Anti-Crisprs and CRISPR-Cas: Characterization and Biotechnology

Anti-CRISPRs and CRISPR-Cas: Characterization and biotechnology

Marios Mejdani

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Department of Biochemistry

University of Toronto

Anti-CRISPRs and CRISPR-Cas: Characterization and biotechnology

Marios Mejdani

Doctor of Philosophy

Department of Biochemistry University of Toronto

2019

Abstract

Bacteria and phages (bacterial specific viruses) have been undergoing an evolutionary arms race for billions of years, whereby bacteria evolve mechanisms to inhibit phage infection and phages evolve mechanisms to evade bacterial defenses. Consequently, there are several mechanisms used by bacteria to inhibit phage infection. These include inhibiting phage adsorption, restriction modification systems, and the more recently discovered CRISPR-Cas immune systems.

Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR) loci together with their accompanying CRISPR-associated (Cas) genes form the only known bacterial adaptive defense mechanism that effectively protects against the transfer of mobile genetic elements (MGEs) such as bacteriophages. CRISPR-Cas systems use an RNA-guided nuclease to bind and cleave foreign

DNA, presenting a powerful barrier to phage infection.

This strong evolutionary barrier led phages to evolve small protein inhibitors of CRISPR-Cas called anti-CRISPRs. In the first section of my work, I characterize the structure, function, and mechanism of action for an anti-CRISPR that inhibits the type I-E CRISPR-Cas system of

Pseudomonas aeruginosa strain 4386. I show that beyond simply inhibiting the CRISPR-Cas system, anti-CRISPR AcrIE2 converts the CRISPR-Cas system from a DNA degradation

ii complex to a transcriptional regulator. This modification suggests that anti-CRISPR proteins may function to do more than simply inhibit CRISPR-Cas targeting.

Although CRISPR-Cas systems are a manifestation of the evolutionary arms race between bacteria and phages, CRISPR-Cas systems have also been used for genome editing in various organisms for research purposes. Considering this previous work, I developed a type I-E and type II-A CRISPR-Cas genome editing system to manipulate the genome of different P. aeruginosa strains for research. In the second section of my work, I discuss the methods I developed to efficiently edit the genome of P. aeruginosa . My work on genome editing in P. aeruginosa has and will allow for the development of new P. aeruginosa mutants for research purposes.

Collectively, my work provides insight into the evolutionary interactions between phages and bacteria in the context of CRISPR-Cas and anti-CRISPRs. Moreover, it provides a genome editing tool for future P. aeruginosa studies.

iii

Acknowledgments

I would like to acknowledge my supervisor, Dr. Alan Davidson, for his excellent support and mentorship over the last five years. Alan has steered me in the right direction both within the lab and outside of it. I have certainly benefited from his advice over the years. I would also like to thank Dr. Karen Maxwell for her expertise in suggesting ideas and helping troubleshoot experiments.

To my supervisory committee composed of Dr. William Navarre and Dr. Angus McQuibban. I want to thank you for your insightful ideas. Will, the ability to walk into your office and just throw ideas at you when Alan wasn’t around was truly invaluable. Angus, your relaxed personality and attention to detail has helped me reach my Ph. D. today.

Now to thank the members of the lab. Thank you everyone for creating a wonderful atmosphere to work in. A special thanks to April Pawluk for training me at the start of my journey. I would also like to thank Joe Bondy-Denomy for his insight and scientific advice in my early years.

Kristina deserves a special thanks in this acknowledgment, thank you for helping me fully develop the Cas9 genome editing tool and more so, thank you for being a friend I could always count on. Also, I would like to thank Kristina for helping me put overnights in the incubator when I needed it. To Vasu, Chidozie, Eric, and Brian, thank you for your help dealing with issues both within and outside of the lab, you are all true friends.

Now for people and friends outside the lab, I want to thank people from Grant Brown’s lab for the help, care, and support they gave me relatively early in my M. Sc. / Ph. D. I would also like to generally acknowledge people in the Moraes lab, the Nodwell lab, the Cowen lab, and the

Ensminger lab.

Finally, and importantly my parents. I have watched them work so hard during their life all so that I could have opportunities not afforded to them. These are true heroes in today’s world and nothing short of it. They have taught me to work hard and work smart, be confident in myself and my abilities, and allow no one to stand in my way. Lessons that I consider to be the most invaluable in the world I see today.

