrated for correcting haemophilia112 and

sickle-cell in mice113; as well as 1-antitrypsin deficiency in iPSCs114; and Parkinson’s disease-

associated alpha-synuclein (SNCA) gene in patient fibroblast-derived iPSCs115. These

technologies have since been largely replaced by more powerful and robust programmable

nucleases. However, there are a number of active and completed clinical trials based on the

pioneering ZFNs due to their chronological primacy. For example, Phase I clinical trials

(NCT00842634) have investigated the safety and efficacy of transplantation of autologous CD4+

T cells edited by ZFNs ex vivo to disrupt the CCR5 gene for the treatment of HIV-1116. However,

treatment elicited severe adverse effects in some patients and variable efficacies, as viremia was

undetectable in only a single patient by the endpoint of the trial. Other active Phase I clinical trials

include those for the treatment of mucopolysaccharidosis (MPS) (NCT02702115,

NCT03041324), and HPV-positive cervical intraepithelial neoplasia (NCT02800369).

- 10 -

1.3.2 TALENs

Transcription activator-like effector (TALE) proteins are critical virulence factors of the

phytopathogenic bacterial genus Xanthomonas, which mediate host cell reprogramming by

mimicking eukaryotic transcription factors117–120. TALEs are DNA-binding proteins characterized

by repeating arrays of 33-35 amino acids, where each array recognizes a single base pair. The

repeat-variable di-residue (RVD) region harbours two amino acids that determine the specificity

of base pair recognition within each repeat121–123. Furthermore, modular TALE repeats can be

linked to FokI in order to generate unique site-specific endonucleases that recognize programmed

DNA sequences, analogous to ZFNs. In contrast to ZFNs, however, the simplicity of RVD base

pair recognition enables straightforward theoretical design of TALENs for targeting user-defined

DNA sequences. TALENs have also demonstrated a greater DNA targeting range with improved

specificities124–126. The principle impediments of TALEN usage are large coding sizes (~10 kb)

and the repeating nature of the RVDs, which impede efficient assembly and packaging of dimeric

TALENs into size-constrained vectors for mammalian cell delivery127. However, investigators

have devised superior methods by which to assemble TALENs, and have concurrently

demonstrated that TALENs have the capacity to target virtually any DNA sequence128–130.

Accordingly, TALEN libraries targeting over 18,000 human protein-coding genes have been

constructed using high-throughput Golden-Gate cloning131.

As with ZFNs, TALENs have been broadly applied for biotechnology and biomedicine, as they

have been applied for genome editing in roundworm132, silkworm133, fruitfly134, frog135, cricket136,

zebrafish137, rat138, cow and swine139. Delivery of two TALEN pairs targeting the same

chromosome has been demonstrated to elicit large indels (>6 kb) in pigs139. These authors used

TALENs to generate mono- and biallelic LDLR knockout pigs as models of familial

hypercholesterolemia139. Interestingly, TALENs have also been used to disrupt particular loci in

the rice genome to confer resistance to pathogenic TALEs in Xanthomonas infection (the source

from which this technology was initially discovered)140. After demonstrating utility for genome

editing in human somatic125 and pluripotent stem cells141, TALENs were subsequently applied as

preclinical anti-viral therapies for HIV infection142 and hepatitis B143. Furthermore, Phase I

clinical trials have been approved for the use of TALENs to treat HPV-related malignant

neoplasms (NCT03226470, NCT03057912).

- 11 -

1.3.3 CRISPR

1.3.3.1 Discovery and mechanisms

When investigating chemical mutagenesis in the 1940s, Auerbach and Robson pronounced that,

“…it could be hoped that among chemical mutagens there might be some with particular affinities

for individual genes. Detection of such substances not only would be of high theoretical interest

but would also open up the long-sought-for way to the production of directed mutations.”87 It

turns out that “such substances” had already been discovered eons ago by archaea, and integrated

into an adaptive genetic network nested within an evolutionary arms race against infectious

plasmids and phages. These prokaryotic immune systems are now known as the clustered

regularly interspaced short palindromic repeats (CRISPR).

Although named differently at the time, CRISPR systems were initially discovered

serendipitously through investigating the genetic origins of alkaline phosphatase isozyme

conversion in Escherichia coli144. The authors noted a peculiar set of repetitive sequences with

high sequence homology interspaced by variable spacers downstream of their gene of interest.

This investigation was indeed the first encounter with CRISPR sequences. Advancements in DNA

sequencing technologies in the 1990s uncovered these repetitive sequences in myriads of bacteria

and archaea. Despite numerous theories, however, their function remained largely enigmatic. It

wasn’t until 2000 that Mojica et al. recognized that these sequences must be functionally

related145. Furthermore, accumulating genomic data eventually revealed CRISPR-associated (cas)

genes adjacent to the repetitive CRISPR loci. These cas genes harboured recognizable subunits

indicating functions in DNA repair and recombination, transcriptional regulation and

chromosome segregation. Accordingly, their proximal association suggested they were involved

in generating the CRISPR loci themselves146.

The functional role of CRISPR was first glimpsed independently by a number of groups who

discovered that the variable spacer regions separating the repeats were homologous to phage

DNA147–149. Importantly, they showed that phages are unable to infect host strains containing

CRISPR elements with homology to their genomes. These phenomena were also observed in the

dairy industry, where investigators were selecting industrial bacteria resistant to phage attack.

Here, the authors provided the first empirical evidence for the involvement of cas genes in the

- 12 -

generation of CRISPR loci, and additionally as the enzymatic effectors of acquired phage

resistance150. These discoveries demonstrated that CRISPR can generate inheritable genetic

libraries of foreign genetic material, which function as part of a cellular immune system akin to

eukaryotic RNA interference (RNAi). Empirical evidence validated this model in subsequent

years151.

The CRISPR-Cas systems have been categorized into two classes (I and II), and further

subdivided into six types (I-VI)152. Although these subcategories exhibit distinct enzymes and

nuanced biochemical pathways, they share the same overarching functional architecture:

adaption, expression, and interference. The adaption stage is dependent on Cas1 and Cas2, which

form a multiprotein complex and mediate integration of short regions foreign DNA in between

CRISPR repeats153. These sequences are known as ‘spacers’, and are dependent on the presence

of a neighbouring protospacer-adjacent motif (PAM) on the cognate foreign DNA. PAM motifs

are typically only a few nucleotides long, and they ensure correct target recognition while

avoiding host self-cleavage154,155. The expression stage is characterized by transcription of the

CRISPR array into a precursor transcript (pre-crRNA), which is subsequently bound by multi-

subunit CRISPR RNA (crRNA)-effector complexes156–158, or, in type II systems, by a single

multi-domain Cas9 protein159. Following complex formation, mature crRNA is generated via

RNA endonuclease activity160, or a mechanism involving concerted action of RNase III and trans-

activating CRISPR RNA (tracrRNA)161. These RNA-protein complexes are then responsible for

mediating the interference module. Specifically, the activated crRNA-effector complexes

recognize the cognate target DNA (i.e., a foreign virus or plasmid) through specific base pair

interactions between the target DNA and the homologous spacer region of the crRNA. Target

DNA cleavage is then facilitated by a variety of nuclease domains embodied within the crRNA-

effector complexes, thus degrading the foreign DNA159,162.

1.3.3.2 Genome engineering

CRISPR-Cas systems are evidently comprised of multiple interacting enzymes, auxiliary proteins,

RNA and DNA molecules. Amongst the variety of CRISPR-Cas systems, however, the type II

CRISPR-Cas9 system is distinctly parsimonious, requiring only tracrRNA, crRNA, Cas9

- 13 -

nuclease, and host factor RNase III as necessary and sufficient for target DNA cleavage in

vitro163,164 and in vivo165,166. Accordingly, type II CRISPR-Cas9 has emerged as the archetypal

system, and has been the most widely repurposed and engineered. In fact, Jinek et al. showed that

tracrRNA and crRNA can be combined into a chimeric single-guide RNA (sgRNA)163. This

effectively reduced the necessary components for Cas9-directed DNA cleavage and has therefore

increased the versatility for biotechnological applications. The modern genome engineering

toolkit now consists of ectopic Cas9 (typically purified and codon optimized from Streptococcus

pyogenes), and a sgRNA harbouring 20 nt of complementarity to the target DNA sequence.

Selection of target DNA sequences are limited only by the presence of a neighbouring PAM

sequence, which in the case of Cas9 is only ‘NGG’. This system is therefore highly versatile,

allowing programmable and directed DNA cleavage at virtually any region within the genome of

a variety of organisms.

CRISPR/Cas9 has numerous advantages over preceding technologies like ZFNs and TALENs.

Namely, both ZFNs and TALENs must be expressed as pairs for effective cleavage, and require

design and production of novel proteins for targeting distinct genomic loci. In contrast, a single

Cas9 and sgRNA are sufficient for DNA cleavage, and targeting novel loci simply requires the

production of new sgRNA sequences (~20 nt). The nature of RNA-guided targeting also allows

for robust multiplexing approaches by expressing multiple sgRNAs, whereas multiplexing with

ZFNs or TALENs was severely limited by cloning constraints. The diverse targeting capacities

and ease of CRISPR/Cas9 design and production thus render this technology superior to preceding

nucleases. Accordingly, once the requisite components for DNA cleavage had been understood163,

CRISPR/Cas9 was rapidly applied for genomic engineering in bacteria167, yeast168, as well as

mouse and human cell lines169,170. These technologies have also profoundly influenced transgenic

animal production in the livestock industry, as CRISPR/Cas9 has already been used to generate

transgenic swine171,172, cattle173, sheep174, and goats175,176.

1.3.3.3 CRISPR/Cas9 biochemistry and engineered Cas9 variants

Despite the evident utility of CRISPR/Cas9 for genome engineering, a number of salient

limitations are hindering its clinical applications, namely, specificity and efficiency. Accordingly,

- 14 -

a number of research groups have sought to engineer more specific and efficient Cas9 variants.

To these ends, an understanding of Cas9 structural biochemistry has proven indubitably useful.

Cas9 is a multi-domain structure which can be broadly dissected into its recognition (REC) and

nuclease (NUC) lobes. The REC lobe is essential for sgRNA and target DNA binding, and

evidently lacks structural homology with other known proteins and is therefore Cas9-specific177.

In contrast, the NUC lobe is comprised of RuvC and HNH nuclease domains, which share

structural homology with the superfamily of retroviral integrases such as Escherichia coli RuvC178

and Thermus thermophilus RuvC179; and with phage T4 endonuclease180 and Vibrio vulnificus

nuclease181, respectively. The NUC lobe also harbours the PAM-interacting (PI) domain, which

recognizes PAM sequences on the non-complementary strand and confers PAM specificity177.

Following sgRNA:Cas9 complex formation, the active ribonucleoprotein (RNP) complex scans

dsDNA for PAM sequences. PAM recognition then stimulates destabilization of the adjacent

dsDNA and subsequent R-loop formation182 (Fig. 1). During this process, the sgRNA:DNA

heteroduplex is stabilized within a positively charged groove at the interface between the REC

and NUC lobes of Cas9. As the Cas9 complex interrogates DNA for complementarity, sequential

extension of the sgRNA:DNA heteroduplex proceeds distally from the PAM site183. Successful

target recognition is highly dependent on complementarity with the first 10-12 nucleotides

directly adjacent to the PAM sequence, known as the sgRNA ‘seed region’163,169,170,184,185.

Accordingly, mismatches detected early in directional melting prematurely destabilize the ternary

complex and prevent Cas9 nuclease activation183. Following successful target recognition, the

HNH and RuvC nuclease domains cleave the complementary and non-complementary strands,

respectively163,177.

- 15 -

Figure 1. CRISPR/Cas9-directed genomic DNA cleavage. The active sgRNA:Cas9 RNP complex scans dsDNA for PAM sequences (green). PAM recognition stimulates destabilization of the adjacent dsDNA and R-loop formation ensues. The sgRNA:DNA heteroduplex is stabilized within a positively charged groove, and sequential extension of the heteroduplex proceeds distally from the PAM site. sgRNA complementarity with the first 10-12 nucleotides adjacent to the PAM sequence (the seed region) is essential for successful target recognition and nuclease activation. Correct sequence complementarity stimulates the HNH and RuvC nuclease domains to cleave the complementary and non-complementary strands, respectively.

1.3.3.3.1 dCas9

The conservation of RuvC and HNH Cas9 nuclease domains has greatly assisted directed

mutagenesis to generate novel Cas9 functions. Notably, D10A and H840A mutations in the RuvC

and HNH domains, respectively, inactivate the catalytic activity of each respective nuclease. This

has allowed for the generation of the catalytically-inactive dead-Cas9 (dCas9) variant, as well as

Cas9 nickases. dCas9 was initially utilized for endogenous promoter binding and transcriptional

repression in Escherichia coli186 and human cells187. Since then, dCas9 has been widely

repurposed as a fusion protein platform for the recruitment of effector domains to targeted

genomic loci. For example, synthetic transcriptional activators constituted by dCas9-VP64 or -

p65 (subunit of nuclear factor kappa B (NF-B)) fusions have been successfully employed to

activate genes in human cells188,189. Researchers have recently used a similar approach to

simultaneously activate multiple endogenous neurogenic genes in transgenic mice, resulting in

the efficient conversion of astrocytes into functional neurons in vivo190. dCas9 has also been used

T A CCA TGGT

TGGTGACC

TAT C GGTCACA C

TGGTGACCTATACCGGTCACsgRNA

PAM

R-loopGG A GT GCA

complementary strand

RuvC

NGG

HNH

ACC

- 16 -

for targeted epigenomic modifications by fusing dCas9 to histone acetyltransferases (HATs)191.

Furthermore, fusions to the Krüppel-associated box (KRAB) mediates site-specific recruitment

of a heterochromatin-forming complex that facilitates histone methylation and deacetylation192.

dCas9 variants have also been employed to address off-target edits and increase the specificity of

Cas9-induced DNA cleavage. For these purposes, dCas9 has been fused to the unbiased FokI

nuclease. Because FokI is an obligate dimer, effective DSB induction is dependent on mutual on-

target activity of two dCas9-FokI fusions guided by two distinct sgRNAs to adjacent DNA

loci193,194. Although this approach reduces off-target effects, it comes at a cost of reduced

efficiency. A similar strategy has been demonstrated by multiplexing Cas9 nickases, where each

contain a single point mutation (either D10A or H840A), thus relying on mutual on-target activity

of both Cas9 variants for cutting195,196. Other efforts to increase Cas9 specificity have involved

directed mutagenesis to generate Cas9 variants with reduced DNA binding energies, which thus

require more stringent target interactions for nuclease activation. To this end, eSpCas9 has been

engineered by alanine mutagenesis to neutralize positively charged grooves that stabilize DNA

during target interrogation197. Furthermore, targeted alanine mutations that disrupt hydrogen

bonding with the DNA phosphate backbone have generated the SpCas9-HF variant with improved

specificities198. Distinct approaches to improving editing specificity include small molecule- and

light-inducible Cas9 variants199–201. These studies demonstrate that controlling the temporal

window within which Cas9 is active can substantially alleviate off-target edits.

1.3.3.3.2 Base pair editors

Significant engineering efforts have also been invoked to generate programmable single base-pair

editors. Pioneering work by Komor et al. generated the first of such enzymes by fusing dCas9 to

rAPOBEC1 (a cytidine deaminase), which became the first-generation base editor (BE1)202. This

was a significant step forward for genome engineering, as it allowed for directed C→T (or G→A)

base pair substitutions, while alleviating the necessity for DSBs. Although BE1 could generate

efficient substitutions in vitro, in vivo editing efficiencies were up to 36-fold lower. Second- and

third-generation BEs were engineered by adding a uracil DNA glycosylase inhibitor (UGI), and

using a Cas9 nickase instead of dCas9. BE3 has demonstrated superior utility, eliciting permanent

mutations in up to 75% of human cells with minimal indel formation (1%)202. Further

modifications have generated more efficacious fourth-generation base editors203. These

- 17 -

technologies have since been utilized for diverse applications, including targeted base editing in

mouse embryos204; wheat and maize205; as well as improving herbicide resistance in rice and

generating marker-free gene edits in tomato206.

1.3.3.3.3 sgRNA engineering

Another effort of Cas9 engineering has been to manipulate the chemical and structural properties

of the sgRNA, as means to maximize both cleavage efficiency and target specificity. It was

initially found that extending the canonical 20 nucleotide spacer region of the sgRNA does not

improve specificity196. In contrast, sgRNA reduction to 17 or 18 nucleotides can generally

ameliorate off-target effects while maintaining on-target cutting efficiencies for most sgRNAs

tested207,208. However, these truncated sgRNAs have proven incompatible with engineered Cas9

variants like SpCas9-HF due to marginal on-target efficiencies198. The influence of non-canonical

sgRNA nucleotide chemistry on Cas9 genome editing has also been investigated. For instance,

2’-O-methyl (M), 2’-O-methyl 3’phosphorothioate (MS), and 2’-O-methyl 3’thioPACE (MSP)

modifications of terminal nucleotides improves sgRNA intracellular stability, and significantly

increases Cas9 editing efficiencies in CD34+ HSPCs and human primary T cells209. However,

additional off-target activity accompanied these improvements. Further modifications to these

sgRNAs including interspaced 2’-fluoro groups increased stability and specificity but reduced on-

target efficiencies210.

1.3.3.4 CRISPR/Cas9 in genetics and disease modeling

In its brief history, CRISPR/Cas9 has already significantly impacted the way in which we study

genetic networks and the role of genetics in disease pathogenesis. For example, the explosion of

massive genomic data sets that has occurred in this century require powerful high-throughput

methods for annotating genomes. To this end, CRISPR has proven immensely successful for

conducting genome-scale knockout screens in mammalian cell cultures, eliciting substantial

phenotypic effects and high validation rates211–216. dCas9-derived base editors have also been

repurposed for generating diverse libraries of point mutations within targeted windows of

genomic loci, enabling high-throughput screening of genetically-diverse functional variants217,218.

The authors demonstrated the utility of such technologies by applying them to identify novel

- 18 -

mutations conferring resistance to the cancer therapeutics imatinib217 and bortezomib218. For these

high-throughput genetic screens, CRISPR supersedes chemical mutagenesis and RNAi screens.

As chemical mutagenesis is undirected, the genetic origins of measurable phenotypes is unknown,

and can be costly and laborious to investigate. Furthermore, although RNAi is high-throughput

and directed, this approach suffers from low signal-to-noise ratios due to high off-target activity

and incomplete gene knockout219–221.

CRISPR systems have also made significant contributions to stem cell research, endowing

researchers with the capacity to effectively knockout and knock-in genes in ESCs and iPSCs222–

225, as well as transcriptionally regulate genes in ESCs226,227, NSCs228, and iPSCs229. Notably, due

to the simplicity of multiplexing sgRNAs, many of these studies demonstrate the feasibility of

simultaneously modulating multiple genetic pathways. CRISPR has also been extensively applied

for disease modeling and drug screening in ESCs230,231, iPSCs232,233, and a number of organoids.

For example, investigators have utilized CRISPR to generate mutant kidney organoids234, as well

as intestinal organoids as disease models to study congenital disorders235 and colorectal cancer236–

239.

1.3.3.5 CRISPR/Cas9 in gene and cell therapies

The first applications of these tools in human cells were hastily welcomed as the dawn of

biomedical utopias in which human disease will be eradicated under the auspices of

CRISPR/Cas9. Although many of these claims were facile and hyperbolic, these rapidly evolving

technologies will undoubtedly have a profound influence on biology and medicine. The low cost

and technological simplicity of CRISPR/Cas9 endows researchers with unprecedented capacities

for manipulating the genetic profiles of clinically-relevant PSCs, autologous primary cells, and

diseased tissues.

CRISPR/Cas9 has already begun to show promise as powerful gene editing tool for gene and cell

therapy applications, as evidenced by several preclinical studies and recently emerging clinical

trials. For example, LNPs and AAV vectors have been used to deliver Cas9 mRNA and sgRNAs

to the liver of a mouse model of tyrosinemia, achieving up to 6% of genetically corrected

hepatocytes and full rescue of liver damage48. Furthermore, LNPs have also been used to deliver

- 19 -

CRISPR to the murine inner ear, with the capacity to edit the genome of 20% of hair cells47.

Interestingly, the power of CRISPR-induced indels has been demonstrated by the correction of a

splicing defect in the LAMA2 gene240. This gene encodes the D2 chain (Lama2) of the laminin-

211 complex expressed in muscle and Schwann cells. Splicing defects in Lama2 result from exon

skipping, which in turn produce truncated proteins and ultimately give rise to congenital muscular

dystrophy type 1A (MDC1A). Using CRISPR, the authors were able to excise the intronic region

containing the mutation, in turn creating a functional donor splice site to create full-length Lama2

protein. Furthermore, AAV-mediated delivery of this CRISPR system in a mouse model of

MDC1A resulted in substantial improvements in neuromuscular pathology240.

These technologies have also made their way into therapies for cardiovascular diseases. Most

notably, investigators have applied the power of CRISPR-induced indels to disrupt the PCSK9

gene. Naturally occurring loss of function mutations in PCSK9 reduce circulating LDL levels and

attenuate the risk of coronary heart disease by up to 88%241. For these reasons, Ding et al. injected

AdV carrying the requisite CRISPR components targeting exon 1 of hepatic PCSK9 into mice242.

Significant hepatic PCSK9 disruption resulted in a ~90% reduction in circulating PCSK9, and a

consequent ~40% reduction in total blood cholesterol. These studies have since been successfully

reproduced using AAV vectors243, as well as in humanized mouse liver in vivo244.

CRISPR has also made significant contributions to cancer therapeutics, namely chimeric antigen

receptor (CAR) T cell immunotherapies, which have been transformative for the treatment of

refractory blood cancers245. Directed gene manipulations facilitated by CRISPR technology have

generated synthetic CAR T cell variants with enhanced anti-tumour efficacy in vivo246,247, as well

as engineered iPSC-derived natural killer (NK) cells for the treatment of solid tumours248.

Advancements in CRISPR-edited CAR T cell therapies have since translated into clinical trials

for the treatment of relapsed or refractory leukemia and lymphoma (NCT03398967). CRISPR has

additionally been used to disrupt the PDCD1 gene encoding programmed death-1 (PD-1) receptor,

an immune checkpoint receptor which downregulates cytotoxic T cell proliferation and activity.

The activating ligand for this receptor can be overexpressed in tumor cells to subvert the immune

system. Accordingly, CRISPR-induced disruption of PDCD1 in cancer patient-derived primary

T cells enhanced cytotoxic anti-tumor activity249, and in vivo injection of these modified T cells

improved the survival of tumor-bearing mice250. The efficacy of these treatments has since led

- 20 -

to clinical trials for the treatment of Epstein-Barr virus (EBV)-positive advanced stage

malignancies (NCT03044743).

As previously discussed (section 1.2), editing HSPCs for inherited blood disorders are a major

effort of gene and cell therapy applications. Autologous HSPCs can be extracted from patients,

edited ex vivo, and subsequently transplanted back into patients. Patient-derived autologous

HSPCs can now be electroporated ex vivo with CRISPR-expressing plasmids251, mRNA209, or

Cas9:sgRNA RNPs252. The use of RNPs presents a particularly attractive strategy, as it manifests

a ‘hit and run’ approach, effectively minimizing nuclease exposure and therefore toxicity and off-

target edits209. De Ravin et al. utilized this approach to correct mutations in CYBB, the gene

encoding the catalytic centre of NADPH oxidase, which gives rise to X-linked chronic

granulomatous disease (X-CGD)253. Specifically, this group edited X-CGD patient-derived

CD34+ HSPCs with the requisite CRISPR components, in turn restoring NADPH oxidase function

in myeloid cells differentiated from the extracted progenitors in vitro. Subsequent transplantation

of corrected HSPCs into NSG mice resulted in effective engraftment and sustainable production

of functional mature human myeloid and lymphoid cells. These experiments represent a

significant step forward from similar studies using ZFNs, which failed to attain appreciable

correction rates following transplantation into NSG mice113,254.

These applications have also extended into anti-viral therapy, in particular for disrupting the

CCR5 gene to generate HIV-resistant human primary T-cells255,256 and iPSCs142,257. Notably, Xu

et al. have shown that CRISPR can be used to disrupt the CCR5 gene in human CD34+ HSPCs,

and that subsequent transplantation into NPG mice can generate multi-lineage progeny and confer

resistance to HIV-1 infection258. These experiments have precipitated the approval of Phase I

clinical trials (NCT03164135).

Substantial empirical evidence has also emerged in support of our current model of CRISPR-

based CF gene therapy. Namely, CRISPR/Cas9 has been used to correct the 'F508 mutation in

CF patient-derived iPSCs259. Corrected iPSCs were able to differentiate into mature airway

epithelial cells exhibiting physiologically normal CFTR function. Moreover, intestinal stem cells

from 'F508 CF patients have been corrected ex vivo, and subsequently able to form clonally

expanded intestinal organoids with restored CFTR function260. These are indeed promising

results. However, genome editing of airway stem cells in vivo will require significant

- 21 -

improvements in CRISPR-mediated transgene knock-in efficiencies. For these purposes, it is

necessary to understand the DSB repair mechanisms used by mammalian cells to repair Cas9-

induced genomic breaks.

1.4 DNA double-strand breaks

As with all biological macromolecules, DNA is subject to destabilizing and decomposing forces.

However, as the hereditary molecule of life, accurate reconstitution of DNA in response to

structural insults is essential to the faithful reproduction of the cells, multicellular networks,

organisms, and species in which it encodes. Of the various types of damage DNA can endure,

DSBs are indeed the most genotoxic and structurally threatening. DSBs can arise due to

extraneous stimuli like ionizing radiation and radiomimetic drugs, or as a consequence of

intracellular reactive oxygen species (ROS) or replication fork collapse261. Other DSBs are

programmed and essential to the reproduction and normal physiology of an organism, such as

meiotic crossover and V(D)J recombination, respectively. In an analogous manner, artificial

nucleases can be programmed to induce DSBs for deliberate genome editing. However, in contrast

to bottom-up, genetically-encoded programmed DSBs, those induced for genome editing are top-

down, encoded by scientists and the social institutions thereof. Accordingly, our deliberate

genomic manipulations will not have been tested and refined by eons of evolutionary pressures.

Understanding the molecular consequences of our programmed DSBs is therefore pertinent to

ensuring they are repaired in a manner conducive to our desired outcomes.

Because DSBs are highly genotoxic lesions, employment of the appropriate repair mechanism is

tightly regulated and crucial for maintaining genome integrity. In mammalian cells, DSBs are

predominantly repaired by one of two pathways, broadly classified as NHEJ and HDR. NHEJ-

related pathways commonly repair DSBs by direct re-ligation of the broken ends following some

or no processing on the ends themselves. Although NHEJ pathways are ostensibly error-prone

due to end processing and template-independent repair, they are probably accurate in many

cases262. On the contrary, HDR encompasses various distinct subpathways, which share the

common feature of requiring a homologous DNA sequence as a template for repair. Accordingly,

it is largely accepted that HDR pathways are exclusively error-free.

- 22 -

1.4.1 Non-homologous end-joining

Following a genomic DSB, the kinases ataxia telangiectasia mutated (ATM), ATM and Rad3-

related (ATR), and DNA-dependent protein kinase catalytic subunit (DNA-PKcs) rapidly

phosphorylate chromatin resident histone H2AX surrounding the break site263–266. This generates

H2AX, which acts as a DNA damage signal. In resting or quiescent vertebrate cells (G0/G1), the

Ku70/Ku80 heterodimer (Ku) is first to bind to free DNA ends generated by a DSB, likely due to

its shear abundance (~400,000 molecules/cell) and strong dissociation constant (~10-9 M)267,268.

However, Ku competes for DNA binding with the less abundant MRE11:RAD50:NBS1 (MRN)

complex, which promotes alternative end-joining pathways in G1269–271. In canonical NHEJ, the

Ku:DNA complex serves as a scaffold for the recruitment of additional repair proteins. The

Artemis:DNA-PKcs complex rapidly binds to Ku:DNA ends. Subsequent autophosphorylation of

Artemis:DNA-PKcs regulates access of other NHEJ proteins and activates the nucleolytic activity

of Artemis, thus initiating DNA end processing272–274. End processing functions to generate short

regions of microhomology (4 nucleotides) to serve as a template for ligation. Polymerases pol µ

and pol , which are capable of flexible template-independent and -dependent polymerase

activity, respectively, are then recruited to extend the processed DNA ends275,276. The

XLF:XRCC4:DNA ligase IV complex subsequently mediates ligation of the two DNA ends (Fig.

2). This enzymatic complex is a flexible ligase with the capacity to ligate complex and

incompatible DNA ends across gaps277. Depending on the chemistry of the break, additional

auxiliary proteins may be required for repair, including polynucleotide kinase (PNK), aprataxin,

and tyrosyl DNA phosphodiesterase 1 (TDP1)278.

Although described here as constrained and sequential mechanistic steps, NHEJ is more

accurately depicted as a semi-haphazard process involving malleable iterations of nuclease,

polymerase, and ligase activity278. It is evident that these enzymatic pathways and the proteins

involved are necessarily flexible. That is, it has proven evolutionarily adaptive to employ robust

biochemical pathways capable of repairing a diverse range of broken DNA substrates, as these

pathways are largely conserved from yeast to vertebrates.

- 23 -

1.4.2 Homology-directed repair

HDR encompasses a number of distinct repair subpathways, namely HR, synthesis-dependent

strand annealing (SDSA), and break-induced replication (BIR). These pathways are commonly

initiated by the generation of H2AX surrounding the DSB site, which function to recruit mediator

of DNA damage checkpoint 1 (MDC1) and the MRN complex279,280. These complexes in turn

scaffold the break site to recruit additional HDR factors. All HDR pathways are similarly

dependent on the generation of long regions of single-stranded DNA (ssDNA) surrounding the

break site. Specifically, ssDNA generation is catalyzed by the endonucleotic activity of the MRN

complex, which itself is dependent on the activity of carboxyl-terminal binding protein (CtBP)-

interacting protein (CtIP)281–283. Following MRN endonucleotic cleavage, exonuclease resection

proceeds bi-directionally via MRN 3’ – 5’exonuclease activity, and long range (~1-3 kb) 5’ – 3’

resection mediated by the redundant activities of EXO1 and BLM/DNA2 nucleases284–286. The

resulting ssDNA is then rapidly bound by replication protein A (RPA)287. The RPA coating

ssDNA is eventually replaced by RAD51 nucleoprotein filaments, aided by the concerted action

of PALB2, BRCA1, and the recombination mediator BRCA2288,289. RAD51 nucleofilaments are

the necessary substrates to search for and invade homologous DNA. Following homologous DNA

strand invasion, the subset of HDR pathways begin to diverge (Fig. 3).

Canonical HR is a consequence of both strands of the DNA break invading the homologous

template. Subsequent polymerization generates joint DNA molecules bearing a double-Holliday

junction (HJ), which can undergo considerable branch migration before dissolution. Double-HJ

dissolution and consequent separation of the conjoined DNA molecules is dependent on the

mutual activity of topoisomerase III and the BLM:RMI1:RMI2 (BTR) complex290. Processing

by BTR gives rise to non-crossover events that avoid loss of heterozygosity by inter-homologue

exchanges291,292. In contrast, if the double-HJs are resolved by the endonucleases MUS81-EME1,

SLX1-SLX4 and GEN1, crossovers are more likely to occur293.

SDSA can occur if one or both strands of the DNA break invades the homologous template, from

which extended D-loops are formed instead of double-HJs. Following homology-dependent

polymerization of the invading strand, D-loops are processed by helicases and the invading strand

is subsequently re-ligated to the other end of the DSB break294. In contrast, BIR results exclusively

from one-end invasion, and replication can proceed for hundreds of kilobases until the end of the

- 24 -

chromosome295,296. Another DSB repair pathway distinct from both HDR and NHEJ is

microhomology-mediated end-joining (MMEJ), which is highly error-prone. Although

categorically distinct, MMEJ shares similar features with HDR in that it requires CtIP-dependent

MRN nucleotyic action. In contrast to HDR, however, MMEJ ensues following limited resection

by MRN and subsequent alignment and ligation of short ssDNA ends, in the absence of any

homologous strand invasion297 (Fig. 2).

1.4.3 DSB repair pathway choice

The choice of DSB repair mechanism is largely dependent on cell cycle phase298. More

specifically, due to the dependence on a homologous sequence, HDR pathways are restricted to

S/G2 when a sister chromatid template is available for repair. On the contrary, NHEJ pathways

are active throughout the cell cycle. The cell cycle-regulated control of these repair pathways is

largely modulated by cyclin-dependent kinases (CDKs), which themselves are master regulators

of cell cycle progression. CDKs have extensive regulatory control over DNA end resection, which

is generally viewed as the committed step for HDR and functions as the primary regulatory node

for DSB repair pathway choice299.

p53-binding protein 1 (53BP1) is a key proximal regulator of DSB repair pathway choice. 53BP1

effectively inhibits end resection, and therefore HDR, principally through its concerted action

with Rap1 interacting factor 1 homolog (RIF1). In response to DSBs in G0/G1, ATM kinase-

mediated phosphorylation of 53BP1 induces recruitment of RIF1 to DSB sites. RIF1 DSB foci in

turn inhibit the accumulation of BRCA1 at DSBs, which ultimately inhibits end resection and

promotes repair by NHEJ300–304 (Fig. 2). The HDR-inhibitory action of 53BP1 is counteracted by

upstream CDK activity as cells enter S phase. More specifically, CDK-mediated phosphorylation

of CtIP in S/G2 promotes CtIP:BRCA1 complex formation, in turn relieving the 53BP1-

dependent, RIF1-mediated inhibition of end resection305. This regulation is also dependent on

CDK-mediated phosphorylation of BRCA1, which promotes UHRF1-mediated ubiquitylation

and consequent degradation of RIF1306. Chromatin remodeling has also recently been implicated

in this regulatory axis. More specifically, CDK- and ATM-mediated phosphorylation of the

chromatin remodeler Cockayne syndrome group B protein (CSB) promotes nucleosome

- 25 -

disassembly, which consequently inhibits and promotes RIF1 and BRCA1 accumulation,

respectively307 (Fig. 3).

In addition to the relief of 53BP1:RIF1 inhibition, positive stimulatory regulation is also required

to promote end resection. In this respect, CtIP has been recognized as a critical regulatory hub for

DSB repair pathway choice in mammalian cells, whose activity is a necessary prelude for end

resection and therefore HDR298,299,308–310. The involvement of CtIP in HDR regulation is

exemplified by low steady-state levels in G1 maintained by ubiquitin-dependent, proteasome-

mediated degradation311–313. CtIP degradation is subsequently suppressed during the G1/S

transition, in concordance with the upregulation of HDR upon S-phase entry. Its localization at

DSBs is also highly regulated. In particular, RNF138 ubiquitin ligase activity displaces Ku from

DSB sites while recruiting CtIP, and this process is necessary for the inhibition of NHEJ and

promotion of HDR314,315.

CtIP is also heavily phosphorylated by upstream kinases in S/G2, in particular by CDK1. For

example, CDK1 phosphorylates CtIP at Thr847 and Ser327 in S/G2, in turn facilitating its

interaction with BRCA1, promoting end resection, and ultimately stimulating HDR282,316–318.

CDKs also phosphorylates CtIP at 5 other residues, termed 5mCDK. These modifications do not

directly promote HDR, but instead facilitate subsequent CtIP phosphorylation at T859 by the

ATM kinase283. T859 phosphorylation promotes CtIP to stimulate end resection and recruit BLM

and EXO1, which in turn catalyze the extended end resection necessary for HDR283. In addition

to phosphorylation, CtIP is also constitutively acetylated at Lys432, Lys526, and Lys604319.

Following DNA damage, CtIP is deacetylated by sirtuin 6 (SIRT6), in turn promoting the ability

of CtIP to mediate end resection and therefore HDR319. Furthermore, loss of SIRT6 activity results

in severe HDR defects in human cells, exemplifying its role in stimulating CtIP-mediated DSB

end resection319,320. In fact, artificially expressing SIRT6 has been shown to rescue the decline of

HDR associated with replicative senescence320.

Limited resection by MRN:CtIP is sufficient for promoting the error-prone MMEJ pathway.

Extended end resection, however, mediated principally via the redundant activities of EXO1 and

BLM/DNA2 nucleases, is necessary for commitment to HDR pathways284–286. Accordingly,

EXO1 exonuclease activity is stimulated by CDK-mediated phosphorylation in S/G2, specifically

at residues Ser639, Thr732, Ser815, and Thr824321. The regulation of DSB repair proteins also

- 26 -

appears to be mutually interdependent, in that the activity of EXO1 is also stimulated by BLM,

MRN, and RPA286,322. These regulatory pathways effectively restrict long-range end resection to

S/G2, and therefore serve as additional levels of HDR regulation.

- 27 -

Figure 2. NHEJ is used to repair DSBs in G1 when CDK activity is low. In G1, ATM-mediated phosphorylation of 53BP1 promotes 53BP1 interactions with RIF1 and PTIP. These interactions in turn inhibit the accumulation of HDR factors such as CtIP, BRCA1, EXO1, BLM and DNA2 at DSB sites. Ku competes for binding with the MRN complex at DSB ends. Stable Ku binding invariably leads to NHEJ with the concerted actions of Artemis:DNA-PKcs, pol µ and , and DNA ligase IV. Limited end processing mediated by MRN:CtIP can lead to mutagenic MMEJ.

NHEJ MMEJ

Artemis

DNA-PKcs

pol µ

pol

Modified from Hustedt & Durocher; Nat Cell Biol. (2017) 19:1-9

- 28 -

Figure 3. HDR is used to repair DSBs in S/G2 when CDK activity is high. CDK-mediated phosphorylation of BRCA1 and CtIP promotes their accumulation at DSB sites by relieving the HDR inhibitory effects of 53BP1:RIF1. Chromatin remodelling directed by pCSB further promotes BRCA1 assembly while inhibiting RIF1. Active CtIP and MRN catalyze rate-limiting short-range end resection, and CDK-mediated phosphorylation of EXO1 stimulates long-range end-resection. RAD51 replaces RPA on ssDNA and catalyzes homologous strand invasion, which can subsequently result in SDSA, HR or BIR

OR

Non-crossover

Crossover HR SDSA BIR

CDK

CDK activity high

Modified from Hustedt & Durocher; Nat Cell Biol. (2017) 19:1-9

CSB

- 29 -

1.5 Enhancing gene targeting efficiency by perturbing DSB

repair

The power of programmable nucleases for genome editing lies in their ability to generate directed

DSBs. Furthermore, the precise nature of the edit depends on the repair pathway employed to

repair the DSB. Gene knockouts can be generated when NHEJ is used to repair the break, as this

pathway is prone to generating small mutations or indels. In contrast, due to the dependence of

HDR on a homologous template, this pathway can be harnessed to insert exogenous DNA

sequences into precise genomic locations. By flanking transgene cassettes – such as CFTR, for

example – with loci-specific sequence homology, exogenous DNA can be used as the template

for HDR in order to knock-in desired genetic elements. This approach is broadly referred to as

gene targeting.

The principle impediment to gene targeting is the inherently low frequency of HDR events. That

is, due to the fact that NHEJ is the predominant DSB repair mechanism, the efficiency of precise

gene targeting is exceedingly low. In mammalian cell lines, primary cells, PSCs and zygotes,

precise gene targeting efficiencies can range anywhere from 0.1-20% (with few exceptions

beyond this range), depending on the cell type, the genomic locus being targeted, method of

delivery, as well as the size and chemistry of the repair template. This represents a considerable

barrier for effective in vivo gene therapy, and more specifically, our CF gene therapy approach to

knock-in the CFTR gene into airway stem cells. Here, the low efficiency of gene targeting is

further compounded by practical barriers to gene delivery and the host immune response.

Accordingly, our approach requires significantly higher gene targeting efficiencies (GTE) to be

clinically useful.

It is worth recalling here the broadly applicable preclinical efficacies of CRISPR/Cas9 outlined

previously. It should be noted that the vast majority of those cases were using CRISPR/Cas9 to

generate knockouts by NHEJ. In the cases where HDR was necessary, small insertions or low

efficiencies sufficed for the application. Due to the evident limitations associated with

CRISPR/Cas9 gene targeting by HDR, a number of groups have sought to improve current

methods and enhance targeting efficiencies. The most widely used approach has been to

- 30 -

administer the small molecule SCR7, a DNA ligase IV inhibitor, in combination with the requisite

factors for CRISPR-mediated gene targeting (Cas9, sgRNA, and a HDR template). As previously

discussed, DNA ligase IV is responsible for ligating broken ends in DSB repair by NHEJ.

Accordingly, by inhibiting this enzyme, investigators have been able to upregulate compensatory

HDR and therefore GTE anywhere from ~2 to 20-fold in mammalian cell lines and zygotes323,324.

However, these results are often not reproducible325–329. Analogous approaches to small molecule

inhibition of NHEJ have been demonstrated using NU7441 and Ku-0060648 to inhibit DNA-

PKcs, resulting in ~2 to 4-fold enhancements in human cell lines and mouse embryonic fibroblasts

(MEFs)330. The RAD51-stimulating small molecule RS-1 can also elicit up to 5-fold

enhancements in human cell lines325 and rabbit embryos328. Furthermore, the small molecules

Brefeldin A and L755507, an ER-Golgi transport inhibitor and a E3-adrenergic receptor agonist,

respectively, enhance gene targeting efficiency ~2 to 9-fold in mammalian cell lines and hPSCs

by unknown mechanisms331.

The cell cycle-regulated control of DSB repair pathway choice also presents an opportunity to

potentiate the use of HDR in response to CRISPR/Cas9-generated DSBs. There are several well-

established mechanisms for synchronizing proliferating cells in specific cell cycle phases. Yang

et al. have shown that the use of nocodazole to synchronize cells in G2/M can enhance gene

targeting efficiency ~2 to 4-fold in human cell lines332. This approach has also demonstrated utility

in hPSCs and neural precursor cells329. Another interesting strategy was to fuse Cas9 to a fragment

of the human Geminin peptide, which is degraded by the proteosome pathway in G1. This

effectively restricts Cas9 expression to S/G2 when HDR is most active, in turn enhancing GTE,

albeit modestly326. Other approaches to enhancing CRISPR/Cas9 GTE have focused on the nature

of the donor template. More specifically, a number of studies have demonstrated greater targeting

efficiencies in mice333–335 and rats336–338 using ssDNA donors as opposed to canonical dsDNA

donor templates.

- 31 -

Chapter 2 Hypothesis and Objectives

2.1 Rationale

Researchers have evidently made significant progress in enhancing CRISPR/Cas9 GTE. It should

be noted that despite these improvements, large insertions (like those necessary to knock-in

CFTR) are more resistant to HDR enhancements. Furthermore, many of these advancements have

come with the use of small molecules, which cannot be packaged into viral vectors and are

therefore less amenable to in vivo gene therapy. Genetically-encoded factors would be more

beneficial for these purposes. In this respect, Orthwein et al. has shown that concurrently

overexpressing a CtIP and PALB2 mutant can enhance CRISPR/Cas9 GTE up to 4-fold in non-

dividing U2OS 53BP1 cells339. This study provided proof of principle for genetically-encoded

means by which to perturb DSB repair and enhance CRISPR/Cas9 GTE, which could be

effectively applied to viral-based gene therapies.

2.1.1 CDK1

Aforementioned studies have evidently demonstrated the utility of perturbing DSB repair pathway

choice as effective means by which to enhance CRISPR/Cas9-mediated gene targeting. In line

with this reasoning, overexpressing CDK1 is an attractive strategy to increase GTE due its

overwhelming control over the balance between HDR and NHEJ.

2.1.2 Post-translational modification (PTM)-mimics

An alternative strategy for increasing GTE would be to specifically harness the influence of CDKs

on DSB repair pathway choice, without directly altering CDK expression or activity. This can be

accomplished by specifically overexpressing the relevant phospho-products of CDK catalysis that

- 32 -

promote HDR. Many of the CDK DSB repair substrates up-regulate HDR and/or inhibit NHEJ in

response to CDK-mediated phosphorylation of specific residues. Furthermore, mutating such

residues to Asp or Glu can generate ‘phospho-mimics’ that recapitulate the activity of the

phosphorylated proteins. In addition to phosphorylation, DSB repair proteins endure other PTMs

such as acetylation and ubiquitylation, which also regulate their capacity to promote HDR in a

cell cycle-specific manner. Importantly, mutating acetylated and ubiquitylated residues to Arg can

generate DSB repair proteins that mimic their active states, regardless of cell cycle position. These

properties make phospho-, as well as acetylated- and ubiquitylated-mimics ideal candidates for

effectively potentiating GTE. Crucial HDR proteins that can be ectopically expressed as PTM-

mimics are outlined in detail below and summarized in Table 1.

2.1.3 CtIP

As previously outlined, CtIP is recognized as an essential regulatory hub for DSB repair pathway

choice in mammalian cells. CtIP activity is negatively regulated in resting cells by ubiquitin-

dependent, proteasome-mediated degradation311–313. However, these studies have shown that

mutating ubiquitylated residues S276, T315, and Y842 to alanine can block ubiquitylation, in turn

generating a hyperactive CtIP variant that constitutively promotes HDR in human cell lines. The

relief of G1 ubiquitin-directed, proteasome-mediated degradation of CtIP is endogenously elicited

as cells enter S phase. CtIP is then activated by PTMs in S/G2, namely by CDK and ATM-

mediated phosphorylation and SIRT6-catalyzed deacetylation. It has been shown that such

activity can be effectively mimicked by site-directed mutagenesis to artificially promote HDR

throughout the cell cycle. For example, the requirement of Thr847 phosphorylation can be

circumvented by a CtIP phospho-mimic in which Thr847 is replaced by Glu (T847E). This mutant

demonstrates normal end resection and HDR in the absence of CDK-mediated phosphorylation318,

and exogenous expression of CtIP T847E increases aberrant HDR while inhibiting NHEJ305,317.

Furthermore, CDK-mediated Ser327 phosphorylation promotes CtIP association with BRCA1,

which is necessary to promote end resection316,340. A Ser327 phospho-mimic (S327E) would

therefore presumably have similar HDR-promoting activities. In addition, 5mCDK CtIP

phosphorylation facilitates subsequent ATM-mediated T859 phosphorylation of CtIP, which in

- 33 -

turn stimulates end resection and the recruitment of additional necessary HDR factors283. Wang

et al. has demonstrated that a T859E CtIP phospho-mimic bypasses the requirement of 5mCDK

phosphorylation, and can alleviate the G1-repression of HDR283.

In addition to phosphorylation, CtIP is also dependent on SIRT6 deacetylase activity to promote

end resection319. Importantly, a CtIP deacetylated mimic, in which Lys432, 526, and 604 are

mutated to Arg (3KR), relieve the requirement of SIRT6-dependent deacetylation319. This CtIP

3KR mutant would presumably have a greater propensity to promote HDR than its wild-type

counterpart.

CtIP evidently endures multiple levels of PTMs that control its ability to promote HDR in

accordance with cell cycle phase and DNA damage. It appears that each individual modification

contributes to the activation of CtIP, where the collective influence of specific PTMs results in

robust HDR-promoting activity. Furthermore, the studies outlined above demonstrate that these

modifications can be effectively mimicked by expressing an artificial CtIP harbouring multiple

mutations.

2.1.4 EXO1

Following CtIP/MRN-mediated short-range DSB end resection (~50-100 bp), EXO1 is recruited

to the partially-resected ends to catalyze long-range end resection (~1-3 kb), a process that is

necessary for HDR341. Similar to CtIP, EXO1 exonuclease activity is stimulated by CDK-

mediated phosphorylation in S/G2, specifically at residues Ser639, Thr732, Ser815, and

Thr824321. This effectively restricts long-range end resection to S/G2, and therefore serves as an

additional level of HDR regulation. Furthermore, Tomimatsu et al. demonstrated that an EXO1

phospho-mimic (EXO1-4D) not only rescues the HDR defect associated with CDK inhibition, but

also significantly up-regulates HDR in wild-type cells321.

- 34 -

Table 1. DSB repair proteins and associated mutations Protein Mutations CDK1 CtIP-WT

- -

CtIP-3EA S327E, T847E, T859E, S276A, T315A, Y842A CtIP-M S327E, T847E, T859E, K432R, K526R, K604R CtIP-M2A S327E, T847E, T859E, K432R, K526R, K604R, S276A, T315A CtIP-M3A S327E, T847E, T859E, K432R, K526R, K604R, S276A, T315A, Y842A EXO1-4D S639D, T732D, S815D, T824D

2.2 Hypothesis

Overexpressing CDK1, as well as CtIP and EXO1 variants can enhance the efficiency of

CRISPR/Cas9-mediated precise gene targeting.

2.3 Objectives

1: Develop an assay to measure gene targeting efficiency.

2: Assess the utility of CDK1, as well as CtIP and EXO1 variants for enhancing CRISPR/Cas9

gene targeting efficiency.

3: Generate an IB3-1 cell line to verify precise gene targeting by a colour-switch assay using

ssDNA donors.

- 35 -

Chapter 3 Materials and Methods

3.1 Plasmids

The CRISPR/Cas9 plasmid for the mCherry AAVS1 integration assay was generated by cloning

the sgRNA (5’-ACCCCACAGTGGGGCCACTA-3’) into pSpCas9(BB)-2A-GFP (PX458)

(Addgene #48138) by conventional restriction cloning. The homology arms on the promoterless

mCherry donor construct were obtained by PCR from BAC clone: RP11 384-G4. The splice

accepter (SA) and T2A sequence were added upstream of mCherry-poly(A) (from pTRE3G-

mCherry) by PCR. The homology arms and SA-T2A-mCherry-p(A) cassette were then cloned

into an empty pSEAP backbone342 by In-Fusion cloning (In-Fusion HD Cloning Plus, ClonTech,

cat. #638911). The lacZ plasmid was constructed as previously described342. The CDK1 plasmid

was obtained from Addgene (Cdc2-HA, #1888). CtIP and EXO1 cDNAs were obtained from

Transomic Technologies (cat. #BC030590-seq, and BC007491-seq, respectively). FLAG tags

were inserted in frame with each cDNA by PCR. Each cDNA-FLAG cassette was then inserted

under the control of a chicken -actin (CBA) promoter in an empty pSEAP backbone342 by In-

Fusion cloning. Mutagenesis was performed using QuikChange Lightning Multi Site-Directed

Mutagenesis kit (Agilent Technologies, cat. #210515).

The CRISPR/Cas9 plasmid used for generating the IB3-1 cell line was obtained from OriGene

Technologies (cat. #GE100002) containing the sgRNA (5’-GAAGGTGGCCAACCGAGCTT-3’)

targeting exon1 of the CFTR gene. The donor plasmid harbouring the EGFP-puromycin cassette

flanked by CFTR homology was custom made from OriGene Technologies on a pUC vector. The

CRISPR/Cas9 plasmids for CMV-mCherry integration into the genomic EGFP-CFTR locus was

generated by cloning the sgRNAs (#1: 5’-AGCACTGCACGCCGTAGGTC-3’;

#2: 5’-CTCGTGACCACCCTGACCTA-3’; #3: 5’-GGGCACGGGCAGCTTGCCGG-3’) into

pSpCas9(BB)-2A-GFP (PX458) (Addgene #48138) by conventional restriction cloning, and the

GFP tag was removed by EcoR1 digestion. The donor plasmid designed to integrate CMV-

mCherry into the EGFP-CFTR genomic locus was assembled by restriction digestion of CMV-

- 36 -

mCherry-p(A) cassette, while the EGFP homology arms were obtained by PCR. The three

fragments were subsequently cloned into an empty pSEAP backbone342 by In-Fusion cloning. The

T7 promoter was inserted in the reverse orientation downstream of the right homology arm by

PCR.

3.2 Flow cytometry

3x105 HEK293 cells (cultured in MEM (Thermo Fisher Scientific, cat. #11095080) + 10% FBS

(Wisent Bioproducts, cat. #080-450) + 1% Pen/Strep (Thermo Fisher Scientific, cat. #15070063)

in a humidified 37 C, 5% CO2 incubator) were seeded into tissue culture-treated polystyrene 6-

well plates (Corning, cat. #353046), and transfected the following day (~70-80% confluency)

using jetPRIME (Polyplus-transfection, cat. #114-07) according to the manufacturer’s protocol

(jetPRIME:DNA = 2:1) (CRISPR/Cas9: 500 ng; Donor: 1 µg; lacZ, CtIP, EXO1, CDK1: 2 µg).

For SCR7-treated cells, SCR7 (Xcess Biosciences, cat. #M60082-2) was diluted to the indicated

concentrations in DMSO and added to the culture media 18 hr post-transfection. 48 hr post-

transfection, cells were either harvested for flow cytometry or passed 1/8 into new 6-well plates.