Table of Contents Contents

Acknowledgments ...... iv

Table of Contents ...... vi

List of Tables ...... ix

List of Figures ...... x

List of Abbreviations ...... xii

List of Appendices (if any) ...... xiii

Chapter 1 ...... 1

1.1 Overview ...... 1

1.2 Bacteria and Mobile Genetic Elements (MGEs) ...... 2

1.3 Bacteriophages ...... 4

1.4 Co-evolution of bacteria and phages ...... 7

1.5 CRISPR-Cas systems (Class I and Class II) ...... 8

1.5.1 Type I-E and Type II-A CRISPR-Cas mechanisms ...... 10

1.6 Discovery of CRISPR-Cas inhibitors ...... 13

1.6.1 Anti-CRISPR mechanisms ...... 14

1.7 CRISPR-Cas: As a biotechnological tool ...... 17

1.8 CRISPR-Cas used for genome editing ...... 17

1.8.1 Type I-E CRISPR-Cas mechanism for biotechnology ...... 20

1.8.2 Type II-A CRISPR-Cas mechanism for biotechnology ...... 21

1.9 Thesis Objectives ...... 21

1.10 Thesis Outline ...... 22

Chapter 2 ...... 23

Anti-CRISPR AcrIE2: A novel mechanism of CRISPR-Cas inhibition ...... 23

2.1 Acknowledgements ...... 23

2.2 Abstract ...... 23

2.3 Anti-CRISPR AcrIE2 ...... 24

2.4 Results ...... 26

2.4.1 AcrIE2 functional residues...... 26

2.4.2 AcrIE2 interacting partners...... 35

2.4.3 AcrIE2 mutants ...... 36

2.4.4 AcrIE2 inhibits DNA degradation but permits DNA binding ...... 38

2.4.5 AcrIE2 is responsible for CRISPR-Cas dependent transcriptional repression and activation ...... 44

2.5 Discussion...... 48

2.6 Materials and Methods ...... 52

2.6.1 Phage propagation and bacterial growth ...... 52

2.6.2 Phage spotting assays ...... 53

2.6.3 Lysogen formation ...... 53

2.6.4 Site-directed mutagenesis and SOE-PCR ...... 54

2.6.5 AcrIE2 expression in E. coli and protein purification ...... 55

2.6.6 Circular dichroism spectroscopy ...... 55

2.6.7 P. aeruginosa AcrIE2 affinity purification ...... 56

2.6.8 Silver staining ...... 56

2.6.9 Transcriptional reporter assay ...... 57

2.6.10 Twitching motility assays ...... 58

2.6.11 RT-qPCR experiments ...... 59

Chapter 3 ...... 60

CRISPR-Cas genome editing in P. aeruginosa ...... 60

3.1 Acknowledgments ...... 60

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3.2 Abstract ...... 60

3.3 Results ...... 61

3.3.1 Type I-E CRISPR-Cas genome editing methods for P. aeruginosa ...... 61

3.4 P. aeruginosa tCas9 plasmid ...... 66

3.4.1 HR independent Cas9 genome editing method for P. aeruginosa ...... 67

3.4.2 P. aeruginosa Cas9 genome editing with homologous recombination ...... 71

3.5 Discussion...... 76

3.6 Materials and Methods ...... 78

3.6.1 P. aeruginosa competence and transformation ...... 78

3.6.2 Type I-E crRNAs and type II-A sgRNAs used in this work ...... 79

3.6.3 SOE-PCR protocol ...... 80

3.6.4 Additional protocols...... 80

Chapter 4 ...... 81

General Discussion ...... 81

4.1 Summary ...... 81

4.2 Future directions ...... 83

4.2.1 Anti-CRISPR characterization and biotechnological tools...... 83

4.2.2 P. aeruginosa genome editing ...... 84

4.3 Closing Remarks ...... 85

References or Bibliography (if any) ...... 86

Appendices (if any) ...... 96

viii

List of Tables

Table 1: Type I-F and type I-E CRISPR-Cas sequence comparison ...... 10

Table 2: Point mutations of AcrIE2 ...... 26-27

Table 3: Mutagenesis of multiple AcrIE2 residues ...... 28-29

Table 4: Mass spectrometry results summary ...... 36

Table 5: P. aeruginosa Cas9 genome editing ...... 76-77

List of Figures

Figure 1: Bacterial horizontal gene transfer ...... 3

Figure 2: Caudovirales phages ...... 5