Passaged cells were subsequently passed 5 additional times over the course of 3 weeks before

harvesting for flow cytometry. When harvesting for flow cytometry, cells were detached by

incubating in 0.5 mL trypsin-EDTA (0.05%) (Thermo Fisher Scientific, cat. #25300062) for 1

min at 37 C (incubator), suspended in serum-containing media, and spun down at 220 g. Cells

were then washed with PBS (1X, pH 7.4), and subsequently re-suspended in PBS. Dead cell

staining was performed using LIVE/DEAD Fixable Violet Dead Cell Stain Kit (Thermo Fisher

Scientific, cat. #L34955) according to the manufacturer’s protocol for unfixed cells. Briefly, cells

were washed twice with PBS + 1% FBS, and re-suspended in 1 mL PBS + 1% FBS. Cells were

filtered through 70 µM nylon mesh into 5 mL round-bottom polystyrene tubes (Falcon, cat.

#352052). Flow cytometry was performed using an LSRII-CFI (Becton Dickinson), and data

analysis was done using FlowJo v.10.

- 37 -

3.3 Junction PCR

HEK293 cells were cultured, seeded, transfected and passaged as detailed in flow cytometry

methods (section 3.2). Following transfection and subsequent passaging for 3 weeks, cells were

trypsinized (as in flow cytometry methods), spun down at 220 g, and washed with PBS. Cells

were subsequently lysed in 500 µL of RIPA buffer (25 mM Tris-HCl pH 7.6, 150 mM NaCl, 1%

NP-40, 1% sodium deoxycholate, 0.1% SDS) + 650 µg/mL proteinase K (Thermo Fisher

Scientific, cat. #EO0491) + 100 µg/mL RNase A at 55 C with gentle shaking overnight. Genomic

DNA was purified by phenol-chloroform extraction and ethanol precipitation. Specifically, 1X

volume of phenol:chloroform:isoamyl alcohol (25:24:1, v/v) (Thermo Fisher Scientific, cat. #

15593031) was added to cell lysates, briefly vortexed, and spun down at 14,000 rpm for 2 min.

The top aqueous layer was extracted into a fresh microfuge tube. 0.5X volume of 7.5 M

ammonium acetate and 2X volume of 100% ethanol were added and incubated at -20 C for at

least 1 hr. The mixture was then spun down at 14,000 rpm at 4 C for 5 min and subsequently

washed with 1 mL 70% ethanol. Purified genomic DNA was re-suspended in Tris-EDTA (TE)

buffer (pH 8.0). 1 µg of genomic DNA was used as template in each PCR reaction, and PCR

products were run on non-denaturing 1% agarose gels. PCR primers illustrated in Fig. 5

(primer set #1: Fwd: 5’-GCTTTGCCACCCTATGCTGACAC-3’,

Rev: 5’- TGTGCACCTTGAAGCGCATGAACTC - 3’;

primer set #2: Fwd: 5’- TTAGCCACTCTGTGCTGACCACTCTG - 3’,

Rev: 5’-AGTCACCCAGAGACAGTGACCAAC - 3’). Band intensities were quantified using

ImageJ.

3.4 Generating the IB3-1-CFTR-EGFP cell line

3.4.1 Transfection

2x105 IB3-1 cells (cultured in LHC-8 (Thermo Fisher Scientific, cat. #12678017) + 10% FBS

(Wisent Bioproducts, cat. #080-450) + 1% Pen/Strep (Thermo Fisher Scientific, cat. #15070063)

in a humidified 37 C, 5% CO2 incubator) were seeded into a tissue culture-treated polystyrene

- 38 -

6-well plate (Corning, cat. #353046). The following day, seeded cells were transfected with

CRISPR/Cas9 targeting CFTR exon1 and the EGFP-puromycin donor DNA (1 µg each) using

FuGENE HD Transfection Reagent (Promega, cat. #E2311) according to the manufacturer’s

protocol (FuGENE:DNA=3:1).

3.4.2 FACS

Transfected cells were expanded to 10 cm plates and passaged for 2 weeks. One of the 10 cm

plates was trypsinized (as in flow cytometry methods, section 3.2) and spun down at 220 g. Cells

were then washed with PBS, and subsequently re-suspended in 1 mL PBS. Single EGFP+ cells

were then sorted into 10 collagen-coated (type I bovine collagen, Advanced Biomatrix, cat.

#5005), tissue culture-treated, polystyrene 96-well plates (Corning, cat. #C353227) containing

LHC-8 + 20% FBS + 1% Pen/Strep culture media.

3.4.3 Puromycin selection and limited dilution

Transfected cells were incubated in LHC-8 + 10% FBS + 1% Pen/Strep supplemented with 1

µg/mL puromycin for 2 weeks. Every 2 days, plates were washed with PBS (1X, pH 7.4) and

fresh puromycin-containing media added. Fluorescence microscopy was then used to identify

EGFP+, puromycin-resistant colonies. Sterile cylinders were placed around each individual

colony to allow for isolated trypsinization, re-suspension in serum-containing media

(supplemented with 1 µg/mL puromycin), and subsequent transfer to 48-well plate. These

colonies were expanded to 6-well plates. Each colony was then diluted in LHC-8 + 10% FBS +

1% Pen/Strep (supplemented with 1 µg/mL puromycin) to 5 cells/mL, and 100 µL pipetted into

96-well plates. Plates were monitored 12 hr later to mark wells which only contained a single cell.

These clones were allowed proliferate and then expanded to larger culture vessels. A total of 6

clones were successfully expanded and healthy at this stage.

- 39 -

3.5 Generating ssDNA donors

T7 in vitro transcription was performed using TranscriptAid T7 High Yield Transcription Kit

(Thermo Fisher Scientific, cat. #K0441) according to the manufacturer’s protocol using the CMV-

mCherry donor (targeting EGFP-CFTR) as a template. Reaction products were then digested with

1 µL DNaseI at RT for 15 min, followed by enzyme inactivation with 1 µL 25 mM EDTA at 65

C for 10 min. RNA was then purified using the PureLink RNA Mini Kit (Thermo Fisher

Scientific, cat. #12183018A) according to the manufacturer’s protocol for purification from liquid

samples. Purified RNA was then used for gene-specific reverse transcription with the SuperScript

IV Reverse Transcriptase (Thermo Fisher Scientific, cat. #18090010) according to the

manufacturer’s protocol. Products of this reaction were then digested with 1 µL RNaseH at 37 C

for 20 min to remove template RNA. RNaseH was inactivated by incubation at 65 C for 10 min.

The remaining ssDNA was purified using the PureLink RNA Mini Kit according to the

manufacturer’s protocol for purifying from liquid samples.

3.6 T7E1 Assay

2x105 IB3-1-CFTR-EGFP cells (from each of the 6 clones) were seeded into tissue culture-treated

polystyrene 12-well plates (Corning, cat. #C353225), and the following day transfected with

CRISPR/Cas9 (targeting EGFP-puromycin) (650 ng) and ssDNA mCherry donor (350ng) using

FuGENE HD Transfection Reagent (Promega, cat. #E2311) according to the manufacturer’s

protocol (Fugene:DNA = 3:1). 3 different CRISPR/Cas9 plasmids containing various sgRNAs

targeting different EGFP sequence motifs were tested for each clone (see plasmids methods,

section 3.1). 2 days post-transfection, transfectants were expanded to 6-well plates. 3 days later,

genomic DNA from each condition was harvested and purified using the GeneArt Genomic

Cleavage Detection Kit (Thermo Fisher Scientific, cat. #A24327) according to the manufacturer’s

protocol. PCR was used to amplify the EGFP sequence from genomic DNA. PCR products were

purified using the NucleoSpin Gel and PCR Cleanup Kit (ClonTech, cat. #740609). Purified PCR

products were subsequently denatured and randomly reannealed using a thermocycler (95 C for

5 min, 95→85 C (-2 C/sec), 85→25 C (-0.1 C/sec), 4 C hold). Randomly reannealed

- 40 -

amplicons were then digested with 1 µL of T7E1 (New England Biolabs, cat. #M0302) for 45

min at 37 qC, and subsequently run on a non-denaturing 2% agarose gel.

3.7 Data analysis

Graphical presentation and statistical analysis were performed using Prism 6 (GraphPad). All data

are presented as mean r SEM. Statistics were performed using unpaired, two-tailed Student’s t-

tests. Differences between means were considered statistically significant when p<0.05 and

p<0.01, and were represented graphically as * and **, respectively.

- 41 -

Chapter 4 Results

4.1 Flow cytometry assay for measuring gene targeting

efficiency

An assay to measure GTE requires two plasmid constructs: one for expressing CRISPR/Cas9 and

a sgRNA, and another harbouring the donor DNA to be integrated into the Cas9 cleavage site.

The Cas9 protein is co-expressed with a sgRNA from a single construct, as seen in Fig. 4A. Cas9

is expressed as a single transcript with EGFP, and the intervening T2A sequence facilitates

cleavage during protein translation, giving rise to individual Cas9 and EGFP proteins343,344. EGFP

fluorescence can therefore be used as a measure of transfection efficiency. The sgRNA is designed

to target Cas9 to induce a DSB within the first intron of the human PPP1R12C gene on

chromosome 19. This genomic region has been commonly referred to as the AAVS1 locus since

the observation that AAV integrates here345. AAV infection is not associated with any known

pathophysiology, and both hES and hiPS cells with disrupted PPP1R12C genes retain

pluripotency111,346. The AAVS1 locus is therefore considered a ‘genomic safe harbour’ for

transgene integration. Furthermore, this gene is ubiquitously expressed in all primary human cells

studied, as well as in commonly used transformed cell lines. For these reasons, the AAVS1 locus

is an ideal genomic locus to study precise gene targeting.

The donor DNA plasmid contains a promoterless mCherry cassette, which is flanked on both sides

by ~1 kb sequences that are homologous to the AAVS1 genomic DNA (Fig. 4B). These homology

arms facilitate site-specific integration of intervening plasmid sequence via HDR at the target

DSB site170. A splice acceptor site and T2A signal were inserted upstream of the mCherry ORF,

with a poly(A) signal downstream. Since AAVS1 exon 1 contains the translation start site,

insertion of this cassette into intron 1 gives rise to a single transcript driven by the endogenous

AAVS1 promoter. Because of the T2A signal upstream of mCherry, translation produces two

separate polypeptides: exon 1 of AAVS1, and mCherry. The absence of an upstream promoter on

the donor plasmid ensures that mCherry is only expressed from the endogenous AAVS1 promoter

- 42 -

following a successful integration event. This approach is analogous to gene trapping strategies347.

mCherry fluorescence is therefore a reliable measure of precise gene targeting, which can be

quantified by flow cytometry. This is demonstrated by the fact that transfection of either the donor

or CRISPR/Cas9 plasmid alone produces negligible mCherry expression (Fig. 4C). In contrast,

appreciable levels of mCherry expression are only attained following co-transfection of both

donor and CRISPR/Cas9 plasmids.

- 43 -

Figure 4. Schematic diagram of mCherry gene integration into the AAVS1 locus. a) CRISPR/Cas9 gene expression cassette containing the sgRNA and Cas9 flanked by a FLAG tag, NLS, T2A-EGFP, and a poly(A) signal. b) Promoterless mCherry donor construct containing the mCherry ORF downstream of a splice acceptor site and T2A, and upstream of a poly(A) signal. The entire cassette is flanked on either side by ~1kb of homologous sequence to the AAVS1 locus. Following a successful gene integration event, mCherry expression is driven by the endogenous AAVS1 promoter. c) HEK293 cells were transfected with the indicated plasmids and subsequently harvested 48 hr post-transfection for analysis by flow cytometry.

Exon1 Exon2AAVS1

Cas9

Exon1 Exon2AAVS1

a

b

Donor:CRISPR:

+-

-+

++

AAVS1

AAVS1

c

- 44 -

4.2 CtIP and EXO1-4D significantly enhance GTE

CtIP and EXO1 mutants were generated by site-directed mutagenesis (see Table 1 for a full list

of mutants). CtIP and EXO1 mutants, as well as CDK1, were then cloned into expression plasmids

under the control of the chicken E-actin promoter, which is frequently used to drive high levels of

gene expression in mammalian expression vectors. To assess the ability of CDK1, as well as CtIP

and EXO1 variants to enhance GTE, the respective expression plasmids were co-transfected with

the AAVS1 CRISPR/Cas9 and donor construct into HEK293 cells. 48 hr post-transfection, cells

were harvested for flow cytometry analysis. Representative flow cytometry data is shown in Fig.

5A. The depicted populations are gated on GFP+ cells, and the proportion of mCherry+ is

expressed as a percentage of the parent population. Because GFP is constitutively expressed from

the CRISPR/Cas9 plasmid, this effectively controls for transfection efficiency. To assess

enhancements in GTE, data were normalized to the CRISPR + donor condition. Co-transfection

with CtIP-WT, -3EA, -M, -M2A, -M3A as well as EXO1-4D and CDK1 elicited significant

enhancements in GTE up to ~2-3 fold (Fig. 5C). Furthermore, absolute targeting efficiencies

reached up to ~20% (Fig. 5B).

Subsequent studies investigated the possibility that observed enhancements in GTE were due in

part to a greater total amount of transfected DNA in experimental vs control conditions. For these

purposes, a lacZ expression plasmid (which theoretically would not influence GTE) was co-

transfected with either the donor plasmid alone, or together with the CRISPR + donor condition.

From these studies, it was evident that the total amount of DNA was indeed influencing the

observed GTE measurements (Fig. 5D). Furthermore, the increase in mCherry+ cells elicited by

co-transfection with lacZ (in the absence of CRISPR) indicated that there was a significant amount

of background mCherry expression from the donor plasmid. This is puzzling due to the absence

of an upstream promoter, but could be the result of weak cryptic promoter activity in the upstream

homology arm. Although the background expression may hinder the reliability of these data, these

experiments were taken as a sufficient initial screen to demonstrate that CtIP mutants were no

more effective than the wild-type CtIP protein. Accordingly, subsequent experiments were limited

to the investigation of CtIP-WT, CDK1 and EXO1-4D.

- 45 -

a

CtIP-3E

A

CtIP-M

3A

CtIP-M

CtIP-M

2A

CtIP-W

T

EXO1-4D

CDK10

1

2

3

4

%m

Che

rry+

/ %

GFP

+ (n

orm

aliz

ed)

**

*

CtIP-3E

A

CtIP-M

3A

CtIP-M

CtIP-M

2A

CtIP-W

T

EXO1-4D

CDK10

5

10

15

%m

Che

rry+

/ %

GFP

+

Donor: ++CRISPR:

++

Donor: +

+

lacZ lacZ

CRISPR:

Donor: +

+

CtIP-3EA CtIP-M3A CtIP-M

CtIP-M2A CtIP-WT CDK1 EXO1-4D

CRISPR:

b c

d

Figure 5. Enhancement of precise gene targeting efficiency by CDK1, CtIP and EXO1-4D in HEK293 cells. HEK293 cells were transfected with the indicated plasmids and subsequently harvested 48 hr post-transfection for analysis by flow cytometry. a) Representative flow cytometry experiment. The populations displayed are gated on GFP+ cells to control for transfection efficiency. b) Average gene targeting efficiency. c) Fold increase in gene targeting efficiency. All values are normalized to Donor+CRISPR condition. d) lacZ control reveals background mCherry expression from the Donor plasmid. Data presented as mean ± SEM (n=3). * P < 0.05, ** P < 0.01.

- 46 -

In order to dilute background mCherry expression from the donor plasmid, HEK293 cells were

passed 6 times over the course of 3 weeks post-transfection before flow cytometry analysis. This

effectively eliminated residual plasmid in cells and ensured that mCherry expression was only a

consequence of successful integration events. This is supported by the fact that in lacZ controls,

mCherry expression is an order of magnitude higher in the presence of CRISPR, despite equal

amounts of transfected DNA (Fig. 6A, panels 2,4). However, co-transfection of lacZ with

CRISPR + donor enhanced GTE values relative to CRISPR + donor controls (panels 3,4). This

can be explained due to the greater total amount of DNA transfected in the lacZ condition, which

effectively stabilizes CRISPR and donor plasmids, thus allowing greater opportunity for

integration events. Accordingly, GTE enhancements in the lacZ condition were subtracted from

GTE values in experimental samples in order to control for the amount of transfected DNA.

Corrected GTE values were then normalized to the CRISPR + donor condition. These experiments

demonstrated that CtIP-WT and EXO1-4D can elicit up to 3- and 6-fold enhancements in GTE,

respectively, whereas CDK1 had no significant effect (Fig. 6C). Absolute integration efficiencies

measured up to 0.5-1% of total cells (Fig 6B). Furthermore, various combinations of factors were

assessed for potential additive or synergistic effects on GTE enhancement, but revealed no

significant increases (Fig. 6E).

- 47 -

lacZ

lacZ

CtIP-WT+CDK1

EXO1+CDK1

CtIP-WT+EXO1-4D

0.0

0.2

0.4

0.6

0.8

1.0

%mCherry+

Donor: ++CRISPR:

lacZ

lacZ

CDK1

CtIP-W

T

EXO1-4D

0.0

0.5

1.0

1.5

%m

Che

rry+

Donor: ++CRISPR:

++

CDK1

CtIP-W

T

EXO1-4D

0

2

4

6

8

%m

Che

rry+

(nor

mal

ized

)

***

lacZ

lacZ

CDK1

CtIP-W

T

EXO1-4D

0.0

0.5

1.0

1.5

%m

Che

rry+

Donor: ++CRISPR:

++

CDK1

CtIP-W

T

EXO1-4D

0

2

4

6

8

%m

Che

rry+

(nor

mal

ized

)

***

CtIP-W

T + CDK1

CDK1 + EXO1-4

D

CtIP-W

T + EXO1-4

D0

2

4

6

%m

Che

rry+

(fol

d ch

ange

)

++

lacZ

lacZ

CDK1

CtIP-W

T

EXO1-4D

0.0

0.5

1.0

1.5

%m

Che

rry+

Donor: ++CRISPR:

++

CDK1

CtIP-W

T

EXO1-4D

0

2

4

6

8

%m

Che

rry+

(nor

mal

ized

)***nse

Figure 6. Enhancement of precise gene targeting by CtIP and EXO1-4D in HEK293 cells as measured by flow cytometry. HEK293 cells were transfected with the indicated plasmids and subsequently passed 6 times over the course of 3 weeks. Cells were then harvested for analysis by flow cytometry. a) Representative flow cytometry experiment. b) Average gene targeting efficiency of controls and individual factors. c) Fold change in gene targeting efficiency induced by individual factors. d) Average gene targeting efficiency of controls and combinations of factors. e) Fold change in gene targeting efficiency induced by combinations of factors. Data presented as mean ± SEM (n=3 or 4). * P < 0.05, ** P < 0.01.

lacZ CtIP EXO1-4D

lacZ

Donor: +

+CRISPR:

lacZ CDK1 CtIP-WT EXO1-4D

Panel #: 1 2 3 4 5 6 7

a

b c

lacZ

lacZ

CtIP-WT+CDK1

EXO1+CDK1

CtIP-WT+EXO1-4D

0.0

0.2

0.4

0.6

0.8

1.0

%mCherry+

Donor: ++CRISPR:

lacZ

lacZ

CtIP-WT+CDK1

EXO1+CDK1

CtIP-WT+EXO1-4D

0.0

0.2

0.4

0.6

0.8

1.0

%mCherry+

Donor: ++CRISPR:

d

- 48 -

Figure 7. Enhancement of precise gene targeting efficiency by SCR7 in HEK293 cells. a) HEK293 cells were transfected with the indicated plasmids. SCR7 was added to the culture media 18 hr post-transfection at the indicated concentrations. Cells were passed 6 times over the course of 3 weeks, and then harvested for analysis by flow cytometry. b) Comparing the respective fold changes associated with SCR7, CtIP-WT, and EXO1-4D as measured by flow cytometry 3 weeks post-transfection. Data presented as mean ± SEM (n=3). * P < 0.05.

DMSO0.0

1 0.1 10

1

2

3

4

%m

Che

rry+

(nor

mal

ized

) **

Donor: +

+

SCR7 (µM)

CRISPR:

0.1µM

SCR7

CtIP-W

T

EXO1-4D

0

2

4

6

8

rela

tive

enha

ncem

ent *

+

+

a b

SCR7 is a small molecule inhibitor of DNA ligase IV that has been widely utilized for

enhancing GTE. Investigators have been able to upregulate HDR and therefore GTE anywhere

from ~2 to 20-fold in mammalian cell lines and zygotes using SCR7315,316. Here, we

investigated the utility of SCR7 using our flow cytometry assay and compared these results to

those obtained by CtIP-WT and EXO1-4D. HEK293 cells treated with various concentrations

of SCR7 were similarly passed 6 times over the course of 3 weeks, and subsequently harvested

for flow cytometry analysis. These data demonstrate that 0.1µM SCR7 was the most effective

concentration used, eliciting up to 3-fold enhancements in GTE (Fig. 7A). Thus, the increase in

GTE evoked by CtIP-WT over-expression is comparable to that of SCR7, while EXO1-4D is

significantly more effective in this respect (Fig. 7B).

- 49 -

In order to confirm the GTE enhancements observed by flow cytometry, junction PCR was

performed at the AAVS1 integration site. Briefly, HEK293 cells were transfected with AAVS1

CRISPR/Cas9 and mCherry donor plasmids, as well as either CtIP-WT, EXO1-4D, or CDK1.

Transfectants were passed 6 times over the course of 3 weeks, and genomic DNA was

subsequently harvested and purified. Limited-cycle PCR was performed to amplify the junction

site using the indicated primers (Fig 8A). Band intensities for each condition were quantified and

normalized to their respective control amplicons (Fig. 8B). As in the flow cytometry experiments,

GTE enhancements in the lacZ condition were subtracted from GTE values in experimental

samples in order to control for the amount of transfected DNA. Corrected GTE values were then

normalized to the CRISPR + donor condition. These data largely corroborate those obtained from

flow cytometry experiments, demonstrating ~3- and 3.5-fold enhancements by CtIP-WT and

EXO1-4D, respectively. Furthermore, there was no significant enhancement by CDK1, and no

additive or synergistic effects by combining various factors (Fig. 8C).

- 50 -

a

c

Donor:

CRISPR:

+

+

CtIP-W

T

EXO1-4D

CDK1

CtIP-W

T + EXO1-4

D

CtIP-W

T + CDK1

CDK1+EXO1-4

D-1

0

1

2

3

4

5

band

inte

nsity

(nor

mal

ized

)

****

*

Donor: +

+CRISPR:

LeftArm RightArmpAmCherry2AGenomicDNA

----------------------------------------------1.45kb

--------------------------------------------------------------------------------------------------------------------------2.5kb

1:2:

b

1:

2:Prim

er s

et

Figure 8. Junction PCR demonstrating the enhancement of precise gene targeting efficiency by CtIP and EXO1-4D in HEK293 cells. a) Representation of genomic DNA containing a successfully integrated mCherry cassette. Primer set 1 is specific for the inserted cassette, while primer set 2 amplifies the wild-type AAVS1 locus. b) Representative junction PCR experiment. HEK293 cells were transfected with the indicated plasmids and subsequently passed 6 times over the course of 3 weeks. Genomic DNA was then harvested and purified from each treatment condition. Primer set 1 was used to amplify the mCherry insert, and primer set 2 was used as a PCR control. c) Fold change in gene targeting efficiency as measured by junction PCR. Data presented as mean ± SEM (n=3 or 4). * P < 0.05, ** P < 0.01.

Prim

er s

et

- 51 -

4.3 IB3-1-CFTR-EGFP cell line

One of the main limitations of the AAVS1 mCherry assay is that it cannot distinguish between

on-target and potential off-target integration events. Although the assay was designed so that

mCherry expression is driven by the endogenous AAVS1 promoter, it could also be inserted under

the control of an off-target promoter. We sought to generate a cell line that could be used for an

assay to distinguish between these distinct integration events. For these purposes, we generated a

cell line from IB3-1 cells, which are immortalized bronchial epithelial cells from a CF patient49.

To generate the cell line, CRISPR/Cas9 was programmed to cut immediately upstream of CFTR

exon 1 and insert an EGFP-puromycin cassette (Fig. 9A). Once the cell line had been obtained,

CRISPR/Cas9 was programmed to cut within the EGFP locus and insert an mCherry expression

cassette (Fig. 9B). A precise integration event would therefore knockout EGFP expression and

induce mCherry expression, whereas an off-target insertion would be double positive. These

distinct integration events could be readily measured by flow cytometry.

In order to generate the cell line, CRISPR/Cas9 targeting CFTR exon 1 and the requisite donor

DNA (Fig. 9A) were co-transfected into IB3-1 cells. Initially, we tried to isolate clonal integrants

by sorting EGFP+ cells using FACS. However, despite sorting close to 1000 single cells, not a

single well contained a growing colony after 3 weeks post-sorting. Puromycin selection was

thereafter performed on the remaining transfected plates in order to isolate clonal integrated cells.

Subsequent limited dilution allowed for single cell pipetting into 96-well plates. Single cell clones

were allowed proliferate (Fig. 10A), and were then expanded to larger culture vessels (Fig. 10B).

To this stage, 6 colonies were successfully expanded and healthy.

- 52 -

Exon1CFTRCas9

Exon2

Donor GFP pA puCMV PGK-puromycin-pA RightArmLeftArm

a

Figure 9. Schematic representation of the IB3-1-CFTR-EGFP cell line and the associated colour-switch assay. a) To generate the cell line, Cas9 was programmed to cut immediately upstream of CFTR exon1. The donor DNA to be integrated harbours an EGFP-puromycin expression cassette flanked on either side by ~500 bp of sequence homologous to the CFTR locus. b) Cas9 is programmed to cut the IB3-1-CFTR-EGFP cell line within the EGFP sequence, and subsequently insert an mCherry expression cassette.

Exon 1CFTR GFP pACMV

Cas9

GFP-mCherry+

GFP+mCherry-

GFP+mCherry+

Precise insertion:No integration: Random integration:

Left Arm pAmCherryCMV Right Arm

PGK-puromycin-pA

Donor T7

b

Exon 1CFTR GFP pACMV PGK-puromycin-pA

Exon 1CFTR GFP pACMV

Cas9

GFP-mCherry+

GFP+mCherry-

GFP+mCherry+

Precise insertion:No integration: Random integration:

Left Arm pAmCherryCMV Right Arm

PGK-puromycin-pA

Donor T7

Exon 1CFTR GFP pACMV

Cas9

GFP-mCherry+

GFP+mCherry-

GFP+mCherry+

Precise insertion:No integration: Random integration:

Left Arm pAmCherryCMV Right Arm

PGK-puromycin-pA

Donor T7

- 53 -

Figure 10. Investigating the identity and CRISPR cleavage efficiency of IB3-1-CFTR-EGFP cell line clones. a) Fluorescence microscopy image of clone #2 isolated from puromycin selection (20X magnification) and b) following 10 passages grown to confluency (5X magnification). c) PCR amplification of the wild-type CFTR locus and d) the transgenic EGFP sequence from purified genomic DNA of each clone. e) T7E1 measuring CRISPR cleavage efficiency in each clone. Each clone was transfected with CRISPR (targeting EGFP) and single-stranded mCherry donor DNA (Fig. 11), and genomic DNA was harvested and purified one week post-transfection. PCR was used to amplify EGFP from each clone, the products of which were denatured and randomly re-annealed. Re-annealed amplicons were then incubated with T7E1 endonuclease to cleave amplicons with base-pair mismatches, and then run on a 2% agarose gel.

a b

4% 6% 17% 19% 15% 13% 26% 34%Clone #: 2 3 1 2 3 4 5 6

negative controls

positivecontrol

Cleavageefficiency:

e

Clone #: 1 2 3 4 5 6 1 2 3 4 5 6

dc

- 54 -

In order to verify that the EGFP cassette had been inserted in the correct locus, we tried to amplify

the EGFP-CFTR junction from the 6 clones obtained from selection. However, despite being able

to amplify the wild-type CFTR locus (Fig. 10C) and the EGFP sequence (Fig. 10D), we were

unable to amplify the EGFP-CFTR junction. A myriad of primer sets and PCR conditions were

attempted with no success for any of the 6 clones obtained, indicating off-target integrations.

Because all 6 clones appeared to have off-target integrations, we speculated that the homology

arms flanking the EGFP-puromycin cassette on the donor DNA may not have perfect sequence

homology to the IB3-1 CFTR locus, as this is a transformed cell line. To investigate this

possibility, we PCR amplified the CFTR locus from genomic DNA isolated from IB3-1 cells, and

Sanger sequencing was used to analyze the PCR products. However, both homology arms used

on the donor DNA in fact harboured 100% identity to the IB3-1 CFTR genomic DNA (Fig. 11).

However, off-target insertions should not necessarily prohibit the use of these cell lines for the

colour switch assay, as the requisite homology arms on the mCherry donor do not span beyond

the EGFP cassette (Fig. 9B). Furthermore, numerous studies have demonstrated the utility of

linear single-stranded donor DNA (as opposed to circular double-stranded plasmids), for

CRISPR/Cas9-mediated gene targeting333–338. Therefore, we also wanted to assess the utility of

CtIP and EXO1-4D for enhancing GTE using ssDNA donors. In order to generate ssDNA from

the mCherry donor, a T7 promoter was cloned in the reverse orientation downstream of the

homology arms (Fig 9B). T7 in vitro transcription followed by DNaseI digestion and RNA

purification generates large quantities of donor transcripts. These transcripts are subsequently

used for gene-specific reverse transcription, followed by RNaseH digestion and ssDNA

purification. Products of each step were run on an agarose gel and can be seen in Fig. 12. In order

to verify the identity of the ssDNA, it was used as a template for PCR, purified, and sequenced.

- 55 -

GAGGAGGAAGGCAGGCTCCGGGGAAGCTGGTGGCAGCGGGTCCTGGGTCTGGCGGACCCTGACGCGAAGGAGGGTCTAGGAAGCTCTCCGGGGAGCCGGTTCTCCCGCCGGTGGCTTCTTCTGTCCTCCAGCGTTGCCAACTGGACCTAAAGAGAGGCCGCGACTGTCGCCCACCTGCGGGATGGGCCTGGTGCTGGGCGGTAAGGACACGGACCTGGAAGGAGCGCGCGCGAGGGAGGGAGGCTGGGAGTCAGAATCGGGAAAGGGAGGTGCGGGGCGGCGAGGGAGCGAAGGAGGAGAGGAGGAAGGAGCGGGAGGGGTGCTGGCGGGGGTGCGTAGTGGGTGGAGAAAGCCGCTAGAGCAAATTTGGGGCCGGACCAGGCAGCACTCGGCTTTTAACCTGGGCAGTGAAGGCGGGGGAAAGAGCAAAAGGAAGGGGTGGTGTGCGGAGTAGGGGTGGGTGGGGGGAATTGGAAGCAAATGACATCACAGCAGGTCAGAGAAAAAGGGTTGAGCGGCAGGCACCCAGAGTAGTAGGTCTTTGGCATTAGGAGCTTGAGCCCAGACGGCCCTAGCAGGGACCCCAGCGCCCGAGAGACC

ATCATTCATTGTTTTGAAAGAAAATGTGGGTATTGTAGAATAAAACAGAAAGCATTAAGAAGAGATGGAAGAATGAACTGAAGCTGATTGAATAGAGAGCCACATCTACTTGCAACTGAAAAGTTAGAATCTCAAGACTCAAGTACGCTACTATGCACTTGTTTTATTTCATTTTTCTAAGAAACTAAAAATACTTGTTAATAAGTACCTAAGTATGGTTTATTGGTTTTCCCCCTTCATGCCTTGGACACTTGATTGTCTTCTTGGCACATACAGGTGCCATGCCTGCATATAGTAAGTGCTCAGAAAACATTTCTTGACTGAATTCAGCCAACAAAAATTTTGGGGTAGGTAGAAAATATATGCTTAAAGTATTTATTGTTATGAGACTGGATATATCTAGTATTTGTCACAGGTAAATGATTCTTCAAAAATTGAAAGCAAATTTGTTGAAATATTTATTTTGAAAAAAGTTACTTCACAAGCTATAAATTTTAAAAGCCATAGGAATAGATACCGAAGTTATATCCAACTGACATTTAATAAATTGTATTCATAGCCTAATGTGATGAGCCACAGAAGCTTGCAAACTTTAATGAG

Figure 11. IB3-1 CFTR homology arm sequencing. Genomic DNA from IB3-1 cells was harvested as described in methods. PCR was used to amplify the region corresponding to the homology arms on the IB3-1 CFTR donor plasmid. PCR products were purified and analyzed by Sanger sequencing.

Exon1CFTRCas9

Left HA Right HA

Exon2

- 56 -

Lane: 1 2 3 4 5 6 7

CRISPR/Cas9 (targeting EGFP) and the ssDNA mCherry donor were co-transfected into each

of the 6 IB3-1-CFTR-EGFP clones. 3 different sgRNAs targeting different EGFP sequence

motifs were tested for each clone. One week post transfection, genomic DNA from each

condition was harvested and purified. The T7E1 genomic cleavage assay was performed to

assess CRISPR/Cas9 cutting efficiency. This assay relies on the fact that Cas9-induced DSBs

are frequently repaired by NHEJ, resulting in indels at the break site. The sequence

surrounding the break site is then amplified from genomic DNA by PCR, the products of

which are then denatured and randomly re-annealed. This results in mutated sequence (a

consequence of Cas9 cleavage) pairing with wild-type sequence. The randomly annealed

amplicons are then incubated with T7E1 endonuclease, which cleaves mismatched dsDNA.

The cleaved amplicons can then be visualized as two distinct bands on an agarose gel, and are

a measure of CRISPR/Cas9 cutting efficiency (Fig. 10E). Only one of the three sgRNAs tested

resulted in appreciable cutting efficiencies. Despite successful cutting, however, when the

transfected cells were visualized by fluorescence microscopy prior to harvesting genomic

DNA, no mCherry fluorescence was evident.

Figure 12. Single-stranded mCherry donor production. Lane 1 is an RNA ladder. T7 in vitro transcription was performed using the mCherry donor plasmid (Fig. 6B), followed by DNaseI digestion and RNA purification (product run in lane 2). RNA was then used as a template for primer-specific reverse transcription, followed by RNaseH digestion (product run in lane 3), and ssDNA purification (product run in lane 4). Lane 5 is a DNA ladder. Lanes 6 and 7 are the same as lanes 3 and 4, respectively, except they were produced using a different primer for reverse transcription.

- 57 -

Chapter 5 Discussion

The technological power of CRISPR/Cas9 gene editing has already made a substantial impact on

genetics and genomics; agriculture and livestock; stem cell research and tissue engineering; the

development of organoids and disease models; as well as a range of gene and cell therapies. Of

particular interest is the applicability of CRISPR/Cas9 gene editing for in vivo gene therapies.

More specifically, the power to generate permanent genomic changes in diseased tissues without

the use of integrating viral vectors. This represents a tremendous technological leap for our ability

to correct heritable diseases at their genetic origins. Our group has sought to apply these

technologies specifically for CF gene therapy. Using our HD-Ad vectors, our goal is to deliver

CRISPR/Cas9 and the wild-type CFTR gene to permanently correct the CFTR genetic defect in

airway stem/progenitor cells, for which we have recently demonstrated successful gene delivery

in vivo85. However, the utility of this approach is still limited by the inherently low efficiency of

CRISPR/Cas9 gene targeting by HDR. In these studies, we have discovered genetically-encoded

means by which to significantly enhance GTE, which may improve the clinical utility of in vivo

gene therapies.

Our approach to enhancing GTE was largely based on previous studies demonstrating the

feasibility of perturbing DSB repair in favour of HDR. Accordingly, we sought to overexpress

CDK1, which has extensive regulatory control over the balance between DSB repair pathways,

whose activity promotes HDR while inhibiting NHEJ. There was also strong theoretical and

empirical support to investigate more proximal regulators of DNA end resection, namely CtIP

and EXO1, and site-specific mutants thereof. Our AAVS1-mCherry integration assay initially

appeared to be a reliable measure by which to test these factors (Fig. 4C). Initial screens using

this assay suggested that CDK1, as well as the CtIP and EXO1 variants could enhance GTE by

~2 to 3-fold (Fig. 5C). However, further investigation revealed that there was significant

background mCherry expression from the donor plasmid itself (Fig. 5D). This is puzzling, as there

is no upstream promoter on the mCherry donor cassette (Fig. 4B). The background expression is

therefore likely a result of weak cryptic promoter activity in the upstream homology arm.

- 58 -

However, there is no documented evidence for promoter activity in intron 1 of the PPP1R12C

gene in the literature.

Due to the background expression from the donor plasmid, we reasoned that harvesting

transfectants 48 hr post-transfection for flow cytometry was not the most reliable measure of

targeting efficiencies. However, these experiments were taken as a sufficient initial screen to

demonstrate that CtIP mutants were no more effective than the wild-type protein for enhancing

GTE (Fig. 5C). Transfectants of subsequent experiments were passaged over the course of 3

weeks prior to flow cytometry. This effectively diluted the background mCherry expression, such

that mCherry expression was only a consequence of successful integration events (Fig. 6). These

experiments revealed that CtIP-WT and EXO1-4D can significantly enhance GTE by ~3 and 6-

fold, respectively. CDK1 surprisingly had a negative impact on GTE with respect to controls, but

this reduction was not statistically significant.

There are a number of potential reasons why CDK1 did not upregulate GTE in these studies.

Firstly, the catalytic activity of CDK1 is dependent on binding to its cognate cyclin proteins,

which are themselves transcriptionally restricted to S/G2348,349. Therefore, ectopic CDK1

expression would presumably only upregulate HDR in the cell cycle phases in which HDR is

already most active. Such upregulation is also conditional on the relative concentrations of CDK1

to cyclins. That is, if cyclin proteins are endogenously limiting, ectopically overexpressed CDKs

would in fact not potentiate CDK activity, and therefore have no influence on GTE. Furthermore,

although the effect was not statistically significant, there was a trend towards reduced GTE in

response to CDK1 overexpression (Fig. 6). This can potentially be explained by the fact that

CDK1 has, in some cases, been shown to down-regulate HDR. For example, CDK1 activity has

been shown to inhibit RAD51 nucleoprotein filament formation in M phase Xenopus extracts350,

and forced activation of CDK1 in interphase cells can attenuate HDR in human cell lines351. CDKs

also control a multitude of metabolic processes, epigenetic modifications and transcriptional

programs352. Therefore, their overexpression may be cytotoxic, in turn reducing GTE in

transfected cells.

Based on our initial screen using the AAVS1-mCherry assay, the CtIP mutants tested were no

more effective in potentiating GTE than the wild type protein (Fig. 5). Accordingly, they were

- 59 -

not included in subsequent experiments. Retrospectively, however, the reliability of these

experiments appears to have been severely hampered by background expression from the donor

plasmid. For example, the direction of the CDK1 effect was reversed following donor plasmid

dilution (Fig. 5C vs Fig. 6C). Flow cytometry following vector dilution was likely a more accurate

measure of GTE, and this effect was confirmed by semi-quantitative junction PCR (Fig. 8). In

hindsight, therefore, one cannot utilize the initial screens as a reliable measure of GTE, nor

potential enhancements thereof.

With that in mind, it may not be reasonable to speculate on why the CtIP mutants were not more

effective than the wild-type protein, as it is evident that they were not reliably tested. Future

studies should therefore subject these mutants to the experimental protocol used in Fig 6.

However, it is worthwhile to retrospectively speculate on the shortcomings of the CtIP mutants

generated in these studies. Firstly, the mutants tested concurrently harboured a number of

mutations. This may destabilize the native structure of the protein and therefore preclude its

potential utility for perturbing DSB repair. Such a phenomenon could be verified by RT-qPCR

and western blot. For these reasons, a more suitable approach would be to create a small library

of CtIP mutants, each possessing individual mutations and various combinations thereof.

However, assuming these combinatorial mutants did not influence native protein folding, there

are still limitations to this approach due to the quaternary structure of CtIP. That is, CtIP functions

as a homotetramer353, and therefore ectopically-expressed mutant peptides may readily form

quaternary structures with wild type conspecifics. This would presumably subject these proteins

to wild type regulatory stimuli, thus reducing potential efficacies of these mutants. A viable

solution to this limitation would be to co-express siRNAs targeting endogenous CtIP with siRNA-

resistant CtIP mutants.

It is evident that the results obtained here for the potential GTE-enhancing effects of the CtIP

mutants studied were inconclusive. Perhaps surprisingly, however, CtIP-WT significantly

enhanced GTE up to ~3-fold, as evidenced by flow cytometry (Fig. 6) and junction PCR (Fig. 8).

This may be a consequence of overcoming the endogenous down-regulation of CtIP-dependent

end resection natively mediated by ubiquitylation311–313. That is, CtIP overexpression may saturate

or exceed the capacities of the cognate E3 ubiquitin ligases to ubiquitylate CtIP, thus overcoming

the associated suppression of end resection.

- 60 -

The substantial increase in GTE in response to EXO1-4D overexpression is unsurprising, as it has

been shown to possess hyper-resective activity and potentiate HDR321. However, it is evident that

the magnitude of the ~6-fold increase observed using flow cytometry was not entirely

recapitulated by the ~3.5-fold increase observed using junction PCR (Fig. 6C vs Fig. 8C). This

discrepancy may be a consequence of off-target insertions. The flow cytometry assay measures

loci-independent mCherry expression within cells, whereas a positive junction PCR product is

dependent on a precise insertion at the AAVS1 locus. The 6-fold enhancement measured by flow

cytometry in response to EXO1-4D may therefore reflect an up-regulation of both on- and off-

target insertions. If this is indeed the case, the observed enhancements for CtIP would not have

been a consequence of off-target insertions, as the fold increases measured by flow cytometry and

junction PCR were virtually identical at ~3-fold (Fig. 3C, Fig. 5C). The discrepancy between

junction PCR and flow cytometry data for EXO1-4D could also be explained by the semi-

quantitative nature of junction PCR. Due to the exponential amplification of DNA in PCR,

saturation of amplicons may reduce the dynamic range of absolute band intensities, and thus the

fold enhancement in GTE.

We also investigated potential synergistic or additive effects associated with co-expression of

CtIP-WT and EXO1-4D (Fig. 6D,E). However, although these combinatorial approaches

increased GTE, the effects were not statistically significant, and thus no more effective than the

individual factors. Because CtIP and EXO1 function in contiguous biochemical pathways in

HDR, their co-overexpression may lead to dramatic hyper-resection phenotypes and severe

genomic instability. Concurrently overexpressing these factors may therefore be synthetically

lethal and undesirable for our purposes.

The IB3-1-CFTR-EGFP cell line would have been a useful model to test the efficacies of CtIP

and EXO1-4D, as it would have been able to distinguish between on- and off-target integration

events. Perhaps ironically, it was evident that all 6 clones obtained for the cell line harboured

EGFP-puromycin cassettes knocked in at off-target genomic loci. This could not be explained by

lack of sequence identity between the donor homology arms and CFTR genomic DNA, as the

genomic DNA was sequenced and revealed 100% identity. The results of this cell line generation

underscore another significant challenge facing CRISPR technology, that is, specificity. Although

not the focus of these studies, the ability to eliminate (or at least minimize) off-target genomic

- 61 -

cleavage events will be especially important for CRISPR therapeutics. This exemplifies the utility

of highly specific Cas9 variants such as eSpCas9 and SpCas9-HF197,198, which should be used in

these studies going forward.

Despite the fact that IB3-1 cell line clones harboured off-target insertions, we reasoned that this

would not abolish the utility of the clones. The requisite homology arms on the mCherry donor

do not span beyond the EGFP cassette (Fig. 9B), and therefore these clones could still theoretically

be used to assess precise insertions. Accordingly, we sought to test the utility of ssDNA donors

using this assay, as they have become the preferred donor template for many CRISPR gene

targeting applications333–338. However, upon co-transfection of the mCherry ssDNA donor and

CRISPR/Cas9 into each of these clones, no mCherry expression was evident by fluorescence

microscopy. This may have been a result of the integrity of the ssDNA production. As seen in

Fig. 12, the purified ssDNA migrates as a smear and faster than the corresponding RNA of similar

size. It remains unclear if this is simply the nature of how long (>2kb) ssDNA migrates on a non-

denaturing agarose gel, or if the product is a heterogeneous mixture of various species. Future

experiments should perform linear PCR (using a single primer) on the plasmid mCherry donor

template to generate ssDNA, in order to observe its migratory behavior on an agarose gel. This

will help evaluate the identity of the ssDNA. Furthermore, plasmid donor DNA should be co-

transfected with the CRISPR/Cas9 plasmid into the IB3-1 cell line clones and monitored for

mCherry expression. These experiments will help explain the inability to elicit mCherry

expression in the IB3-1 cell line clones using ssDNA donors in these studies.

5.1 Future directions

Further studies will undoubtedly need to confirm the efficacy of these approaches in primary

human cells to evaluate their applicability for clinical therapies. For such purposes, these factors

will need to be cloned into our HD-Ad vectors together with CRISPR/Cas9 and donor DNA.

Changing the context of the donor DNA from plasmids to HD-Ad vectors may influence the utility

of these factors, as it has been shown that the type of DSB repair pathway employed can be

- 62 -

dependent on the nature of the repair template354–356. Indeed, there are evident differences in the

behavior of viral DNA and small circular plasmids within the nucleus357,358.

If the efficacies of these factors are translatable to viral vectors in the context of primary cells and

animal models, this will have a tremendous impact on clinical applications for CF gene therapy.

Preliminary data from our laboratory suggests that we can achieve permanent transgene

integration in ~10% of transformed airway cells using our CRISPR HD-Ad vectors, without CtIP

or EXO1-4D (unpublished data). If the addition of CtIP or EXO1-4D to our CRISPR HD-Ad

vectors elicits GTE enhancements consistent with the data obtained here, we may be able to

achieve therapeutically-significant levels of gene editing in airway stem/progenitor cells.

Additional barriers to this goal include in vivo gene delivery and the host immune response. To

these ends, we have recently shown that our vectors can effectively target airway stem/progenitor

cells in primary culture, as well as in mouse and pig lungs in vivo85. Furthermore, studies have

shown that ~10% of wild type CFTR expression in CF airways is sufficient to correct lung

pathology359,360. Therefore, if the GTE enhancing strategies discovered here can be applied to our

HD-Ad vectors, in parallel with improvements in subverting the host immune response, we may

be able to achieve permanent correction of CF lung pathology.

Additional genetically-encoded approaches to enhancing CRISPR/Cas9 GTE have recently been

documented. For example, Canny et al. has generated a small ubiquitin peptide variant, i53, that

inhibits 53BP1 accumulation at DSB sites, in turn down-regulating NHEJ and promoting

compensatory HDR361. The authors demonstrate that i53 can be used to enhance CRISPR/Cas9

GTE by up to 5.6-fold in mouse and human cell lines using ds- and ssDNA donors. Furthermore,

the upregulation of HDR using i53 was largely dependent on the presence of CtIP. This suggests

that potential synergistic effects on GTE could be elicited by co-expressing i53 and CtIP (or

mutant CtIP variants). This may indeed be true for EXO1 as well, although not tested in this study.

Due to the large cloning capacity of HD-Ad vectors (~36 kb), i53 could be readily packaged into

our vectors together with either CtIP or EXO1-4D, in order to further enhance CRISPR/Cas9

GTE.

Despite the advancements made here and elsewhere for enhancing CRISPR/Cas9 GTE, there are

still unanswered questions and further room for improvement. It is commonly understood in the

- 63 -

research community that the favourability of NHEJ over HDR represents the sole and principle

impediment to CRISPR/Cas9 GTE. However, there are likely additional cellular and molecular

barriers to GTE that have been unexplored and largely overlooked, yet may be equally relevant.

These include the bias for sister chromatids as templates for repair in HDR, and the necessity of

exogenous donor DNA to spatially co-localize with the integration site for successful gene

targeting.

When HDR is employed to repair DSBs in post-replicative cells, the sister chromatid template is

physically linked to the DSB by the cohesin complex and associated factors362. Cohesin maintains

spatial co-localization of sister chromatids, and is crucial for efficient HDR in yeast and

mammalian cells363–365. Accordingly, cohesin is readily recruited to DSBs via the DNA damage

response366,367, and genome-wide sister chromatid cohesion is up-regulated in response to DSBs

in S/G2368. Furthermore, the ability of a particular sequence to function as a template for

recombinational repair is primarily limited by its proximity to the site of DNA damage369.

Unsurprisingly, the frequency at which sister chromatids are used as templates for HDR is orders

of magnitude greater than that of ectopic sequences370–372. Efficient HDR is therefore dependent

on the spatiotemporal co-localization of the template sequence with the DSB. It logically follows

that during the stage of the cell cycle in which HDR is most efficient (S/G2), the sister chromatid

physically associated with the DSB will predominate as the template for repair over exogenous

DNA that lacks a spatial bias for the genomic integration site. This is supported by the fact that

down-regulating essential cohesin subunits dramatically increases aberrant gene conversion

between homologous chromosomes373. The biased use of sister chromatids as templates for HDR

may therefore be a significant barrier suppressing CRISPR/Cas9 GTE.

The challenge of the sister chromatid bias is further compounded by the fact that exogenous DNA

does not have the liberty of unrestricted nuclear diffusion. In fact, some reports have shown that

plasmid DNA is >100-fold less mobile than soluble proteins of similar dimensions357, and that

DNA ranging from 21 bp to 6 kb is completely immobile374. Similarly, others have reported that

plasmids >5 kb accumulate in stagnant clusters at the periphery of the nucleus at sites of nuclear

entry375. On the contrary, some reports suggest that DNA <400 bp rapidly diffuses throughout the

nucleus and cytoplasm376, whereas others have demonstrated that plasmid mobility and

localization depends on its transcriptional output377,378. It appears that the spatiotemporal

- 64 -

dynamics of exogenous DNA within the nucleus remains relatively unclear. However, one could

reasonably conclude that an exogenous donor template would not have the liberty of unrestricted

nuclear diffusion, thus further amplifying the limitation of lacking a spatial bias for the genomic

integration site.

The impetus of conventional wisdom in this field has restricted investigative efforts on

potentiating compensatory HDR by inhibiting NHEJ. This is straightforwardly obvious, as gene

targeting depends on HDR. This approach also subverts the sister chromatid bias by artificially

upregulating HDR in G1 – i.e., in the absence of a sister chromatid. However, the lack of an active

spatial bias of the donor template for the target site raises additional concerns in this context. That

is, if HDR is employed at targeted DSBs in G1 in the absence of a spatiotemporally associated

donor template, ectopic recombination between non-allelic genomic sequences is a foreseeable

consequence. Such recombination events can lead to loss of heterozygosity (LOH), gross

chromosomal rearrangements (GCRs) and tumorogenesis379,380. These potential consequences

deserve more attention from the research community investigating ways to enhance CRISPR-

mediated gene targeting.

A number of potential strategies independent of DSB repair pathway choice could be employed

to circumvent the prospective deleterious consequences mentioned above. One idea would be to

minimize the use of sister chromatids as templates for repair in HDR in S/G2, thus raising the

probability that exogenous sequences would be used as templates for repair. This could be

accomplished by targeting critical regulators of cohesin dynamics, such as sororin and WAPL, to

reduce the physical association of sister chromatids in S/G2381,382. However, this approach

introduces the possibility of adverse cytogenetic effects that are observed in cohesinopathies383.

A potential solution to this would be to localize perturbations in cohesin dynamics to the target

integration site. This could be accomplished by using dCas9 as a fusion platform to direct cohesin

regulators to target sites, analogous to the use of dCas9 for transcriptional and epigenetic

regulation. This approach would additionally require the use of an orthogonal programmable

nuclease to induce the targeted DSB. The principle drawback here would therefore be the

necessity for excessive molecular machinery and attendant cloning capacities.

- 65 -

There may be a more parsimonious and effective way to overcome a number of these challenges

simultaneously. That is, active spatiotemporal recruitment of donor DNA molecules to the target

integration site. The approach would involve covalently linking ssDNA donors to the Cas9

enzyme inducing the targeted DSB. This could be effectively accomplished by fusing Cas9 to

SNAP-tag, a synthetic mutant of the DNA repair protein O6-alkylguanine-DNA alkyltransferase

(AGT) that reacts specifically with benzylguanine (BG) derivatives to generate covalent

linkages384. BG derivatives can themselves be covalently linked to ssDNA donors if the ssDNA

is generated using terminal amine-modified oligos. The covalent linkage between Cas9-SNAP

and BG-ssDNA can then be generated in vivo following co-transfection of Cas9-SNAP and BG-

ssDNA, or by in vitro methods (see online New England Biolabs, SNAP-tag Technologies for

details). Interestingly, studies have shown that Cas9 remains strongly associated with DSB

cleavage products following nucleolytic catalysis183. Cas9-SNAP-BG-ssDNA would therefore

establish and maintain donor co-localization following DSB induction, thus increasing the

probability that the localized donor will be used as a template for subsequent repair. Cas9-SNAP-

BG-ssDNA would be a large nucleoprotein complex, theoretically capable of making targeted

genomic DSBs while delivering the template donor DNA to the target locus. Using this engineered

approached, therefore, not only would Cas9 function as a programmable nuclease, but additionally

as a molecular motor that guides transgene cassettes to their cognate target loci.