Figure 3: Lytic and lysogenic cycles ...... 6

Figure 4: General representation of CRISPR-Cas function ...... 9

Figure 5: Type I-E and type I-F CRISPR-Cas targeting mechanism ...... 11

Figure 6: Cas9 recognition and targeting ...... 13

Figure 7: Mechanisms of CRISPR-Cas inhibition ...... 16

Figure 8: CRISPR-Cas mechanisms for genome editing ...... 19

Figure 9: NMR structure of AcrIE2...... 25

Figure 10: Point mutations of AcrIE2 ...... 26

Figure 11: Functional residues of AcrIE2 ...... 30

Figure 12: Protein expression of AcrIE1, AcrIE2, and AcrIE2 inactive mutants from P. aeruginosa ...... 31

Figure 13: E. coli purification and FPLC of AcrIE2 and AcrIE2 mutants for circular dichroism ...... 33

Figure 14: Circular dichroism analysis of AcrIE2 mutants...... 34

Figure 15: AcrIE2 affinity purification and interactors ...... 37

Figure 16: CRISPR-Cas dependent transcriptional repression of phzM ...... 38-40

Figure 17: AcrIE2 dependent transcriptional regulation of phzM ...... 41-42

Figure 18: CRISPR-Cas dependent transcriptional repression of pilA ...... 43-44

Figure 19: AcrIE2 dependent transcriptional regulation of pilA ...... 45-46

Figure 20: AcrIE2 model of activity ...... 50

Figure 21: HR dependent type I-E CRISPR-Cas genome editing ...... 61

Figure 22: P. aeruginosa 4386 knockout strains ...... 64-65

Figure 23: Plasmid map of tCas9 ...... 66

Figure 24: tCas9 + sgRNA transformation efficiency in SMC 4386 ...... 68

Figure 25: HR independent tCas9 mutation ...... 69

Figure 26: HR independent tCas9 knockout ...... 70

Figure 27: HR dependent type II-A CRISPR-Cas genome editing ...... 71

Figure 28: tCas9 + sgRNA transformation efficiency in PAO1 ...... 73

Figure 29: HR dependent type II-A CRISPR-Cas production of JBD68 gp24 Δ1-150 phage ...... 74-75

List of Abbreviations

6xHis six histidine tag

Acr anti-CRISPR

BLASTp basic local alignment search tool – protein

Bp base pair

PCR Polymerase chain reaction

CRISPR Clustered regularly interspaced short palindromic repeats

Cas CRISPR-associated

CASCADE CRISPR-associated complex for anti-viral defense crRNA CRISPR RNA sgRNA Synthetic guide ribonucleic acid

DNA Deoxyribonucleic acid

NHEJ Non-homologous end joining

RGD Random genetic drift

HGT Horizontal gene transfer

IPTG Isopropyl β-D-1-thiogalactopyranoside

LB Lysogeny broth

HR Homologous recombination

xii

List of Appendices (if any)

xiii

Chapter 1

Introduction

1.1 Overview

Bacteria evolve through random genetic drift (RGD) and horizontal gene transfer (HGT). HGT is a mechanism of lateral gene transfer that allows bacteria to acquire and integrate foreign DNA into their genome. Phages (bacterial specific viruses) are a major contributor to HGT as they can effectively bind bacterial cells and inject DNA into the cytoplasm. This vector-based mechanism of HGT comes with the significant caveat of potential lysis and death for the bacterial host.

To defend against phage infection and other lateral gene transfer elements bacteria have evolved mechanisms to degrade foreign DNA sequences. One of these mechanisms is CRISPR-Cas or

Clustered Regularly Interspaced Short Palindromic Repeats and CRISPR associated genes.

Although there are a variety of different CRISPR-Cas systems they all use an RNA guided protein complex that can anneal to a foreign DNA and/or RNA sequence and through nuclease activity induce degradation. As expected, through this mechanism, CRISPR-Cas poses a significant threat to phage infection.