This approach would simultaneously overcome a number of limitations currently facing efficient

CRISPR-mediated gene targeting. Namely, this would impose a spatiotemporal bias on the donor

DNA for its cognate target integration site. Accordingly, this would increase the probability that

exogenously supplied donor DNA is used as a template for Cas9 DSB repair over the sister

chromatid in S/G2. This could also be used in parallel with strategies that enhance the use of HDR

in G1. In this context, spatiotemporal co-localization of donor DNA would mitigate the possibility

of deleterious recombination events at the target site in the absence of a sister chromatid. This

approach would also presumably reduce the frequency of off-target insertions, as the donor DNA

would be spatially restricted to its target site, and therefore less available for recombination at

distant off-target loci.

The main limitation of this approach would be the requirement for chemical modifications on the

donor DNA which cannot be genetically encoded. Therefore, these could not be collectively

- 66 -

packaged into viral vectors. However, Cas9-SNAP-BG-ssDNA nucleoprotein complexes could

be readily synthesized in vitro and packaged into LNPs for ex vivo and in vivo delivery. These

components could also be packaged separately and assembled in vivo. That is, Cas9-SNAP could

be encoded and delivered in viral vectors in parallel with LNP-delivered BG-ssDNA. LNPs are

able to effectively deliver proteins and nucleic acids to the brain385, retina45,46, inner ear47, liver48,

as well as the nasal epithelium56,386 and lung54,387. This approach could therefore be amenable to

a variety of gene therapy applications, including lung gene therapy for CF.

5.2 Conclusion

Gene therapies have attracted substantial attention from physicians, scientists and the public alike,

and have found broad ranging therapeutic applications. However, many gene therapy clinical

trials have failed in the last few decades due to insufficient understanding of the major challenges

and a lack of the requisite technologies. Classical approaches involving -RTs were initially

effective but elicited deleterious genotoxic effects due to random genomic insertions. On the

contrary, superior integrating vectors like SIN--RVs and lentiviruses with improved safety

profiles have demonstrated utility in a number of clinical trials. However, they still rely on semi-

random integration, and therefore carry the risk of insertional mutagenesis and oncogenic

transformation. Semi-random integration also gives rise to transgene expression heterogeneity

amongst targeted cells, thus generating cells with differential potency for cell therapies. Although

non-integrating vectors such as AdVs and AAVs subvert these risks and therefore possess better

safety profiles, their efficacies for gene therapy are limited by the half-life and finitude of their

cognate target cells.

The advent of programmable nucleases such as CRISPR/Cas9 will greatly attenuate these risks

and overcome impeding limitations. The targeting capacities of CRISPR/Cas9 to permanently

correct gene code will improve the efficacies of non-integrating vectors, while mitigating the

genotoxic risks of insertional mutagenesis associated with integrating viruses. Directed genome

edits also have the advantage of preserving endogenous regulatory elements and therefore the

native spatiotemporal patterns of gene expression. Furthermore, Cas9 has proven to be a robust

- 67 -

platform for rationally-engineered mutants with novel genomic targeting functions. As a

consequence, this technology has opened the door to inducing genetic manipulations that were

hitherto unfeasible, such as making single base pair edits and targeted epigenetic modulations.

Despite the power of CRISPR/Cas9, immanent challenges remain for the application of these

technologies into the clinic. In particular, hijacking HDR of targeted DSBs to insert therapeutic

transgenes has thus far remained inefficient with limited solutions. Here, we present the use of

CtIP and EXO1-4D as novel, genetically-encoded strategies to perturb DSB repair in favour of

HDR and consequently enhance GTE. These studies have demonstrated that CtIP and EXO1-4D

overexpression can upregulate CRISPR/Cas9 GTE up to ~3- and 6-fold, respectively, in human

cell lines. These discoveries represent a significant step forward for our CF gene therapy strategy,

but future studies will be necessary to address a host of lingering unanswered questions. In

particular, assessing the utility of these strategies in the context of viral vectors in primary cells

and in vivo animal models will be pertinent to clinical translations.

Our efforts to generate the IB3-1-CFTR-EGFP cell line also illuminate the potential for insidious

off-target edits, which remain a preeminent concern for CRISPR therapeutics. Furthermore, the

long-term consequences of transiently manipulating DSB repair have yet to be thoroughly

investigated. The appropriate clinical implementations of CRISPR/Cas9 technologies will be

incumbent on our fastidious attention to these salient risk factors. Notwithstanding these persistent

challenges, however, CtIP and EXO1-4D may be clinically significant for viral gene therapy for

genetic diseases such as CF. Our high capacity HD-Ad vectors provide a unique opportunity to

co-package these factors with CRISPR/Cas9 and donor DNA on a single vector, which can readily

target airway stem/progenitor cells. The realization of this approach with the requisite

technologies may ultimately provide a cure for CF, the most common genetic disease in the

Caucasian population.

- 68 -

References

1. Bobadilla, J. L., Macek, M., Fine, J. P. & Farrell, P. M. Cystic fibrosis: A worldwide

analysis of CFTR mutations - Correlation with incidence data and application to

screening. Hum. Mutat. 19, 575–606 (2002).

2. O’Sullivan, B. P. & Freedman, S. D. Cystic fibrosis. The Lancet 373, 1891–1904 (2009).

3. MacKenzie, T., Gifford, A. H., Sabadosa, K. A., Quinton, H. B., Knapp, E. A., Goss, C.

H. & Marshall, B. C. Longevity of patients with cystic fibrosis in 2000 to 2010 and

beyond: Survival analysis of the Cystic Fibrosis Foundation Patient Registry. Ann. Intern.

Med. 161, 233–241 (2014).

4. Marshall, B., Elbert, A., Petren, K., Rizvi, S., Fink, A., Ostrenga, J., Sewall, A., Loeffler,

D. Patient Registry: Annual Data Report 2015. Cyst. Fibros. Found. Patient Regist. 94

(2016).

5. Elborn, J. S. Cystic fibrosis. The Lancet 388, 2519–2531 (2016).

6. Locher, K. P. Mechanistic diversity in ATP-binding cassette (ABC) transporters. Nat.

Struct. Mol. Biol. 23, 487–493 (2016).

7. Hunt, J. F., Wang, C. & Ford, R. C. Cystic fibrosis transmembrane conductance regulator

(ABCC7) structure. Cold Spring Harb. Perspect. Med. 3, a009514 (2013).

8. Hwang, T. C. & Kirk, K. L. The CFTR Ion channel: Gating, regulation, and anion

permeation. Cold Spring Harb. Perspect. Med. 3, a009498 (2013).

9. Chong, P. A., Kota, P., Dokholyan, N. V. & Forman-Kay, J. D. Dynamics intrinsic to

cystic fibrosis transmembrane conductance regulator function and stability. Cold Spring

Harb. Perspect. Med. 3, a009522 (2013).

10. Dalemans, W., Barbry, P., Champigny, G., Jallat, S., Jallat, S., Dott, K., Dreyer, D.,

Crystal, R. G., Pavirani, A., Lecocq, J.P. & Lazdunski, M. Altered chloride ion channel

kinetics associated with the ΔF508 cystic fibrosis mutation. Nature 354, 526–528 (1991).

- 69 -

11. Li, C., Ramjeesingh, M., Reyes, E., Jensen, T., Chang, X., Rommens, J. M. & Bear, C. E.

The cystic fibrosis mutation (ΔF508) does not influence the chloride channel activity of

CFTR. Nat. Genet. 3, 311–316 (1993).

12. Lukacs, G. L., Chang, X. B., Bear, C., Kartner, N., Mohamed, A., Riordan, J. R. &

Grinstein, S. The ΔF508 mutation decreases the stability of cystic fibrosis transmembrane

conductance regulator in the plasma membrane: Determination of functional half-lives on

transfected cells. J. Biol. Chem. 268, 21592–21598 (1993).

13. Protasevich, I., Yang, Z., Wang, C., Atwell, S., Zhao, X., Emtage, S., Wetmore, D., Hunt,

J. F. & Brouillette, C. G. Thermal unfolding studies show the disease causing F508del

mutation in CFTR thermodynamically destabilizes nucleotide-binding domain 1. Protein

Sci. 19, 1917–1931 (2010).

14. Wang, X. & Li, C. Decoding F508del misfolding in cystic fibrosis. Biomolecules 4, 498–

509 (2014).

15. Farinha, C. M. & Canato, S. From the endoplasmic reticulum to the plasma membrane:

mechanisms of CFTR folding and trafficking. Cell. Mol. Life Sci. 74, 39–55 (2017).

16. Grasemann, H. CFTR Modulator Therapy for Cystic Fibrosis. N. Engl. J. Med. 377,

2085–2088 (2017).

17. Ong, T. & Ramsey, B. W. Update in cystic fibrosis 2014. Am. J. Respir. Crit. Care Med.

192, 669–675 (2015).

18. Haq, I. J., Gray, M. A., Garnett, J. P., Ward, C. & Brodlie, M. Airway surface liquid

homeostasis in cystic fibrosis: pathophysiology and therapeutic targets. Thorax. 71, 284–

287 (2015).

19. Boucher, R. C., Stutts, M. J., Knowles, M. R., Cantley, L. & Gatzy, J. T. Na+ transport in

cystic fibrosis respiratory epithelia. Abnormal basal rate and response to adenylate

cyclase activation. J. Clin. Invest. 78, 1245–1252 (1986).

- 70 -

20. Boucher, R. C. Evidence for airway surface dehydration as the initiating event in CF

airway disease. J. Intern. Med. 261, 5–16 (2007).

21. Kreda, S. M., Davis, C. W. & Rose, M. C. CFTR, mucins, and mucus obstruction in

cystic fibrosis. Cold Spring Harb. Perspect. Med. 2, a009589 (2012).

22. Cohen-Cymberknoh, M., Shoseyov, D. & Kerem, E. Managing cystic fibrosis: Strategies

that increase life expectancy and improve quality of life. Am. J. Respir. Crit. Care Med.

183, 1463–1471 (2011).

23. Snell, G., Reed, A., Stern, M. & Hadjiliadis, D. The evolution of lung transplantation for

cystic fibrosis: A 2017 update. J. Cyst. Fibros. 16, 553–564 (2017).

24. McIntyre, K. Gender and survival in cystic fibrosis. Curr. Opin. Pulm. Med. 19, 392–397

(2013).

25. Ramsey, B. W. et al. A CFTR potentiator in patients with cystic fibrosis and the G551D

mutation. N. Engl. J. Med. 365, 1663–1672 (2011).

26. Van Goor, F., Hadida, S., Grootenhuis, P. D. J., Burton, B., Stack, J. H., Straley, K. S.,

Decker, C. J., Miller, M., McCartney, J., Olson, E. R., Wine, J. J., Frizzell, R. A.,

Ashlock, M. & Negulescu, P. A. Correction of the F508del-CFTR protein processing

defect in vitro by the investigational drug VX-809. Proc. Natl. Acad. Sci. 108, 18843–

18848 (2011).

27. Boyle, M. P., Bell, S. C., Konstan, M. W., McColley, S. A., Rowe, S. M., Rietschel, E.,

Huang, X., Waltz, D., Patel, N. R. & Rodman, D. A CFTR corrector (lumacaftor) and a

CFTR potentiator (ivacaftor) for treatment of patients with cystic fibrosis who have a

phe508del CFTR mutation: A phase 2 randomised controlled trial. Lancet Respir. Med. 2,

527–538 (2014).

28. Wainwright, C. E. et al. Lumacaftor–Ivacaftor in Patients with Cystic Fibrosis

Homozygous for Phe508del CFTR. N. Engl. J. Med. 373, 220–231 (2015).

29. Kerem, B., Rommens, J. M., Buchanan, J. A., Markiewicz, D., Cox, T. K., Chakravarti,

- 71 -

A., Buchwald, M. & Tsui, L. C. Identification of the cystic fibrosis gene: genetic analysis.

Science 245, 1073–1080 (1989).

30. Hacein-Bey-Abina, S. et al. Insertional oncogenesis in 4 patients after retrovirus-

mediated gene therapy of SCID-X1. J. Clin. Invest. 118, 3132–3142 (2008).

31. Howe, S. J. et al. Insertional mutagenesis combined with acquired somatic mutations

causes leukemogenesis following gene therapy of SCID-X1 patients. J. Clin. Invest. 118,

3143–3150 (2008).

32. Stein, S. et al. Genomic instability and myelodysplasia with monosomy 7 consequent to

EVI1 activation after gene therapy for chronic granulomatous disease. Nat. Med. 16, 198–

204 (2010).

33. Kang, H. J., Bartholomae, C. C., Paruzynski, A., Arens, A., Kim, S., Yu, S. S., Hong, Y.,

Joo, C. W., Yoon, N. K., Rhim, J. W., Kim, J. G., Von Kalle, C., Schmidt, M., Kim, S. &

Ahn, H. S. Retroviral gene therapy for X-linked chronic granulomatous disease: results

from phase I/II trial. Mol. Ther. 19, 2092–2101 (2011).

34. Braun, C. J. et al. Gene therapy for Wiskott-Aldrich syndrome-long-term efficacy and

genotoxicity. Sci. Transl. Med. 6, (2014).

35. Aiuti, A. et al. Lentiviral hematopoietic stem cell gene therapy in patients with wiskott-

aldrich syndrome. Science 341, (2013).

36. Hacein-Bey Abina, S. et al. Outcomes Following Gene Therapy in Patients With Severe

Wiskott-Aldrich Syndrome. JAMA 313, 1550 (2015).

37. Hacein-Bey-Abina, S. et al. A Modified γ-Retrovirus Vector for X-Linked Severe

Combined Immunodeficiency. N. Engl. J. Med. 371, 1407–1417 (2014).

38. Cavazzana-Calvo, M. et al. Transfusion independence and HMGA2 activation after gene

therapy of human β-thalassaemia. Nature 467, 318–322 (2010).

39. Cartier, N. et al. Hematopoietic stem cell gene therapy with a lentiviral vector in X-linked

- 72 -

adrenoleukodystrophy. Science 326, 818–823 (2009).

40. Biffi, A. et al. Lentiviral hematopoietic stem cell gene therapy benefits metachromatic

leukodystrophy. Science 341, (2013).

41. Ellis, J. Silencing and Variegation of gammaretrovirus and lentivirus vectors. Hum. Gene

Ther. 16, 1241–1246 (2005).

42. Rivière, I., Dunbar, C. E. & Sadelain, M. Hematopoietic stem cell engineering at a

crossroads. Blood 119, 1107–1116 (2012).

43. von Kalle, C., Deichmann, A. & Schmidt, M. Vector integration and tumorigenesis. Hum.

Gene Ther. 25, 475–481 (2014).

44. Cullis, P. R. & Hope, M. J. Lipid Nanoparticle systems for enabling gene therapies. Mol.

Ther. 25, 1467–1475 (2017).

45. del Pozo-Rodríguez, A., Delgado, D., Solinís, M. A., Gascón, A. R. & Pedraz, J. L. Solid

lipid nanoparticles for retinal gene therapy: Transfection and intracellular trafficking in

RPE cells. Int. J. Pharm. 360, 177–183 (2008).

46. Delgado, D., Del Pozo-Rodríguez, A., Solinís, M. Á., Avilés, M., Weber, B. H. F.,

Fernández, E., Rodríguez Gascón, A., Soliní, M. Á., Ferná, E. & Rodrí, A. Dextran and

protamine-based solid lipid nanoparticles as potential vectors for the treatment of X

linked juvenile retinoschisis. Hum. Gene Ther. 23, 345–355 (2012).

47. Zuris, J. A., Thompson, D. B., Shu, Y., Guilinger, J. P., Bessen, J. L., Hu, J. H., Maeder,

M. L., Joung, J. K., Chen, Z. Y. & Liu, D. R. Cationic lipid-mediated delivery of proteins

enables efficient protein-based genome editing in vitro and in vivo. Nat. Biotechnol. 33,

73–80 (2015).

48. Yin, H. et al. Therapeutic genome editing by combined viral and non-viral delivery of

CRISPR system components in vivo. Nat. Biotechnol. 34, 328–333 (2016).

49. High, K. H., Nathwani, A., Spencer, T. & Lillicrap, D. Current status of haemophilia

- 73 -

gene therapy. Haemophilia 20, 43–49 (2014).

50. Nathwani, A. C. et al. Long-Term Safety and Efficacy of Factor IX Gene Therapy in

Hemophilia B. N. Engl. J. Med. 371, 1994–2004 (2014).

51. Russell, S. et al. Efficacy and safety of voretigene neparvovec (AAV2-hRPE65v2) in

patients with RPE65-mediated inherited retinal dystrophy: a randomised, controlled,

open-label, phase 3 trial. Lancet 390, 849–860 (2017).

52. Rosenfeld, M. a, Yoshimura, K., Trapnell, B. C., Yoneyama, K., Rosenthal, E. R.,

Dalemans, W., Fukayama, M., Bargon, J., Stier, L. E. & Stratford-Perricaudet, L. In vivo

transfer of the human cystic fibrosis transmembrane conductance regulator gene to the

airway epithelium. Cell 68, 143–55 (1992).

53. Volpers, C. & Kochanek, S. Adenoviral vectors for gene transfer and therapy. J. Gene

Med. 6, S164–S171 (2004).

54. Hyde, S. C., Gill, D. R., Higgins, C. F., Trezise, A. E. O., MacVinish, L. J., Cuthbert, A.

W., Ratcliff, R., Evans, M. J. & Colledge, W. H. Correction of the ion transport defect in

cystic fibrosis transgenic mice by gene therapy. Nature 362, 250–255 (1993).

55. Zabner, J., Couture, L. A., Gregory, R. J., Graham, S. M., Smith, A. E. & Welsh, M. J.

Adenovirus-mediated gene transfer transiently corrects the chloride transport defect in

nasal epithelia of patients with cystic fibrosis. Cell 75, 207–216 (1993).

56. Caplen, N. J., Alton, E. W., Middleton, P. G., Dorin, J. R., Stevenson, B. J., Gao, X.,

Durham, S. R., Jeffery, P. K., Hodson, M. E. & Coutelle, C. Liposome-mediated CFTR

gene transfer to the nasal epithelium of patients with cystic fibrosis. Nat. Med. 1, 39–46

(1995).

57. Crystal, R. G., McElvaney, N. G., Rosenfeld, M. A., Chu, C. S., Mastrangeli, A., Hay, J.

G., Brody, S. L., Jaffe, H. A., Eissa, N. T. & Danel, C. Administration of an adenovirus

containing the human CFTR cDNA to the respiratory tract of individuals with cystic

fibrosis. Nat. Genet. 8, 42–51 (1994).

- 74 -

58. Knowles, M. R., Hohneker, K. W., Zhou, Z., Olsen, J. C., Noah, T. L., Hu, P. C., Leigh,

M. W., Engelhardt, J. F., Edwards, L. J. & Jones, K. R. A controlled study of adenoviral-

vector-mediated gene transfer in the nasal epithelium of patients with cystic fibrosis. N.

Engl. J. Med. 333, 823–831 (1995).

59. Hay, J. G., McElvaney, N. G., Herena, J. & Crystal, R. G. Modification of Nasal

Epithelial Potential Differences of Individuals with Cystic Fibrosis Consequent to Local

Administration of a Normal CFTR cDNA Adenovirus Gene Transfer Vector. Hum. Gene

Ther. 6, 1487–1496 (1995).

60. Zabner, J., Ramsey, B. W., Meeker, D. P., Aitken, M. L., Balfour, R. P., Gibson, R. L.,

Launspach, J., Moscicki, R. A., Richards, S. M. & Standaert, T. A. Repeat administration

of an adenovirus vector encoding cystic fibrosis transmembrane conductance regulator to

the nasal epithelium of patients with cystic fibrosis. J. Clin. Invest. 97, 1504–11 (1996).

61. Bellon, G. et al. Aerosol administration of a recombinant adenovirus expressing CFTR to

cystic fibrosis patients: a phase I clinical trial. Hum. Gene Ther. 8, 15–25 (1997).

62. Harvey, B. G., Leopold, P. L., Hackett, N. R., Grasso, T. M., Williams, P. M., Tucker, A.

L., Kaner, R. J., Ferris, B., Gonda, I., Sweeney, T. D., Ramalingam, R., Kovesdi, I., Shak,

S. & Crystal, R. G. Airway epithelial CFTR mRNA expression in cystic fibrosis patients

after repetitive administration of a recombinant adenovirus. J. Clin. Invest. 104, 1245–55

(1999).

63. Zuckerman, J. B., Robinson, C. B., McCoy, K. S., Shell, R., Sferra, T. J., Chirmule, N.,

Magosin, S. A., Propert, K. J., Brown-Parr, E. C., Hughes, J. V, Tazelaar, J., Baker, C.,

Goldman, M. J. & Wilson, J. M. A phase I study of adenovirus-mediated transfer of the

human cystic fibrosis transmembrane conductance regulator gene to a lung segment of

individuals with cystic fibrosis. Hum. Gene Ther. 10, 2973–85 (1999).

64. Joseph, P. M., O’Sullivan, B. P., Lapey, A., Dorkin, H., Oren, J., Balfour, R., Perricone,

M. A., Rosenberg, M., Wadsworth, S. C., Smith, A. E., St Georg, J. A. & Meeker, D. P.

Aerosol and lobar administration of a recombinant adenovirus to individuals with cystic

fibrosis. I. Methods, safety, and clinical implications. Hum. Gene Ther. 12, 1369–1382

- 75 -

(2001).

65. Perricone, M. A., Morris, J. E., Pavelka, K., Plog, M. S., O’Sullivan, B. P., Joseph, P. M.,

Dorkin, H., Lapey, A., Balfour, R., Meeker, D. P., Smith, A. E., Wadsworth, S. C. & St

Georg, J. A. Aerosol and lobar administration of a recombinant adenovirus to individuals

with cystic fibrosis. II. Transfection efficiency in airway epithelium. Hum. Gene Ther. 12,

1383–1394 (2001).

66. Samulski, R. J. & Muzyczka, N. AAV-Mediated Gene Therapy for Research and

Therapeutic Purposes. Annu. Rev. Virol. 1, 427–451 (2014).

67. Wagner, J. A., Moran, M. L., Messner, A. H., Daifuku, R., Conrad, C. K., Reynolds, T.,

Guggino, W. B., Moss, R. B., Carter, B. J., Wine, J. J., Flotte, T.R. & Gardner, P. A

Phase I/II Study of tgAAV-CF for the Treatment of Chronic Sinusitis in Patients with

Cystic Fibrosis. Stanford University, Stanford, California. Hum. Gene Ther. 9, 889–909

(1998).

68. Wagner, J. a et al. Safety and biological efficacy of an adeno-associated virus vector-

cystic fibrosis transmembrane regulator (AAV-CFTR) in the cystic fibrosis maxillary

sinus. Laryngoscope 109, 266–74 (1999).

69. Wagner, J. A. et al. A Phase II, Double-Blind, Randomized, Placebo-Controlled Clinical

Trial of tgAAVCF Using Maxillary Sinus Delivery in Patients with Cystic Fibrosis with

Antrostomies. Hum. Gene Ther. 13, 1349–1359 (2002).

70. Aitken, M. L., Moss, R. B., Waltz, D. A., Dovey, M. E., Tonelli, M. R., McNamara, S.

C., Gibson, R. L., Ramsey, B. W., Carter, B. J. & Reynolds, T. C. A Phase I Study of

Aerosolized Administration of tgAAVCF to Cystic Fibrosis Subjects with Mild Lung

Disease. Hum. Gene Ther. 12, 1907–1916 (2001).

71. Flotte, T. R., Zeitlin, P. L., Reynolds, T. C., Heald, A. E., Pedersen, P., Beck, S., Conrad,

C. K., Brass-Ernst, L., Humphries, M., Sullivan, K., Wetzel, R., Taylor, G., Carter, B.J. &

Guggino, W.B. Phase I trial of intranasal and endobronchial administration of a

recombinant adeno-associated virus serotype 2 (rAAV2)-CFTR vector in adult cystic

- 76 -

fibrosis patients: a two-part clinical study. Hum. Gene Ther. 14, 1079–1088 (2003).

72. Moss, R. B. et al. Repeated Aerosolized AAV-CFTR for Treatment of Cystic Fibrosis: A

Randomized Placebo-Controlled Phase 2B Trial. Hum. Gene Ther. 18, 726–732 (2007).

73. Kobinger, G. P., Weiner, D. J., Yu, Q. C. & Wilson, J. M. Filovirus-pseudotyped

lentiviral vector can efficiently and stably transduce airway epithelia in vivo. Nat.

Biotechnol. 19, 225–230 (2001).

74. Mitomo, K. et al. Toward gene therapy for cystic fibrosis using a lentivirus pseudotyped

with Sendai virus envelopes. Mol. Ther. 18, 1173–82 (2010).

75. Sinn, P. L., Cooney, A. L., Oakland, M., Dylla, D. E., Wallen, T. J., Pezzulo, A. A.,

Chang, E. H. & McCray, P. B. Lentiviral vector gene transfer to porcine airways. Mol.

Ther. Nucleic Acids 1, e56 (2012).

76. Griesenbach, U. et al. Assessment of F/HN-pseudotyped lentivirus as a clinically relevant

vector for lung gene therapy. Am. J. Respir. Crit. Care Med. 186, 846–856 (2012).

77. Patel, M., Giddings, A. M., Sechelski, J. & Olsen, J. C. High efficiency gene transfer to

airways of mice using influenza hemagglutinin pseudotyped lentiviral vectors. J. Gene

Med. 15, 51–62 (2013).

78. Parks, R. J., Chen, L., Anton, M., Sankar, U., Rudnicki, M. A. & Graham, F. L. A helper-

dependent adenovirus vector system: removal of helper virus by Cre-mediated excision of

the viral packaging signal. Proc. Natl. Acad. Sci. U. S. A. 93, 13565–70 (1996).

79. Toietta, G., Koehler, D. R., Finegold, M. J., Lee, B., Hu, J. & Beaudet, A. L. Reduced

inflammation and improved airway expression using helper-dependent adenoviral vectors

with a K18 promoter. Mol. Ther. 7, 649–658 (2003).

80. Koehler, D. R., Sajjan, U., Chow, Y.-H., Martin, B., Kent, G., Tanswell, A. K., McKerlie,

C., Forstner, J. F. & Hu, J. Protection of Cftr knockout mice from acute lung infection by

a helper-dependent adenoviral vector expressing Cftr in airway epithelia. Proc. Natl.

Acad. Sci. U. S. A. 100, 15364–9 (2003).

- 77 -

81. Farmen, S. L., Karp, P. H., Ng, P., Palmer, D. J., Koehler, D. R., Hu, J., Beaudet, A. L.,

Zabner, J. & Welsh, M. J. Gene transfer of CFTR to airway epithelia: low levels of

expression are sufficient to correct Cl- transport and overexpression can generate

basolateral CFTR. Am. J. Physiol. Lung Cell. Mol. Physiol. 289, L1123-30 (2005).

82. Cao, H., Ouyang, H., Ip, W., Du, K., Duan, W., Avolio, J., Wu, J., Duan, C., Yeger, H.,

Bear, C. E., Gonska, T., Hu, J. & Moraes, T. J. Testing gene therapy vectors in human

primary nasal epithelial cultures. Mol. Ther. - Methods Clin. Dev. 2, 15034 (2015).

83. Cao, H. et al. Efficient gene delivery to pig airway epithelia and submucosal glands using

helper-dependent adenoviral vectors. Mol. Ther. - Nucleic Acids 2, (2013).

84. Rawlins, E. L. & Hogan, B. L. M. Epithelial stem cells of the lung: privileged few or

opportunities for many? Development 133, 2455–65 (2006).

85. Cao, H., Ouyang, H., Grasemann, H., Bartlett, C., Du, K., Duan, R., Shi, F., Estrada, M.,

Seigel, K., Coates, A., Yeger, H., Bear, CE., Gonska, T., Moraes, T.J. & Hu, J.

Transducing airway basal cells with a helper-dependent adenoviral vector for lung gene

therapy. Hum. Gene Ther. (2018) doi:10.1089/hum.2017.201.

86. Muller, H. J. Artificial transmutation of the gene. Science 66, 84–87 (1927).

87. Auerbach, C. & Robson, J. M. Chemical production of mutations. Nature 157, 302

(1946).

88. Scherer, S. & Davis, R. W. Replacement of chromosome segments with altered DNA

sequences constructed in vitro. Proc. Natl. Acad. Sci. U. S. A. 76, 4951–5 (1979).

89. Rothstein, R. J. One-Step Gene Disruption in Yeast. Methods Enzymol. 101, 202–211

(1983).

90. Smithies, O., Gregg, R. G., Boggs, S. S., Koralewski, M. A. & Kucherlapati, R. S.

Insertion of DNA sequences into the human chromosomal β-globin locus by homologous

recombination. Nature 317, 230–234 (1985).

- 78 -

91. Thomas, K. R., Folger, K. R. & Capecchi, M. R. High frequency targeting of genes to

specific sites in the mammalian genome. Cell 44, 419–428 (1986).

92. Latt, S. Sister chromatid exchange formation. Annu. Rev. Genet. 15, 11–55 (1981).

93. Padmore, R., Cao, L. & Kleckner, N. Temporal comparison of recombination and

synaptonemal complex formation during meiosis in S. cerevisiae. Cell 66, 1239–1256

(1991).

94. Plessis, A., Perrin, A., Haber, J. E. & Dujon, B. Site-specific recombination determined

by I-SceI, a mitochondrial group I intron-encoded endonuclease expressed in the yeast

nucleus. Genetics 130, 451–460 (1992).

95. Choulika, A., Perrin, A., Dujon, B. & Ois Nicolas, J. Induction of homologous

recombination in mammalian chromosomes by using the I-scei system of saccharomyces

cerevisiae. Mol. Cell. Biol. 15, 1968–1973 (1995).

96. Rouet, P., Smih, F. & Jasin, M. Introduction of double-strand breaks into the genome of

mouse cells by expression of a rare-cutting endonuclease. Mol. Cell. Biol. 14, 8096–106

(1994).

97. Pavletich, N. P. & Pabo, C. O. Zinc finger-DNA recognition: Crystal structure of a

Zif268-DNA complex at 2.1 Å. Science 252, 809–817 (1991).

98. Klug, A. The Discovery of Zinc Fingers and Their Applications in Gene Regulation and

Genome Manipulation. Annu. Rev. Biochem. 79, 213–231 (2010).

99. Liu, Q., Segal, D. J., Ghiara, J. B. & Barbas, C. F. Design of polydactyl zinc-finger

proteins for unique addressing within complex genomes. Proc. Natl. Acad. Sci. 94, 5525–

5530 (1997).

100. Kim, Y. G., Cha, J. & Chandrasegaran, S. Hybrid restriction enzymes: zinc finger fusions

to Fok I cleavage domain. Proc. Natl. Acad. Sci. 93, 1156–1160 (1996).

101. Isalan, M., Choo, Y. & Klug, A. Synergy between adjacent zinc fingers in sequence-

- 79 -

specific DNA recognition. Proc. Natl. Acad. Sci. U. S. A. 94, 5617–21 (1997).

102. Isalan, M., Klug, A. & Choo, Y. Comprehensive DNA recognition through concerted

interactions from adjacent zinc fingers. Biochemistry 37, 12026–12033 (1998).

103. Miller, J. C. & Pabo, C. O. Rearrangement of side-chains in a zif268 mutant highlights

the complexities of zinc finger-DNA recognition. J. Mol. Biol. 313, 309–315 (2001).

104. Pabo, C., Peisach, E. & Grant, R. Design and selection of novel cys2his2 zinc finger

proteins. Annu. Rev. Biochem. 70, 313–340 (2001).

105. Dutta, S., Madan, S. & Sundar, D. Exploiting the recognition code for elucidating the

mechanism of zinc finger protein-DNA interactions. BMC Genomics 17, (2016) doi:

10.1186/s12864-016-3324-8.

106. Moore, F. E., Reyon, D., Sander, J. D., Martinez, S. A., Blackburn, J. S., Khayter, C.,

Ramirez, C. L., Joung, J. K. & Langenau, D. M. Improved somatic mutagenesis in

zebrafish using transcription activator-like effector nucleases (TALENs). PLoS One 7,

e37877 (2012).

107. Yang, D. et al. Generation of PPARγ mono-allelic knockout pigs via zinc-finger

nucleases and nuclear transfer cloning. Cell Res. 21, 979–982 (2011).

108. Liu, X., Wang, Y., Guo, W., Chang, B., Liu, J., Guo, Z., Quan, F. & Zhang, Y. Zinc-

finger nickase-mediated insertion of the lysostaphin gene into the beta-casein locus in

cloned cows. Nat. Commun. 4, 2565 (2013).

109. Doyon, Y., McCammon, J. M., Miller, J. C., Faraji, F., Ngo, C., Katibah, G. E., Amora,

R., Hocking, T. D., Zhang, L., Rebar, E. J., Gregory, P. D., Urnov, F. D. & Amacher, S.

L. Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases.

Nat. Biotechnol. 26, 702–708 (2008).

110. Carbery, I. D., Ji, D., Harrington, A., Brown, V., Weinstein, E. J., Liaw, L. & Cui, X.

Targeted genome modification in mice using zinc-finger nucleases. Genetics 186, 451–

459 (2010).

- 80 -

111. Hockemeyer, D. et al. Efficient targeting of expressed and silent genes in human ESCs

and iPSCs using zinc-finger nucleases. Nat. Biotechnol. 27, 851–857 (2009).

112. Li, H. et al. In vivo genome editing restores haemostasis in a mouse model of

haemophilia. Nature 475, 217–221 (2011).

113. Hoban, M. D. et al. Correction of the sickle cell disease mutation in human hematopoietic

stem/progenitor cells. Blood 125, 2597–2604 (2015).

114. Yusa, K. et al. Targeted gene correction of α1-antitrypsin deficiency in induced

pluripotent stem cells. Nature 478, 391–394 (2011).

115. Soldner, F. et al. Generation of isogenic pluripotent stem cells differing exclusively at

two early onset parkinson point mutations. Cell 146, 318–331 (2011).

116. Tebas, P. et al. Gene editing of CCR5 in autologous CD4 T cells of persons infected with

HIV. N. Engl. J. Med. 370, 901–910 (2014).

117. Kay, S., Hahn, S., Marois, E., Hause, G. & Bonas, U. A bacterial effector acts as a plant

transcription factor and induces a cell size regulator. Science 318, 648–651 (2007).

118. Sugio, A., Yang, B., Zhu, T. & White, F. F. Two type III effector genes of Xanthomonas

oryzae pv. oryzae control the induction of the host genes OsTFIIAgamma1 and OsTFX1

during bacterial blight of rice. Proc. Natl. Acad. Sci. U. S. A. 104, 10720–5 (2007).

119. White, F. F. & Yang, B. Host and pathogen factors controlling the rice-xanthomonas

oryzae interaction. Plant Physiol. 150, 1677–1686 (2009).

120. Kay, S. & Bonas, U. How Xanthomonas type III effectors manipulate the host plant.

Curr. Opin. Microbiol. 12, 37–43 (2009).

121. Moscou, M. J. & Bogdanove, A. J. A simple cipher governs DNA recognition by TAL

effectors. Science 326, 1501 (2009).

122. Boch, J., Scholze, H., Schornack, S., Landgraf, A., Hahn, S., Kay, S., Lahaye, T.,

Nickstadt, A. & Bonas, U. Breaking the code of DNA binding specificity of TAL-type III

- 81 -

effectors. Science 326, 1509–1512 (2009).

123. Deng, D., Yan, C., Pan, X., Mahfouz, M., Wang, J., Zhu, J. K., Shi, Y. & Yan, N.

Structural basis for sequence-specific recognition of DNA by TAL effectors. Science 335,

720–723 (2012).

124. Christian, M., Cermak, T., Doyle, E. L., Schmidt, C., Zhang, F., Hummel, A.,

Bogdanove, A. J. & Voytas, D. F. Targeting DNA double-strand breaks with TAL

effector nucleases. Genetics 186, 756–761 (2010).

125. Miller, J. C. et al. A TALE nuclease architecture for efficient genome editing. Nat.

Biotechnol. 29, 143–150 (2011).

126. Li, T., Huang, S., Jiang, W. Z., Wright, D., Spalding, M. H., Weeks, D. P. & Yang, B.

TAL nucleases (TALNs): Hybrid proteins composed of TAL effectors and FokI DNA-

cleavage domain. Nucleic Acids Res. 39, 359–372 (2011).

127. Holkers, M., Maggio, I., Liu, J., Janssen, J. M., Miselli, F., Mussolino, C., Recchia, A.,

Cathomen, T. & Gonçalves, M. A. F. V. Differential integrity of TALE nuclease genes

following adenoviral and lentiviral vector gene transfer into human cells. Nucleic Acids

Res. 41, (2013).

128. Reyon, D., Tsai, S. Q., Khgayter, C., Foden, J. A., Sander, J. D. & Joung, J. K. FLASH

assembly of TALENs for high-throughput genome editing. Nat. Biotechnol. 30, 460–465

(2012).

129. Zhang, Z., Zhang, S., Huang, X., Orwig, K. E. & Sheng, Y. Rapid assembly of

customized TALENs into multiple delivery systems. PLoS One 8, e80281 (2013).

130. Cermak, T., Starker, C. G. & Voytas, D. F. Efficient design and assembly of custom

talens using the golden gate platform. Chromosomal Mutagenesis: Second Edition 133–

159 (2014). doi:10.1007/978-1-4939-1862-1_7

131. Kim, Y. et al. A library of TAL effector nucleases spanning the human genome. Nat.

Biotechnol. 31, 251–258 (2013).

- 82 -

132. Wood, A. J., Lo, T. W., Zeitler, B., Pickle, C. S., Ralston, E. J., Lee, A. H., Amora, R.,

Miller, J. C., Leung, E., Meng, X., Zhang, L., Rebar, E. J., Gregory, P. D., Urnov, F. D.

& Meyer, B. J. Targeted genome editing across species using ZFNs and TALENs.

Science 333, 307 (2011).

133. Ma, S., Zhang, S., Wang, F., Liu, Y., Liu, Y., Xu, H., Liu, C., Lin, Y., Zhao, P. & Xia, Q.

Highly efficient and specific genome editing in silkworm using custom TALENs. PLoS

One 7, e45035 (2012).

134. Yu, Z. et al. Various applications of TALEN- and CRISPR/Cas9-mediated homologous

recombination to modify the Drosophila genome. Biol. Open 3, 271–280 (2014).

135. Lei, Y., Guo, X., Liu, Y., Cao, Y., Deng, Y., Chen, X., Cheng, C. H. K., Dawid, I. B.,

Chen, Y. & Zhao, H. Efficient targeted gene disruption in Xenopus embryos using

engineered transcription activator-like effector nucleases (TALENs). Proc. Natl. Acad.

Sci. 109, 17484–17489 (2012).

136. Watanabe, T., Ochiai, H., Sakuma, T., Horch, H. W., Hamaguchi, N., Nakamura, T.,

Bando, T., Ohuchi, H., Yamamoto, T., Noji, S. & Mito, T. Non-transgenic genome

modifications in a hemimetabolous insect using zinc-finger and TAL effector nucleases.

Nat. Commun. 3, 1017 (2012).

137. Sander, J. D., Cade, L., Khayter, C., Reyon, D., Peterson, R. T., Joung, J. K. & Yeh, J. R.

J. Targeted gene disruption in somatic zebrafish cells using engineered TALENs. Nat.

Biotechnol. 29, 697–698 (2011).

138. Tesson, L., Usal, C., Meq́noret, S., Leung, E., Niles, B. J., Remy, S., Santiago, Y.,

Vincent, A. I., Meng, X., Zhang, L., Gregory, P. D., Anegon, I. & Cost, G. J. Knockout

rats generated by embryo microinjection of TALENs. Nat. Biotechnol. 29, 695–696

(2011).

139. Carlson, D. F., Tan, W., Lillico, S. G., Stverakova, D., Proudfoot, C., Christian, M.,

Voytas, D. F., Long, C. R., Whitelaw, C. B. A. & Fahrenkrug, S. C. Efficient TALEN-

mediated gene knockout in livestock. Proc. Natl. Acad. Sci. 109, 17382–17387 (2012).

- 83 -

140. Li, T., Liu, B., Spalding, M. H., Weeks, D. P. & Yang, B. High-efficiency TALEN-based

gene editing produces disease-resistant rice. Nat. Biotechnol. 30, 390–392 (2012).

141. Hockemeyer, D. et al. Genetic engineering of human pluripotent cells using TALE

nucleases. Nat. Biotechnol. 29, 731–734 (2011).

142. Ye, L., Wang, J., Beyer, A. I., Teque, F., Cradick, T. J., Qi, Z., Chang, J. C., Bao, G.,

Muench, M. O., Yu, J., Levy, J. A. & Kan, Y. W. Seamless modification of wild-type

induced pluripotent stem cells to the natural CCR5Δ32 mutation confers resistance to

HIV infection. Proc. Natl. Acad. Sci. 111, 9591–9596 (2014).

143. Bloom, K., Ely, A., Mussolino, C., Cathomen, T. & Arbuthnot, P. Inactivation of

Hepatitis B virus replication in cultured cells and in vivo with engineered transcription

activator-like effector nucleases. Mol. Ther. 21, 1889–1897 (2013).

144. Ishino, Y., Shinagawa, H., Makino, K., Amemura, M. & Nakata, A. Nucleotide sequence

of the iap gene, responsible for alkaline phosphatase isozyme conversion in Escherichia

coli, and identification of the gene product. J. Bacteriol. 169, 5429–33 (1987).

145. Mojica, F. J. M., Díez-Villaseñor, C., Soria, E. & Juez, G. Biological significance of a

family of regularly spaced repeats in the genomes of Archaea, Bacteria and mitochondria.

Mol. Microbiol. 36, 244–246 (2000).

146. Jansen, R., Embden, J. D. A. van, Gaastra, W. & Schouls, L. M. Identification of genes

that are associated with DNA repeats in prokaryotes. Mol. Microbiol. 43, 1565–1575

(2002).

147. Mojica, F. J. M., Díez-Villaseñor, C., García-Martínez, J. & Soria, E. Intervening

sequences of regularly spaced prokaryotic repeats derive from foreign genetic elements.

J. Mol. Evol. 60, 174–182 (2005).

148. Pourcel, C., Salvignol, G. & Vergnaud, G. CRISPR elements in Yersinia pestis acquire

new repeats by preferential uptake of bacteriophage DNA, and provide additional tools

for evolutionary studies. Microbiology 151, 653–663 (2005).

- 84 -

149. Bolotin, A., Quinquis, B., Sorokin, A. & Dusko Ehrlich, S. Clustered regularly

interspaced short palindrome repeats (CRISPRs) have spacers of extrachromosomal

origin. Microbiology 151, 2551–2561 (2005).

150. Barrangou, R., Fremaux, C., Deveau, H., Richards, M., Boyaval, P., Moineau, S.,

Romero, D. A. & Horvath, P. CRISPR provides acquired resistance against viruses in

prokaryotes. Science 315, 1709–1712 (2007).

151. Marraffini, L. A. & Sontheimer, E. J. CRISPR interference limits horizontal gene transfer

in staphylococci by targeting DNA. Science 322, 1843–1845 (2008).

152. Makarova, K. S. et al. An updated evolutionary classification of CRISPR-Cas systems.

Nat. Rev. Microbiol. 13, 722–736 (2015).

153. Nuñez, J. K., Kranzusch, P. J., Noeske, J., Wright, A. V., Davies, C. W. & Doudna, J. A.

Cas1-Cas2 complex formation mediates spacer acquisition during CRISPR-Cas adaptive

immunity. Nat. Struct. Mol. Biol. 21, 528–534 (2014).

154. Deveau, H., Barrangou, R., Garneau, J. E., Labonté, J., Fremaux, C., Boyaval, P.,

Romero, D. A., Horvath, P. & Moineau, S. Phage response to CRISPR-encoded

resistance in Streptococcus thermophilus. J. Bacteriol. 190, 1390–1400 (2008).

155. Shah, S. A., Erdmann, S., Mojica, F. J. M. & Garrett, R. A. Protospacer recognition

motifs: Mixed identities and functional diversity. RNA Biology 10, 891–899 (2013).

156. Staals, R. H. J. et al. Structure and activity of the RNA-targeting type III-B CRISPR-Cas

complex of Thermus thermophilus. Mol. Cell 52, 135–145 (2013).

157. Staals, R. H. J. et al. RNA Targeting by the type III-A CRISPR-Cas csm complex of

Thermus thermophilus. Mol. Cell 56, 518–530 (2014).

158. Tamulaitis, G., Kazlauskiene, M., Manakova, E., Venclovas, Č., Nwokeoji, A. O.,

Dickman, M. J., Horvath, P. & Siksnys, V. Programmable RNA shredding by the type

III-A CRISPR-Cas system of Streptococcus thermophilus. Mol. Cell 56, 506–517 (2014).

- 85 -

159. Jinek, M., Jiang, F., Taylor, D. W., Sternberg, S. H., Kaya, E., Ma, E., Anders, C., Hauer,

M., Zhou, K., Lin, S., Kaplan, M., Iavarone, A. T., Charpentier, E., Nogales, E. &

Doudna, J. A. Structures of Cas9 endonucleases reveal RNA-mediated conformational

activation. Science 343, (2014).

160. Wang, R., Preamplume, G., Terns, M. P., Terns, R. M. & Li, H. Interaction of the Cas6

riboendonuclease with CRISPR RNAs: Recognition and cleavage. Structure 19, 257–264

(2011).

161. Deltcheva, E., Chylinski, K., Sharma, C. M., Gonzales, K., Chao, Y., Pirzada, Z. A.,

Eckert, M. R., Vogel, J. & Charpentier, E. CRISPR RNA maturation by trans-encoded

small RNA and host factor RNase III. Nature 471, 602–607 (2011).

162. Huo, Y., Nam, K. H., Ding, F., Lee, H., Wu, L., Xiao, Y., Farchione, M. D., Zhou, S.,

Rajashankar, K., Kurinov, I., Zhang, R. & Ke, A. Structures of CRISPR Cas3 offer

mechanistic insights into Cascade-activated DNA unwinding and degradation. Nat.

Struct. Mol. Biol. 21, 771–777 (2014).

163. Jinek, M., Chylinski, K., Fonfara, I., Hauer, M., Doudna, J. A. & Charpentier, E. A

programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity.

Science 337, 816–821 (2012).

164. Gasiunas, G., Barrangou, R., Horvath, P. & Siksnys, V. Cas9-crRNA ribonucleoprotein

complex mediates specific DNA cleavage for adaptive immunity in bacteria. Proc. Natl.

Acad. Sci. 109, 2579–2586 (2012).

165. Sapranauskas, R., Gasiunas, G., Fremaux, C., Barrangou, R., Horvath, P. & Siksnys, V.

The Streptococcus thermophilus CRISPR/Cas system provides immunity in Escherichia

coli. Nucleic Acids Res. 39, 9275–9282 (2011).

166. Magadán, A. H., Dupuis, M. È., Villion, M. & Moineau, S. Cleavage of phage DNA by

the Streptococcus thermophilus CRISPR3-Cas system. PLoS One 7, e40913 (2012).

167. Jiang, W., Bikard, D., Cox, D., Zhang, F. & Marraffini, L. A. RNA-guided editing of

bacterial genomes using CRISPR-Cas systems. Nat. Biotechnol. 31, 233–239 (2013).

- 86 -

168. Dicarlo, J. E., Norville, J. E., Mali, P., Rios, X., Aach, J. & Church, G. M. Genome

engineering in Saccharomyces cerevisiae using CRISPR-Cas systems. Nucleic Acids Res.

41, 4336–4343 (2013).

169. Cong, L. et al. Multiplex Genome Engineering Using CRISPR/Cas Systems. Science

339, 819–823 (2013).

170. Mali, P., Yang, L., Esvelt, K. M., Aach, J., Guell, M., DiCarlo, J. E., Norville, J. E. &

Church, G. M. RNA-guided human genome engineering via Cas9. Science 339, 823–826

(2013).

171. Ruan, J., Li, H., Xu, K., Wu, T., Wei, J., Zhou, R., Liu, Z., Mu, Y., Yang, S., Ouyang, H.,

Chen-Tsai, R. Y. & Li, K. Highly efficient CRISPR/Cas9-mediated transgene knockin at

the H11 locus in pigs. Sci. Rep. 5, 14253 (2015).

172. Wang, K., Ouyang, H., Xie, Z., Yao, C., Guo, N., Li, M., Jiao, H. & Pang, D. Efficient

Generation of Myostatin Mutations in Pigs Using the CRISPR/Cas9 System. Sci. Rep. 5,

16623 (2015).

173. Jeong, Y.-H. et al. Knock-in fibroblasts and transgenic blastocysts for expression of

human FGF2 in the bovine β-casein gene locus using CRISPR/Cas9 nuclease-mediated

homologous recombination. Zygote 24, 442–456 (2016).

174. Crispo, M., Mulet, A. P., Tesson, L., Barrera, N., Cuadro, F., Dos Santos-Neto, P. C.,

Nguyen, T. H., Crénéguy, A., Brusselle, L., Anegón, I. & Menchaca, A. Efficient

generation of myostatin knock-out sheep using CRISPR/Cas9 technology and

microinjection into zygotes. PLoS One 10, e0136690 (2015).

175. Ni, W., Qiao, J., Hu, S., Zhao, X., Regouski, M., Yang, M., Polejaeva, I. A. & Chen, C.

Efficient gene knockout in goats using CRISPR/Cas9 system. PLoS One 9, e106718

(2014).

176. Wang, X. et al. Generation of gene-modified goats targeting MSTN and FGF5 via zygote

injection of CRISPR/Cas9 system. Sci. Rep. 5, 13878 (2015).

- 87 -

177. Nishimasu, H., Ran, F. A., Hsu, P. D., Konermann, S., Shehata, S. I., Dohmae, N.,

Ishitani, R., Zhang, F. & Nureki, O. Crystal structure of Cas9 in complex with guide

RNA and target DNA. Cell 156, 935–949 (2014).

178. Ariyoshi, M., Vassylyev, D. G., Iwasaki, H., Nakamura, H., Shinagawa, H. & Morikawa,

K. Atomic structure of the RuvC resolvase: A holliday junction-specific endonuclease

from E. coli. Cell 78, 1063–1072 (1994).

179. Górecka, K. M., Komorowska, W. & Nowotny, M. Crystal structure of RuvC resolvase in

complex with Holliday junction substrate. Nucleic Acids Res. 41, 9945–9955 (2013).

180. Biertümpfel, C., Yang, W. & Suck, D. Crystal structure of T4 endonuclease VII resolving

a Holliday junction. Nature 449, 616–620 (2007).

181. Li, C. L., Hor, L. I., Chang, Z. F., Tsai, L. C., Yang, W. Z. & Yuan, H. S. DNA binding

and cleavage by the periplasmic nuclease Vvn: A novel structure with a known active

site. EMBO J. 22, 4014–4025 (2003).

182. Jiang, F., Taylor, D. W., Chen, J. S., Kornfeld, J. E., Zhou, K., Thompson, A. J., Nogales,

E. & Doudna, J. A. Structures of a CRISPR-Cas9 R-loop complex primed for DNA

cleavage. Science 351, 867–871 (2016).

183. Sternberg, S. H., Redding, S., Jinek, M., Greene, E. C. & Doudna, J. A. DNA

interrogation by the CRISPR RNA-guided endonuclease Cas9. Nature 507, 62–67

(2014).

184. Semenova, E., Jore, M. M., Datsenko, K. A., Semenova, A., Westra, E. R., Wanner, B.,

van der Oost, J., Brouns, S. J. J. & Severinov, K. Interference by clustered regularly

interspaced short palindromic repeat (CRISPR) RNA is governed by a seed sequence.

Proc. Natl. Acad. Sci. 108, 10098–10103 (2011).

185. Hsu, P. D., Scott, D. A., Weinstein, J. A., Ran, F. A., Konermann, S., Agarwala, V., Li,

Y., Fine, E. J., Wu, X., Shalem, O., Cradick, T. J., Marraffini, L. A., Bao, G. & Zhang, F.

DNA targeting specificity of RNA-guided Cas9 nucleases. Nat. Biotechnol. 31, 827–832

(2013).

- 88 -

186. Bikard, D., Jiang, W., Samai, P., Hochschild, A., Zhang, F. & Marraffini, L. A.

Programmable repression and activation of bacterial gene expression using an engineered

CRISPR-Cas system. Nucleic Acids Res. 41, 7429–7437 (2013).

187. Qi, L. S., Larson, M. H., Gilbert, L. A., Doudna, J. A., Weissman, J. S., Arkin, A. P. &

Lim, W. A. Repurposing CRISPR as an RNA-guided platform for sequence-specific

control of gene expression. Cell 152, 1173–1183 (2013).

188. Maeder, M. L., Linder, S. J., Cascio, V. M., Fu, Y., Ho, Q. H. & Joung, J. K. CRISPR

RNA-guided activation of endogenous human genes. Nat. Methods 10, 977–979 (2013).

189. Perez-Pinera, P., Kocak, D. D., Vockley, C. M., Adler, A. F., Kabadi, A. M., Polstein, L.

R., Thakore, P. I., Glass, K. A., Ousterout, D. G., Leong, K. W., Guilak, F., Crawford, G.

E., Reddy, T. E. & Gersbach, C. A. RNA-guided gene activation by CRISPR-Cas9-based

transcription factors. Nat. Methods 10, 973–976 (2013).

190. Zhou, H. et al. In vivo simultaneous transcriptional activation of multiple genes in the

brain using CRISPR–dCas9-activator transgenic mice. Nat. Neurosci. 21, 440–46 (2018).

191. Hilton, I. B., D’Ippolito, A. M., Vockley, C. M., Thakore, P. I., Crawford, G. E., Reddy,

T. E. & Gersbach, C. A. Epigenome editing by a CRISPR-Cas9-based acetyltransferase

activates genes from promoters and enhancers. Nat. Biotechnol. 33, 510–517 (2015).

192. Gilbert, L. A, Larson, M. H., Morsut, L., Liu, Z., Gloria, A., Torres, S. E., Stern-ginossar,

N., Brandman, O., Whitehead, H., Doudna, J. a, Lim, W. A & Jonathan, S. CRISPR-

mediated modular RNA-guided regualtion of transcription in eukaryotes. Cell 154, 442–

451 (2013).

193. Guilinger, J. P., Thompson, D. B. & Liu, D. R. Fusion of catalytically inactive Cas9 to

FokI nuclease improves the specificity of genome modification. Nat. Biotechnol. 32,

577–582 (2014).

194. Tsai, S. Q., Wyvekens, N., Khayter, C., Foden, J. A., Thapar, V., Reyon, D., Goodwin,

M. J., Aryee, M. J. & Joung, J. K. Dimeric CRISPR RNA-guided FokI nucleases for

highly specific genome editing. Nat. Biotechnol. 32, 569–576 (2014).

- 89 -

195. Mali, P., Aach, J., Stranges, P. B., Esvelt, K. M., Moosburner, M., Kosuri, S., Yang, L. &

Church, G. M. Cas9 transcriptional activators for target specificity screening and paired

nickases for cooperative genome engineering. Nat. Biotechnol. 31, 833–838 (2013).

196. Ran, F. A., Hsu, P. D., Lin, C. Y., Gootenberg, J. S., Konermann, S., Trevino, A. E.,

Scott, D. A., Inoue, A., Matoba, S., Zhang, Y. & Zhang, F. Double nicking by RNA-

guided CRISPR-Cas9 for enhanced genome editing specificity. Cell 154, 1380–1389

(2013).

197. Slaymaker, I. M., Gao, L., Zetsche, B., Scott, D. A., Yan, W. X. & Zhang, F. Rationally

engineered Cas9 nucleases with improved specificity. Science 351, 84–88 (2016).

198. Kleinstiver, B. P., Pattanayak, V., Prew, M. S., Tsai, S. Q., Nguyen, N. T., Zheng, Z. &

Joung, J. K. High-fidelity CRISPR-Cas9 nucleases with no detectable genome-wide off-

target effects. Nature 529, 490–495 (2016).

199. Davis, K. M., Pattanayak, V., Thompson, D. B., Zuris, J. A. & Liu, D. R. Small

molecule-triggered Cas9 protein with improved genome-editing specificity. Nat. Chem.

Biol. 11, 316–318 (2015).

200. Zetsche, B., Volz, S. E. & Zhang, F. A split-Cas9 architecture for inducible genome

editing and transcription modulation. Nat. Biotechnol. 33, 139–142 (2015).

201. Nihongaki, Y., Kawano, F., Nakajima, T. & Sato, M. Photoactivatable CRISPR-Cas9 for

optogenetic genome editing. Nat. Biotechnol. 33, 755–760 (2015).

202. Komor, A. C., Kim, Y. B., Packer, M. S., Zuris, J. A. & Liu, D. R. Programmable editing

of a target base in genomic DNA without double-stranded DNA cleavage. Nature 533,

420–424 (2016).

203. Komor, A. C., Zhao, K. T., Packer, M. S., Gaudelli, N. M., Waterbury, A. L., Koblan, L.

W., Kim, Y. B., Badran, A. H. & Liu, D. R. Improved base excision repair inhibition and

bacteriophage Mu Gam protein yields C:G-to-T:A base editors with higher efficiency and

product purity. Sci. Adv. 3, eaao4774 (2017).

- 90 -

204. Kim, K., Ryu, S. M., Kim, S. T., Baek, G., Kim, D., Lim, K., Chung, E., Kim, S. & Kim,

J. S. Highly efficient RNA-guided base editing in mouse embryos. Nat. Biotechnol. 35,

435–437 (2017).

205. Zong, Y., Wang, Y., Li, C., Zhang, R., Chen, K., Ran, Y., Qiu, J. L., Wang, D. & Gao, C.

Precise base editing in rice, wheat and maize with a Cas9-cytidine deaminase fusion. Nat.

Biotechnol. 35, 438–440 (2017).

206. Shimatani, Z., Kashojiya, S., Takayama, M., Terada, R., Arazoe, T., Ishii, H., Teramura,

H., Yamamoto, T., Komatsu, H., Miura, K., Ezura, H., Nishida, K., Ariizumi, T. &

Kondo, A. Targeted base editing in rice and tomato using a CRISPR-Cas9 cytidine

deaminase fusion. Nat. Biotechnol. 35, 441–443 (2017).

207. Fu, Y., Sander, J. D., Reyon, D., Cascio, V. M. & Joung, J. K. Improving CRISPR-Cas

nuclease specificity using truncated guide RNAs. Nat. Biotechnol. 32, 279–284 (2014).

208. Tsai, S. Q., Zheng, Z., Nguyen, N. T., Liebers, M., Topkar, V. V., Thapar, V., Wyvekens,

N., Khayter, C., Iafrate, A. J., Le, L. P., Aryee, M. J. & Joung, J. K. GUIDE-seq enables

genome-wide profiling of off-target cleavage by CRISPR-Cas nucleases. Nat. Biotechnol.

33, 187–198 (2015).

209. Hendel, A., Bak, R. O., Clark, J. T., Kennedy, A. B., Ryan, D. E., Roy, S., Steinfeld, I.,

Lunstad, B. D., Kaiser, R. J., Wilkens, A. B., Bacchetta, R., Tsalenko, A., Dellinger, D.,

Bruhn, L. & Porteus, M. H. Chemically modified guide RNAs enhance CRISPR-Cas

genome editing in human primary cells. Nat. Biotechnol. 33, 985–989 (2015).

210. Rahdar, M., McMahon, M. A., Prakash, T. P., Swayze, E. E., Bennett, C. F. & Cleveland,

D. W. Synthetic CRISPR RNA-Cas9–guided genome editing in human cells. Proc. Natl.

Acad. Sci. 112, e7110–7 (2015)

211. Shalem, O., Sanjana, N. E., Hartenian, E., Shi, X., Scott, D. A., Mikkelsen, T. S., Heckl,

D., Ebert, B. L., Root, D. E., Doench, J. G. & Zhang, F. Genome-scale CRISPR-Cas9

knockout screening in human cells. Science 343, 84–87 (2014).

212. Wang, T., Wei, J. J., Sabatini, D. M. & Lander, E. S. Genetic screens in human cells

- 91 -

using the CRISPR-Cas9 system. Science 343, 80–4 (2014).

213. Koike-Yusa, H., Li, Y., Tan, E. P., Velasco-Herrera, M. D. C. & Yusa, K. Genome-wide

recessive genetic screening in mammalian cells with a lentiviral CRISPR-guide RNA

library. Nat. Biotechnol. 32, 267–273 (2014).

214. Zhou, Y., Zhu, S., Cai, C., Yuan, P., Li, C., Huang, Y. & Wei, W. High-throughput

screening of a CRISPR/Cas9 library for functional genomics in human cells. Nature 509,

487–491 (2014).

215. Liu, S. J. et al. CRISPRi-based genome-scale identification of functional long noncoding

RNA loci in human cells. Science 355, (2017).

216. Montalbano, A., Canver, M. C. & Sanjana, N. E. High-throughput approaches to pinpoint

function within the noncoding genome. Mol. Cell 68, 44–59 (2017).

217. Ma, Y., Zhang, J., Yin, W., Zhang, Z., Song, Y. & Chang, X. Targeted AID-mediated

mutagenesis (TAM) enables efficient genomic diversification in mammalian cells. Nat.

Methods 13, 1029–1035 (2016).

218. Hess, G. T., Frésard, L., Han, K., Lee, C. H., Li, A., Cimprich, K. A., Montgomery, S. B.

& Bassik, M. C. Directed evolution using dCas9-targeted somatic hypermutation in

mammalian cells. Nat. Methods 13, 1036–1042 (2016).

219. Jackson, A. L., Bartz, S. R., Schelter, J., Kobayashi, S. V., Burchard, J., Mao, M., Li, B.,

Cavet, G. & Linsley, P. S. Expression profiling reveals off-target gene regulation by

RNAi. Nat. Biotechnol. 21, 635–637 (2003).

220. Birmingham, A., Anderson, E. M., Reynolds, A., Ilsley-Tyree, D., Leake, D., Fedorov,

Y., Baskerville, S., Maksimova, E., Robinson, K., Karpilow, J., Marshall, W. S. &

Khvorova, A. 3’ UTR seed matches, but not overall identity, are associated with RNAi

off-targets. Nat. Methods 3, 199–204 (2006).

221. Jackson, A. L. & Linsley, P. S. Recognizing and avoiding siRNA off-target effects for

target identification and therapeutic application. Nat. Rev. Drug Discov. 9, 57–67 (2010).

- 92 -

222. Batista, P. J. et al. M6A RNA modification controls cell fate transition in mammalian

embryonic stem cells. Cell Stem Cell 15, 707–719 (2014).

223. Wang, Q., Zou, Y., Nowotschin, S., Kim, S. Y., Li, Q. V., Soh, C. L., Su, J., Zhang, C.,

Shu, W., Xi, Q., Huangfu, D., Hadjantonakis, A. K. & Massagué, J. The p53 family

coordinates wnt and nodal inputs in mesendodermal differentiation of embryonic stem

cells. Cell Stem Cell 20, 70–86 (2017).

224. Hunt, C. P. J., Pouton, C. W. & Haynes, J. M. Characterising the developmental profile

of human embryonic stem cell-derived medium spiny neuron progenitors and assessing

mature neuron function using a CRISPR-generated human DARPP-32WT/eGFP-

AMPreporter line. Neurochem. Int. 106, 3–13 (2017).

225. Adkar, S. S., Willard, V. P., Brunger, J. M., Shiao, K. T., Gersbach, C. A. & Guilak, F.

318. Targeted genome editing of human induced pluripotent stem cells using

CRISPR/Cas9 to generate a knock-in type II collagen reporter for the purification of

chondrogenic cells. Mol. Ther. 24, S128 (2016).

226. Kearns, N. A, Genga, R. M. J., Enuameh, M. S., Garber, M., Wolfe, S. A & Maehr, R.

Cas9 effector-mediated regulation of transcription and differentiation in human

pluripotent stem cells. Development 141, 219–23 (2014).

227. Guo, J., Ma, D., Huang, R., Ming, J., Ye, M., Kee, K., Xie, Z. & Na, J. An inducible

CRISPR-ON system for controllable gene activation in human pluripotent stem cells.

Protein Cell 8, 379–393 (2017).

228. Luo, Y., Xu, X., An, X., Sun, X., Wang, S. & Zhu, D. Targeted inhibition of the MIR-

199a/214 cluster by CRISPR interference augments the tumor tropism of human induced

pluripotent stem cell-derived neural stem cells under hypoxic condition. Stem Cells Int.

2016, 3598542 (2016).

229. Mandegar, M. A. et al. CRISPR Interference Efficiently Induces Specific and Reversible

Gene Silencing in Human iPSCs. Cell Stem Cell 18, 541–553 (2016).

230. Zhang, W. et al. A Werner syndrome stem cell model unveils heterochromatin alterations

- 93 -

as a driver of human aging. Science 348, 1160–1163 (2015).

231. Avior, Y., Lezmi, E., Yanuka, D. & Benvenisty, N. Modeling developmental and

tumorigenic aspects of trilateral retinoblastoma via human embryonic stem cells. Stem

Cell Reports 8, 1354–1365 (2017).

232. Chang, C. J., Kotini, A., Teruya-Feldstein, J., Abdel-Wahab, O., Bradley, R. &

Papapetrou, E. P. Isogenic iPSC models of SRSF2-mutant myelodysplastic syndrome

capture disease phenotypes, splicing defects and drug responses. Blood 128, 962 (2016).

233. Guo, D., Liu, H., Gao, G., Liu, Y., Zhuang, Y., Yang, F., Wang, K., Zhou, T., Qin, D.,

Hong, L., Li, J., Xu, K. & Li, Y.X. Creating a patient carried Men1 gene point mutation

on wild type iPSCs locus mediated by CRISPR/Cas9 and ssODN. Stem Cell Res. 18, 67–

69 (2017).

234. Freedman, B. S. et al. Modelling kidney disease with CRISPR-mutant kidney organoids

derived from human pluripotent epiblast spheroids. Nat. Commun. 6, 8715 (2015).

235. Woo, D. H., Chen, Q., Yang, T. L. B., Glineburg, M. R., Hoge, C., Leu, N. A., Johnson,

F. B. & Lengner, C. J. Enhancing a wnt-telomere feedback loop restores intestinal stem

cell function in a human organotypic model of dyskeratosis congenita. Cell Stem Cell 19,

397–405 (2016).

236. Matano, M., Date, S., Shimokawa, M., Takano, A., Fujii, M., Ohta, Y., Watanabe, T.,

Kanai, T. & Sato, T. Modeling colorectal cancer using CRISPR-Cas9-mediated

engineering of human intestinal organoids. Nat. Med. 21, 256–262 (2015).

237. Drost, J. et al. Sequential cancer mutations in cultured human intestinal stem cells.

Nature 521, 43–47 (2015).

238. Fessler, E. et al. TGFβ signaling directs serrated adenomas to the mesenchymal colorectal

cancer subtype. EMBO Mol. Med. 8, 745–760 (2016).

239. Verissimo, C. S., Overmeer, R. M., Ponsioen, B., Drost, J., Mertens, S., Verlaan-Klink, I.,

Van Gerwen, B., Van Der Ven, M., Van De Wetering, M., Egan, D. A., Bernards, R.,

- 94 -

Clevers, H., Bos, J. L. & Snippert, H. J. Targeting mutant RAS in patient-derived

colorectal cancer organoids by combinatorial drug screening. Elife 5, e18489 (2016).

240. Kemaladewi, D. U. et al. Correction of a splicing defect in a mouse model of congenital

muscular dystrophy type 1A using a homology-directed-repair-independent mechanism.

Nat. Med. 23, 984–989 (2017).

241. Cohen, J. C., Boerwinkle, E., Mosley, T. H. & Hobbs, H. H. Sequence variations in

PCSK9, low LDL, and protection against coronary heart disease. N. Engl. J. Med. 354,

1264–1272 (2006).

242. Ding, Q., Strong, A., Patel, K. M., Ng, S. L., Gosis, B. S., Regan, S. N., Cowan, C. A.,

Rader, D. J. & Musunuru, K. Permanent alteration of PCSK9 with in vivo CRISPR-Cas9

genome editing. Circ. Res. 115, 488–492 (2014).

243. Ran, F. A., Cong, L., Yan, W. X., Scott, D. A., Gootenberg, J. S., Kriz, A. J., Zetsche, B.,

Shalem, O., Wu, X., Makarova, K. S., Koonin, E. V., Sharp, P. A. & Zhang, F. In vivo

genome editing using Staphylococcus aureus Cas9. Nature 520, 186–191 (2015).

244. Wang, X., Raghavan, A., Chen, T., Qiao, L., Zhang, Y., Ding, Q. & Musunuru, K.

CRISPR-Cas9 targeting of PCSK9 in human hepatocytes in vivo - Brief report.

Arterioscler. Thromb. Vasc. Biol. 36, 783–786 (2016).

245. Kalos, M., Levine, B. L., Porter, D. L., Katz, S., Grupp, S. A., Bagg, A. & June, C. H. T

cells with chimeric antigen receptors have potent antitumor effects and can establish

memory in patients with advanced leukemia. Sci. Transl. Med. 3, 95ra73 (2011).

246. Rupp, L. J., Schumann, K., Roybal, K. T., Gate, R. E., Ye, C. J., Lim, W. A. & Marson,

A. CRISPR/Cas9-mediated PD-1 disruption enhances anti-tumor efficacy of human

chimeric antigen receptor T cells. Sci. Rep. 7, 737 (2017).

247. Eyquem, J., Mansilla-Soto, J., Giavridis, T., Van Der Stegen, S. J. C., Hamieh, M.,

Cunanan, K. M., Odak, A., Gönen, M. & Sadelain, M. Targeting a CAR to the TRAC

locus with CRISPR/Cas9 enhances tumour rejection. Nature 543, 113–117 (2017).

- 95 -

248. Blum, R., Arumugam, A., Wu, J., Walcheck, B. & Kaufman, D. Engineering human

pluripotent stem cell-derived natural killer cells to prevent CD16a shedding for enhanced

anti-tumor killing. Blood 128, 1336 (2016).

249. Su, S. et al. CRISPR-Cas9 mediated efficient PD-1 disruption on human primary T cells

from cancer patients. Sci. Rep. 6, 20070 (2016).

250. Su, S., Zou, Z., Chen, F., Ding, N., Du, J., Shao, J., Li, L., Fu, Y., Hu, B., Yang, Y., Sha,

H., Meng, F., Wei, J., Huang, X. & Liu, B. CRISPR-Cas9-mediated disruption of PD-1

on human T cells for adoptive cellular therapies of EBV positive gastric cancer.

Oncoimmunology 6, e1249558 (2017).

251. Mandal, P. K. et al. Efficient ablation of genes in human hematopoietic stem and effector

cells using CRISPR/Cas9. Cell Stem Cell 15, 643–52 (2014).

252. Gundry, M. C., Brunetti, L., Lin, A., Mayle, A. E., Kitano, A., Wagner, D., Hsu, J. I.,

Hoegenauer, K. A., Rooney, C. M., Goodell, M. A. & Nakada, D. Highly efficient

genome editing of murine and human hematopoietic progenitor cells by CRISPR/Cas9.

Cell Rep. 17, 1453–1461 (2016).

253. De Ravin, S. S. et al. CRISPR-Cas9 gene repair of hematopoietic stem cells from patients

with X-linked chronic granulomatous disease. Sci. Transl. Med. 9, (2017).

254. Genovese, P. et al. Targeted genome editing in human repopulating haematopoietic stem

cells. Nature 510, 235–240 (2014).

255. Li, C., Guan, X., Du, T., Jin, W., Wu, B., Liu, Y., Wang, P., Hu, B., Griffin, G. E.,

Shattock, R. J. & Hu, Q. Inhibition of HIV-1 infection of primary CD4+ T-cells by gene

editing of CCR5 using adenovirus-delivered CRISPR/Cas9. J. Gen. Virol. 96, 2381–2393

(2015).

256. Hou, P., Chen, S., Wang, S., Yu, X., Chen, Y., Jiang, M., Zhuang, K., Ho, W., Hou, W.,

Huang, J. & Guo, D. Genome editing of CXCR4 by CRISPR/cas9 confers cells resistant

to HIV-1 infection. Sci. Rep. 5, 15577 (2015).

- 96 -

257. Kang, H. J., Minder, P., Park, M. A., Mesquitta, W. T., Torbett, B. E. & Slukvin, I. I.

CCR5 disruption in induced pluripotent stem cells using CRISPR/Cas9 provides selective

resistance of immune cells to CCR5-tropic HIV-1 virus. Mol. Ther. - Nucleic Acids 4,

e268 (2015).

258. Xu, L. et al. CRISPR/Cas9-Mediated CCR5 Ablation in Human Hematopoietic

Stem/Progenitor Cells Confers HIV-1 Resistance In Vivo. Mol. Ther. 25, 1782–1789

(2017).

259. Firth, A. L., Menon, T., Parker, G. S., Qualls, S. J., Lewis, B. M., Ke, E., Dargitz, C. T.,

Wright, R., Khanna, A., Gage, F. H. & Verma, I. M. Functional gene correction for cystic

fibrosis in lung epithelial cells generated from patient iPSCs. Cell Rep. 12, 1385–1390

(2015).

260. Schwank, G., Koo, B. K., Sasselli, V., Dekkers, J. F., Heo, I., Demircan, T., Sasaki, N.,

Boymans, S., Cuppen, E., Van Der Ent, C. K., Nieuwenhuis, E. E. S., Beekman, J. M. &

Clevers, H. Functional repair of CFTR by CRISPR/Cas9 in intestinal stem cell organoids

of cystic fibrosis patients. Cell Stem Cell 13, 653–658 (2013).

261. Mehta, A. & Haber, J. E. Sources of DNA double-strand breaks and models of

recombinational DNA repair. Cold Spring Harb. Perspect. Biol. 6, a016428 (2014).

262. Bétermier, M., Bertrand, P. & Lopez, B. S. Is Non-Homologous End-Joining Really an

Inherently Error-Prone Process? PLoS Genetics 10, a016428 (2014).

263. Rogakou, E. P., Pilch, D. R., Orr, A. H., Ivanova, V. S. & Bonner, W. M. DNA double-

stranded breaks induce histone H2AX phosphorylation on serine 139. J. Biol. Chem. 273,

5858–5868 (1998).

264. Rogakou, E. P., Boon, C., Redon, C. & Bonner, W. M. Megabase chromatin domains

involved in DNA double-strand breaks in vivo. J. Cell Biol. 146, 905–16 (1999).

265. Paull, T. T., Rogakou, E. P., Yamazaki, V., Kirchgessner, C. U., Gellert, M. & Bonner,

W. M. A critical role for histone H2AX in recruitment of repair factors to nuclear foci

after DNA damage. Curr. Biol. 10, 886–895 (2000).

- 97 -

266. McKinnon, P. J. ATM and ataxia telangiectasia. EMBO Rep. 5, 772–776 (2004).

267. Mimori, T. & Hardin, J. A. Mechanism of interaction between Ku protein and DNA. J.

Biol. Chem. 261, 10375–10379 (1986).

268. Blier, P. R., Griffith, A. J., Craft, J. & Hardin, A. Binding of Ku Protein to DNA. J. Biol.

Chem. 268, 7594–7601 (1993).

269. Makharashvili, N., Tubbs, A. T., Yang, S. H., Wang, H., Barton, O., Zhou, Y.,

Deshpande, R. A., Lee, J. H., Lobrich, M., Sleckman, B. P., Wu, X. & Paull, T. T.

Catalytic and Noncatalytic Roles of the CtIP Endonuclease in Double-Strand Break End

Resection. Mol. Cell 54, 1022–1033 (2014).

270. Masani, S., Han, L., Meek, K. & Yu, K. Redundant function of DNA ligase 1 and 3 in

alternative end-joining during immunoglobulin class switch recombination. Proc. Natl.

Acad. Sci. U. S. A. 113, 1–6 (2016).

271. Chang, H. H. Y., Pannunzio, N. R., Adachi, N. & Lieber, M. R. Non-homologous DNA

end joining and alternative pathways to double-strand break repair. Nat. Rev. Mol. Cell

Biol. 18, 495–506 (2017).

272. Yoo, S. & Dynan, W. S. Geometry of a complex formed by double strand break repair

proteins at a single DNA end: recruitment of DNA-PKcs induces inward translocation of

Ku protein. Nucleic Acids Res. 27, 4679–4686 (1999).

273. Hammarsten, O., DeFazio, L. G. & Chu, G. Activation of DNA-dependent protein kinase

by single-stranded DNA ends. J. Biol. Chem. 275, 1541–1550 (2000).

274. Ma, Y., Pannicke, U., Schwarz, K. & Lieber, M. R. Hairpin opening and overhang

processing by an Artemis/DNA-dependent protein kinase complex in nonhomologous

end joining and V(D)J recombination. Cell 108, 781–794 (2002).

275. Ramadan, K., Maga, G., Shevelev, I. V., Villani, G., Blanco, L. & Hübscher, U. Human

DNA polymerase λ possesses terminal deoxyribonucleotidyl transferase activity and can

elongate RNA primers: Implications for novel functions. J. Mol. Biol. 328, 63–72 (2003).

- 98 -

276. Ramadan, K., Shevelev, I. V, Maga, G. & Hübscher, U. De novo DNA synthesis by

human DNA polymerase lambda, DNA polymerase mu and terminal

deoxyribonucleotidyl transferase. J. Mol. Biol. 339, 395–404 (2004).

277. Gu, J., Lu, H., Tippin, B., Shimazaki, N., Goodman, M. F. & Lieber, M. R. XRCC4:DNA

ligase IV can ligate incompatible DNA ends and can ligate across gaps. EMBO J. 26,

1010–1023 (2007).

278. Lieber, M. R. The Mechanism of Double-Strand DNA Break Repair by the

Nonhomologous DNA End-Joining Pathway. Annu. Rev. Biochem. 79, 181–211 (2010).

279. Stewart, G. S., Wang, B., Bignell, C. R., Taylor, A. M. R. & Elledge, S. J. MDC1 is a

mediator of the mammalian DNA damage checkpoint. Nature 421, 961–966 (2003).

280. Kobayashi, J., Tauchi, H., Sakamoto, S., Nakamura, A., Morishima, K.I, Matsuura, S.,

Kobayashi, T., Tamai, K., Tanimoto, K. & Komatsu, K. NBS1 localizes to γ-H2AX foci

through interaction with the FHA/BRCT domain. Curr. Biol. 12, 1846–1851 (2002).

281. Sartori, A. A., Lukas, C., Coates, J., Mistrik, M., Fu, S., Bartek, J., Baer, R., Lukas, J. &

Jackson, S. P. Human CtIP promotes DNA end resection. Nature 450, 509–514 (2007).

282. Chen, L., Nievera, C. J., Lee, A. Y.-L. & Wu, X. Cell cycle-dependent complex

formation of BRCA1.CtIP.MRN is important for DNA double-strand break repair. J.

Biol. Chem. 283, 7713–7720 (2008).

283. Wang, H., Shi, L. Z., Wong, C. C. L., Han, X., Hwang, P. Y. H., Truong, L. N., Zhu, Q.,

Shao, Z., Chen, D. J., Berns, M. W., Yates, J. R., Chen, L. & Wu, X. The Interaction of

CtIP and Nbs1 Connects CDK and ATM to Regulate HR-Mediated Double-Strand Break

Repair. PLoS Genet. 9, e1003277 (2013).

284. Garcia, V., Phelps, S. E. L., Gray, S. & Neale, M. J. Bidirectional resection of DNA

double-strand breaks by Mre11 and Exo1. Nature 479, 241–244 (2011).

285. Shibata, A. et al. DNA Double-Strand Break Repair Pathway Choice Is Directed by

Distinct MRE11 Nuclease Activities. Mol. Cell 53, 7–18 (2014).

- 99 -

286. Nimonkar, A. V., Genschel, J., Kinoshita, E., Polaczek, P., Campbell, J. L., Wyman, C.,

Modrich, P. & Kowalczykowski, S. C. BLM-DNA2-RPA-MRN and EXO1-BLM-RPA-

MRN constitute two DNA end resection machineries for human DNA break repair.

Genes Dev. 25, 350–362 (2011).

287. Ashton, N. W., Bolderson, E., Cubeddu, L., O’Byrne, K. J. & Richard, D. J. Human

single-stranded DNA binding proteins are essential for maintaining genomic stability.

BMC Mol. Biol. 14, 9 (2013).

288. Roy, R., Chun, J. & Powell, S. N. BRCA1 and BRCA2: Different roles in a common

pathway of genome protection. Nat. Rev. Cancer 12, 68–78 (2012).

289. Park, J. Y., Zhang, F. & Andreassen, P. R. PALB2: The hub of a network of tumor

suppressors involved in DNA damage responses. Biochim. Biophys. Acta. 1846, 263–275

(2014).

290. Cejka, P., Plank, J. L., Bachrati, C. Z., Hickson, I. D. & Kowalczykowski, S. C. Rmi1

stimulates decatenation of double Holliday junctions during dissolution by Sgs1-Top3.

Nat. Struct. Mol. Biol. 17, 1377–1382 (2010).

291. Wu, L. & Hickson, I. O. The Bloom’s syndrome helicase suppresses crossing over during

homologous recombination. Nature 426, 870–874 (2003).

292. Ira, G., Malkova, A., Liberi, G., Foiani, M. & Haber, J. E. Srs2 and Sgs1-Top3 suppress

crossovers during double-strand break repair in yeast. Cell 115, 401–411 (2003).

293. Wyatt, H. D. M., Sarbajna, S., Matos, J. & West, S. C. Coordinated actions of SLX1-

SLX4 and MUS81-EME1 for holliday junction resolution in human cells. Mol. Cell 52,

234–247 (2013).

294. Verma, P. & Greenberg, R. A. Noncanonical views of homology-directed DNA repair.

Genes Dev. 30, 1138–1154 (2016).

295. Donnianni, R. A. & Symington, L. S. Break-induced replication occurs by conservative

DNA synthesis. Proc. Natl. Acad. Sci. 110, 13475–13480 (2013).

- 100 -

296. Saini, N., Ramakrishnan, S., Elango, R., Ayyar, S., Zhang, Y., Deem, A., Ira, G., Haber,

J. E., Lobachev, K. S. & Malkova, A. Migrating bubble during break-induced replication

drives conservative DNA synthesis. Nature 502, 389–392 (2013).

297. Truong, L. N., Li, Y., Shi, L. Z., Hwang, P. Y.-H., He, J., Wang, H., Razavian, N., Berns,

M. W. & Wu, X. Microhomology-mediated end joining and homologous recombination

share the initial end resection step to repair DNA double-strand breaks in mammalian

cells. Proc. Natl. Acad. Sci. 110, 7720–7725 (2013).

298. Hustedt, N. & Durocher, D. The control of DNA repair by the cell cycle. Nat Cell Biol

19, 1–9 (2017).

299. Symington, L. S. & Gautier, J. Double-strand break end resection and repair pathway

choice. Annu. Rev. Genet. 45, 247–271 (2011).

300. Callen, E. et al. 53BP1 mediates productive and mutagenic DNA repair through distinct

phosphoprotein interactions. Cell 153, 1266–1280 (2013).

301. Chapman, J. R., Barral, P., Vannier, J. B., Borel, V., Steger, M., Tomas-Loba, A., Sartori,

A. A., Adams, I. R., Batista, F. D. & Boulton, S. J. RIF1 Is essential for 53BP1-

dependent nonhomologous end joining and suppression of DNA double-strand break

resection. Mol. Cell 49, 858–871 (2013).

302. Di Virgilio, M., Callen, E., Yamane, A., Zhang, W., Jankovic, M., Gitlin, A. D.,

Feldhahn, N., Resch, W., Oliveira, T. Y., Chait, B. T., Nussenzweig, A., Casellas, R.,

Robbiani, D. F. & Nussenzweig, M. C. Rif1 prevents resection of DNA breaks and

promotes immunoglobulin class switching. Science 339, 711–715 (2013).

303. Zimmermann, M., Lottersberger, F., Buonomo, S. B., Sfeir, A. & De Lange, T. 53BP1

regulates DSB repair using Rif1 to control 5′ end resection. Science 339, 700–704

(2013).

304. Feng, L., Fong, K. W., Wang, J., Wang, W. & Chen, J. RIF1 counteracts BRCA1-

mediated end resection during DNA repair. J. Biol. Chem. 288, 11135–11143 (2013).

- 101 -

305. Escribano-Díaz, C., Orthwein, A., Fradet-Turcotte, A., Xing, M., Young, J. T. F., Tkáč,

J., Cook, M. A., Rosebrock, A. P., Munro, M., Canny, M. D., Xu, D. & Durocher, D. A

cell cycle-dependent regulatory circuit composed of 53BP1-RIF1 and BRCA1-CtIP

controls DNA repair pathway choice. Mol. Cell 49, 872–883 (2013).

306. Zhang, H. et al. A cell cycle-dependent BRCA1-UHRF1 cascade regulates DNA double-

strand break repair pathway choice. Nat. Commun. 7, 10201 (2016).

307. Batenburg, N. L., Walker, J. R., Noordermeer, S. M., Moatti, N., Durocher, D. & Zhu, X.

D. ATM and CDK2 control chromatin remodeler CSB to inhibit RIF1 in DSB repair

pathway choice. Nat. Commun. 8, 1921 (2017).

308. Huertas, P. DNA resection in eukaryotes: Deciding how to fix the break. Nat. Struct. Mol.

Biol. 17, 11–16 (2010).

309. Chapman, J. R., Taylor, M. R. G. & Boulton, S. J. Playing the End Game: DNA double-

strand break repair pathway choice. Mol. Cell 47, 497–510 (2012).

310. Makharashvili, N. & Paull, T. T. CtIP: A DNA damage response protein at the

intersection of DNA metabolism. DNA Repair (Amst). 32, 75–81 (2015).

311. Steger, M., Murina, O., Hühn, D., Ferretti, L. P., Walser, R., Hänggi, K., Lafranchi, L.,

Neugebauer, C., Paliwal, S., Janscak, P., Gerrits, B., Del Sal, G., Zerbe, O. & Sartori, A.

A. Prolyl isomerase PIN1 regulates DNA double-strand break repair by counteracting

DNA end resection. Mol. Cell 50, 333–343 (2013).

312. Lafranchi, L., de Boer, H. R., de Vries, E. G. E., Ong, S.-E., Sartori, A. A. & van Vugt,

M. A. T. M. APC/C(Cdh1) controls CtIP stability during the cell cycle and in response to

DNA damage. EMBO J. 33, 2860–79 (2014).

313. Ferretti, L. P., Himmels, S. F., Trenner, A., Walker, C., Von Aesch, C., Eggenschwiler,

A., Murina, O., Enchev, R. I., Peter, M., Freire, R., Porro, A. & Sartori, A. A. Cullin3-

KLHL15 ubiquitin ligase mediates CtIP protein turnover to fine-tune DNA-end resection.

Nat. Commun. 7, 12628 (2016).

- 102 -

314. Ismail, I. H., Gagné, J. P., Genois, M. M., Strickfaden, H., Mcdonald, D., Xu, Z., Poirier,

G. G., Masson, J. Y. & Hendzel, M. J. The RNF138 E3 ligase displaces Ku to promote

DNA end resection and regulate DNA repair pathway choice. Nat. Cell Biol. 17, 1446–

1457 (2015).

315. Schmidt, C. K., Galanty, Y., Sczaniecka-Clift, M., Coates, J., Jhujh, S., Demir, M.,

Cornwell, M., Beli, P. & Jackson, S. P. Systematic E2 screening reveals a UBE2D-

RNF138-CtIP axis promoting DNA repair. Nat. Cell Biol. 17, 1458–1470 (2015).

316. Yu, X. & Chen, J. DNA damage-induced cell cycle checkpoint control requires CtIP, a

phosphorylation-dependent binding partner of BRCA1 C-terminal domains. Mol. Cell.

Biol. 24, 9478–86 (2004).

317. Huertas, P. & Jackson, S. P. Human CtIP mediates cell cycle control of DNA end

resection and double strand break repair. J. Biol. Chem. 284, 9558–9565 (2009).

318. Yun, M. H. & Hiom, K. CtIP-BRCA1 modulates the choice of DNA double-strand-break

repair pathway throughout the cell cycle. Nature 459, 460–463 (2009).

319. Kaidi, A., Weinert, B. T., Choudhary, C. & Jackson, S. P. Human SIRT6 promotes DNA

end resection through CtIP deacetylation. Science 329, 1348–1353 (2010).

320. Mao, Z., Tian, X., Van Meter, M., Ke, Z., Gorbunova, V. & Seluanov, A. Sirtuin 6

(SIRT6) rescues the decline of homologous recombination repair during replicative

senescence. Proc. Natl. Acad. Sci. 109, 11800–11805 (2012).

321. Tomimatsu, N., Mukherjee, B., Catherine Hardebeck, M., Ilcheva, M., Vanessa

Camacho, C., Louise Harris, J., Porteus, M., Llorente, B., Khanna, K. K. um & Burma, S.

Phosphorylation of EXO1 by CDKs 1 and 2 regulates DNA end resection and repair

pathway choice. Nat. Commun. 5, 3561 (2014).

322. Nimonkar, A. V., Ozsoy, A. Z., Genschel, J., Modrich, P. & Kowalczykowski, S. C.

Human exonuclease 1 and BLM helicase interact to resect DNA and initiate DNA repair.

Proc. Natl. Acad. Sci. 105, 16906–16911 (2008).

- 103 -

323. Chu, V. T., Weber, T., Wefers, B., Wurst, W., Sander, S., Rajewsky, K. & Kühn, R.

Increasing the efficiency of homology-directed repair for CRISPR-Cas9-induced precise

gene editing in mammalian cells. Nat. Biotechnol. 33, 543–548 (2015).

324. Maruyama, T., Dougan, S. K., Truttmann, M. C., Bilate, A. M., Ingram, J. R. & Ploegh,

H. L. Increasing the efficiency of precise genome editing with CRISPR-Cas9 by

inhibition of nonhomologous end joining. Nat. Biotechnol. 33, 538–542 (2015).

325. Pinder, J., Salsman, J. & Dellaire, G. Nuclear domain ‘knock-in’ screen for the evaluation

and identification of small molecule enhancers of CRISPR-based genome editing.

Nucleic Acids Res. 43, 9379–9392 (2015).

326. Gutschner, T., Haemmerle, M., Genovese, G., Draetta, G. F. & Chin, L. Post-translational

regulation of Cas9 during G1 enhances homology-directed repair. Cell Rep. 14, 1555–

1566 (2016).

327. Lee, J. S., Grav, L. M., Pedersen, L. E., Lee, G. M. & Kildegaard, H. F. Accelerated

homology-directed targeted integration of transgenes in Chinese hamster ovary cells via

CRISPR/Cas9 and fluorescent enrichment. Biotechnol. Bioeng. 113, 2518–2523 (2016).

328. Song, J., Yang, D., Xu, J., Zhu, T., Chen, Y. E. & Zhang, J. RS-1 enhances

CRISPR/Cas9- and TALEN-mediated knock-in efficiency. Nat. Commun. 7, 10548

(2016).

329. Yang, D., Scavuzzo, M. A., Chmielowiec, J., Sharp, R., Bajic, A. & Borowiak, M.

Enrichment of G2/M cell cycle phase in human pluripotent stem cells enhances HDR-

mediated gene repair with customizable endonucleases. Sci. Rep. 6, 21264 (2016).

330. Robert, F., Barbeau, M., Éthier, S., Dostie, J. & Pelletier, J. Pharmacological inhibition of

DNA-PK stimulates Cas9-mediated genome editing. Genome Med. 7, (2015).

331. Yu, C., Liu, Y., Ma, T., Liu, K., Xu, S., Zhang, Y., Liu, H., La Russa, M., Xie, M., Ding,

S. & Qi, L. S. Small molecules enhance CRISPR genome editing in pluripotent stem

cells. Cell Stem Cell 16, 142–147 (2015).

- 104 -

332. Lin, S., Staahl, B. T., Alla, R. K. & Doudna, J. A. Enhanced homology-directed human

genome engineering by controlled timing of CRISPR/Cas9 delivery. Elife 3, e04766

(2014).

333. Wang, H., Yang, H., Shivalila, C. S., Dawlaty, M. M., Cheng, A. W., Zhang, F. &

Jaenisch, R. One-step generation of mice carrying mutations in multiple genes by

CRISPR/cas-mediated genome engineering. Cell 153, 910–918 (2013).

334. Yang, H., Wang, H., Shivalila, C. S., Cheng, A. W., Shi, L. & Jaenisch, R. One-step

generation of mice carrying reporter and conditional alleles by CRISPR/Cas-mediated

genome engineering. Cell 154, 1370–9 (2013).

335. Wu, Y., Liang, D., Wang, Y., Bai, M., Tang, W., Bao, S., Yan, Z., Li, D. & Li, J.

Correction of a genetic disease in mouse via use of CRISPR-Cas9. Cell Stem Cell 13,

659–662 (2013).

336. Yoshimi, K., Kaneko, T., Voigt, B. & Mashimo, T. Allele-specific genome editing and

correction of disease-associated phenotypes in rats using the CRISPR-Cas platform. Nat.

Commun. 5, 4240 (2014).

337. Ma, Y., Zhang, X., Shen, B., Lu, Y., Chen, W., Ma, J., Bai, L., Huang, X. & Zhang, L.

Generating rats with conditional alleles using CRISPR/Cas9. Cell Research 24, 122–125

(2014).

338. Yoshimi, K., Kunihiro, Y., Kaneko, T., Nagahora, H., Voigt, B. & Mashimo, T. ssODN-

mediated knock-in with CRISPR-Cas for large genomic regions in zygotes. Nat.

Commun. 7, 10431 (2016).

339. Orthwein, A., Noordermeer, S. M., Wilson, M. D., Landry, S., Enchev, R. I., Sherker, A.,

Munro, M., Pinder, J., Salsman, J., Dellaire, G., Xia, B., Peter, M. & Durocher, D. A

mechanism for the suppression of homologous recombination in G1 cells. Nature 528,

422–426 (2015).

340. Yu, X., Wu, L. C., Bowcock, A. M., Aronheim, A. & Baer, R. The C-terminal (BRCT)

domains of BRCA1 interact in vivo with CtIP, a protein implicated in the CtBP pathway

- 105 -

of transcriptional repression. J. Biol. Chem. 273, 25388–25392 (1998).

341. Tomimatsu, N., Mukherjee, B., Deland, K., Kurimasa, A., Bolderson, E., Khanna, K. K.

& Burma, S. Exo1 plays a major role in DNA end resection in humans and influences

double-strand break repair and damage signaling decisions. DNA Repair (Amst). 11, 441–

448 (2012).

342. Chow, Y. H., O’Brodovich, H., Plumb, J., Wen, Y., Sohn, K. J., Lu, Z., Zhang, F.,

Lukacs, G. L., Tanswell, A. K., Hui, C. C., Buchwald, M. & Hu, J. Development of an

epithelium-specific expression cassette with human DNA regulatory elements for

transgene expression in lung airways. Proc. Natl. Acad. Sci. U. S. A. 94, 14695–700

(1997).

343. Kim, J. H., Lee, S.-R., Li, L.-H., Park, H.-J., Park, J.-H., Lee, K. Y., Kim, M.-K., Shin, B.

A. & Choi, S.-Y. High cleavage efficiency of a 2A peptide derived from porcine

teschovirus-1 in human cell lines, zebrafish and mice. PLoS One 6, e18556 (2011).

344. Daniels, R. W., Rossano, A. J., Macleod, G. T. & Ganetzky, B. Expression of multiple

transgenes from a single construct using viral 2A peptides in Drosophila. PLoS One 9,

e100637 (2014).

345. Kotin, R. M., Linden, R. M. & Berns, K. I. Characterization of a preferred site on human

chromosome 19q for integration of adeno-associated virus DNA by non-homologous

recombination. EMBO J. 11, 5071–8 (1992).

346. Smith, J. R., Maguire, S., Davis, L. A., Alexander, M., Yang, F., Chandran, S., ffrench-

Constant, C. & Pedersen, R. A. Robust, persistent transgene expression in human

embryonic stem cells is achieved with AAVS1-targeted integration. Stem Cells 26, 496–

504 (2008).

347. Wiles, M. V., Vauti, F., Otte, J., Füchtbauer, E. M., Ruiz, P., Füchtbauer, A., Arnold, H.

H., Lehrach, H., Metz, T., Von Melchner, H. & Wurst, W. Establishment of a gene-trap

sequence tag library to generate mutant mice from embryonic stem cells. Nature Genet.

24, 13–14 (2000).

- 106 -

348. Gerard, C. & Goldbeter, A. Temporal self-organization of the cyclin/Cdk network driving

the mammalian cell cycle. Proc. Natl. Acad. Sci. 106, 21643–21648 (2009).

349. Malumbres, M. Cyclin-dependent kinases. Genome Biol. 15, 122 (2014).

350. Peterson, S. E., Li, Y., Chait, B. T., Gottesman, M. E., Baer, R. & Gautier, J. Cdk1

uncouples CtIP-dependent resection and Rad51 filament formation during M-phase

double-strand break repair. J. Cell Biol. 194, 705–720 (2011).

351. Krajewska, M., Heijink, A. M., Bisselink, Y. J. W. M., Seinstra, R. I., Silljé, H. H. W.,

De Vries, E. G. E. & Van Vugt, M. A. T. M. Forced activation of Cdk1 via wee1

inhibition impairs homologous recombination. Oncogene 32, 3001–3008 (2013).

352. Lim, S. & Kaldis, P. Cdks, cyclins and CKIs: roles beyond cell cycle regulation.

Development 140, 3079–3093 (2013).

353. Davies, O. R., Forment, J. V., Sun, M., Belotserkovskaya, R., Coates, J., Galanty, Y.,

Demir, M., Morton, C. R., Rzechorzek, N. J., Jackson, S. P. & Pellegrini, L. CtIP

tetramer assembly is required for DNA-end resection and repair. Nat. Struct. Mol. Biol.

22, 150–157 (2015).

354. Nakade, S., Tsubota, T., Sakane, Y., Kume, S., Sakamoto, N., Obara, M., Daimon, T.,

Sezutsu, H., Yamamoto, T., Sakuma, T. & Suzuki, K. I. T. Microhomology-mediated

end-joining-dependent integration of donor DNA in cells and animals using TALENs and

CRISPR/Cas9. Nat. Commun. 5, 5560 (2014).

355. He, X., Tan, C., Wang, F., Wang, Y., Zhou, R., Cui, D., You, W., Zhao, H., Ren, J.,

Feng, B. & Dna, C. C. Knock-in of large reporter genes in human cells via CRISPR/

Cas9-induced homology-dependent and independent DNA repair. Nucleic Acids Res. 19,

1–14 (2016).

356. Paix, A., Folkmann, A., Goldman, D. H., Kulaga, H., Grzelak, M. J., Rasoloson, D.,

Paidemarry, S., Green, R., Reed, R. R. & Seydoux, G. Precision genome editing using

synthesis-dependent repair of Cas9-induced DNA breaks. Proc. Natl. Acad. Sci. 114,

e10745–54 (2017).

- 107 -

357. Mearini, G., Nielsen, P. E. & Fackelmayer, F. O. Localization and dynamics of small

circular DNA in live mammalian nuclei. Nucleic Acids Res. 32, 2642–2651 (2004).

358. Everett, R. D. The spatial organization of DNA virus genomes in the nucleus. PLoS

Pathog. 9, e1003386 (2013).

359. Highsmith, W. E., Burch, L. H., Zhou, Z., Olsen, J. C., Strong, T. V., Smith, T.,

Friedman, K. J., Silverman, L. M., Boucher, R. C., Collins, F. S. & Knowles, M. R.

Identification of a splice site mutation (2789 + 5 G>A) associated with small amounts of

normal CFTR mRNA and mild cystic fibrosis. Hum. Mutat. 9, 332–338 (1997).

360. Kerem, E. et al. A cystic fibrosis transmembrane conductance regulator splice variant

with partial penetrance associated with variable cystic fibrosis presentations. Am. J.

Respir. Crit. Care Med. 155, 1914–1920 (1997).

361. Canny, M. D. et al. Inhibition of 53BP1 favors homology-dependent DNA repair and

increases CRISPR-Cas9 genome-editing efficiency. Nat. Biotechnol. 36, 95–102 (2018).

362. Brooker, A. S. & Berkowitz, K. M. The roles of cohesins in mitosis, meiosis, and human

health and disease. Methods Mol. Biol. 1170, 229–266 (2014).

363. Sjogren, C. & Nasmyth, K. Sister chromatid cohesion is required for postreplicative

double-strand break repair in Saccharomyces cerevisiae. Curr. Biol. 11, 991–995 (2001).

364. Ström, L., Karlsson, C., Lindroos, H. B., Wedahl, S., Katou, Y., Shirahige, K. & Sjögren,

C. Postreplicative formation of cohesion is required for repair and induced by a single

DNA break. Science 317, 242–245 (2007).

365. Whelan, G., Kreidl, E., Peters, J. M. & Eichele, G. The non-redundant function of

cohesin acetyltransferase Esco2: some answers and new questions. Nucleus 3, 330–334

(2012).

366. Ünal, E., Arbel-Eden, A., Sattler, U., Shroff, R., Lichten, M., Haber, J. E. & Koshland, D.

DNA damage response pathway uses histone modification to assemble a double-strand

break-specific cohesin domain. Mol. Cell 16, 991–1002 (2004).

- 108 -

367. Ström, L., Lindroos, H. B., Shirahige, K. & Sjögren, C. Postreplicative recruitment of

cohesin to double-strand breaks is required for DNA repair. Mol. Cell 16, 1003–1015

(2004).

368. Ünal, E., Heidinger-Pauli, J. M. & Koshland, D. DNA double-strand breaks trigger

genome-wide sister-chromatid cohesion through Eco1 (Ctf7). Science 317, 245–248

(2007).

369. Lee, C.-S., Wang, R. W., Chang, H.-H., Capurso, D., Segal, M. R. & Haber, J. E.

Chromosome position determines the success of double-strand break repair. Proc. Natl.

Acad. Sci. 113, e146–154 (2016).

370. Kadyk, L. C. & Hartwell, L. H. Sister chromatids are preferred over homologs as

substrates for recombinational repair in Saccharomyces cerevisiae. Genetics 132, 387–

402 (1992).

371. Johnson, R. D. & Jasin, M. Sister chromatid gene conversion is a prominent double-

strand break repair pathway in mammalian cells. EMBO J. 19, 3398–3407 (2000).

372. Johnson, R. D. & Jasin, M. Double-strand-break-induced homologous recombination in

mammalian cells. Biochem. Soc. Trans. 29, 196–201 (2001).

373. Covo, S., Westmoreland, J. W., Gordenin, D. A. & Resnick, M. A. Cohesin is limiting for

the suppression of DNA damage-induced recombination between homologous

chromosomes. PLoS Genet. 6, 1–16 (2010).

374. Lukacs, G. L., Haggie, P., Seksek, O., Lechardeur, D., Freedman, N. & Verkman, A. S.

Size-dependent DNA mobility in cytoplasm and nucleus. J. Biol. Chem. 275, 1625–1629

(2000).

375. Mieruszynski, S., Digman, M. A., Gratton, E. & Jones, M. R. Characterization of

exogenous DNA mobility in live cells through fluctuation correlation spectroscopy. Sci.

Rep. 5, 13848 (2015).

376. Shimizu, N., Kamezaki, F. & Shigematsu, S. Tracking of microinjected DNA in live cells

- 109 -

reveals the intracellular behavior and elimination of extrachromosomal genetic material.

Nucleic Acids Res. 33, 6296–6307 (2005).

377. Gasiorowski, J. Z. & Dean, D. A. Intranuclear trafficking of episomal DNA is

transcription-dependent. Mol. Ther. 15, 2132–2139 (2007).

378. Larkin, J. D., Papantonis, A. & Cook, P. R. Promoter type influences transcriptional

topography by targeting genes to distinct nucleoplasmic sites. J. Cell Sci. 126, 2052–2059

(2013).

379. Aguilera, A. & Gómez-González, B. Genome instability: A mechanistic view of its

causes and consequences. Nat. Rev. Genet. 9, 204–217 (2008).

380. Delacôte, F. & Lopez, B. S. Importance of the cell cycle phase for the choice of the

appropriate DSB repair pathway, for genome stability maintenance: The trans-S double-

strand break repair model. Cell Cycle 7, 33–38 (2008).

381. Kueng, S., Hegemann, B., Peters, B. H., Lipp, J. J., Schleiffer, A., Mechtler, K. & Peters,

J. M. Wapl controls the dynamic association of cohesin with chromatin. Cell 127, 955–

967 (2006).

382. Nishiyama, T., Ladurner, R., Schmitz, J., Kreidl, E., Schleiffer, A., Bhaskara, V., Bando,

M., Shirahige, K., Hyman, A. A., Mechtler, K. & Peters, J. M. Sororin mediates sister

chromatid cohesion by antagonizing Wapl. Cell 143, 737–749 (2010).

383. Cucco, F. & Musio, A. Genome stability: What we have learned from cohesinopathies.

Am. J. Med. Genet. 172, 171–178 (2016).

384. Keppler, A., Gendreizig, S., Gronemeyer, T., Pick, H., Vogel, H. & Johnsson, K. A

general method for the covalent labeling of fusion proteins with small molecules in vivo.

Nat. Biotechnol. 21, 86–89 (2003).

385. Kurakhmaeva, K. B., Djindjikhashvili, I. A., Petrov, V. E., Balabanyan, V. U., Voronina,

T. A., Trofimov, S. S., Kreuter, J., Gelperina, S., Begley, D. & Alyautdin, R. N. Brain

targeting of nerve growth factor using poly(butyl cyanoacrylate) nanoparticles. J. Drug

- 110 -

Target. 17, 564–574 (2009).

386. Hyde, S. C. et al. Repeat administration of DNA/liposomes to the nasal epithelium of

patients with cystic fibrosis. Gene Ther. 7, 1156–1165 (2000).

387. Alton, E. W. F. W. et al. A phase I/IIa safety and efficacy study of nebulized liposome-

mediated gene therapy for cystic fibrosis supports a multidose trial. Am. J. Respir. Crit.

Care Med. 192, 1389–1392 (2015).