To circumvent CRISPR-Cas systems bacteriophages code for CRISPR-Cas inhibitors called anti-

CRISPRs (Acrs). These Acrs are small phage encoded proteins that can interact and inhibit

CRISPR-Cas nuclease activity, thereby enabling lateral gene transfer. The first focus of this thesis is to characterize in detail the structure and inhibitory function of AcrIE2, an Acr that can inhibit the Type I-E CRISPR-Cas system of P. aeruginosa .

Although CRISPR-Cas systems are a manifestation of the co-evolution between bacteria and phages, CRISPR-Cas systems have also been used for biotechnological purposes. Through the

RNA guided nuclease activity of CRISPR-Cas systems, bacterial DNA sequences can be mutated, resulting in new bacterial strains that can be used for research. In the second section of this thesis, I focus on my work in creating a type I-E and type II-A CRISPR-Cas genome editing tool for P. aeruginosa .

1.2 Bacteria and Mobile Genetic Elements (MGEs)

Two of the main modes for bacterial evolution are RGD and HGT. RGD considers the random mutations that naturally occur in a replicating and dividing bacterial population, these include

DNA polymerase mistakes during division (ex: insertions, deletions etc.), as well as duplications

(Kimura and Maruyama 1969; Drake 1991; Rosche and Foster 2000; Lynch and Force 2000;

Lynch and Conery 2000). HGT focuses on the transfer of genetic material or mobile genetic elements (MGEs) between bacterial organisms by a mechanism other than reproduction (Figure

1). The mechanisms of HGT include natural transformation, conjugation, and transduction

(Doolittle 1999; Garcia-Vallvé et al. 2000; Daubin and Szöllősi 2016; Griffith 1928; Jain et al.

1999).

Figure 1: Bacterial horizontal gene transfer. Representation of the mechanisms for horizontal gene transfer. Upon DNA acquisition the bacterial cell recombines genetic material into the chromosome. This figure was modified from Servier Medical Art, licensed under a Creative Common Attribution 3.0 Generic License. http://smart.servier.com/.

In natural transformation, bacteria take up and recombine exogenous DNA into their chromosomes. Usually the acquired DNA comes from other bacteria found in the vicinity that have lysed and spilled their contents (Avery et al. 1944). In conjugation two bacterial cells form a conjugation bridge that allows for the transfer of DNA from the donor bacterium to the recipient (Trieu-Cuot et al. 1987). Unlike natural transformation and conjugation, transduction is the only known form of DNA transfer that requires a vector (Zinder and Lederberg 1952). In transduction, a bacterial specific virus called a bacteriophage must bind the surface of a bacterial cell and inject its DNA content into the cytoplasm (Hershey and Chase 1952). This DNA content includes phage genes required for viral propagation and may also include additional pieces of

DNA from a previous host. In this manner, genetic material is transferred from one bacterium to another via a vector. Collectively, these methods of horizontal genetic transfer have a significant

impact on the evolution of bacterial cells as they are responsible for transferring genetic material into a variety of bacterial hosts.

1.3 Bacteriophages

Bacteriophages (bacterial specific viruses) outnumber their bacterial hosts by a factor of 10:1, making them the most abundant entities on the planet (Clokie et al. 2011). During infection, bacteriophages transfer their genetic material into the host and can cause bacterial lysis or cell death (Ryan and Rutenberg 2007; d’Herelle 1917; Twort 1915). Moreover, during infection, bacteriophages serve as a mode of HGT making them a significant component of bacterial evolution. In fact, whole genome sequencing has shown that phage sequences are a major source of variability between bacterial strains and that a number of bacterial virulence genes are encoded by phages (Schneider 2017). Given the intricate connection between phages and bacterial evolution, phages are responsible for influencing a myriad of processes including environmental nutrient cycling, the human microbiome, and the global food supply (Sime-

Ngando 2014).

96 % of all phages are found in the order Caudovirales and are composed of an icosahedral head that is attached to a flexible tail (Fokine and Rossmann 2014; Ackermann 2007). Caudovirales are divided into three large phylogenetically related families; Myoviridae represent phages with long contractile tails, Siphoviridae represent phages with long non-contractile tails, and

Podoviridae represent phages with short tails (Ackermann 2001) (Figure 2). During infection, the tail of a phage must first encounter and recognize a receptor on the bacterial cell surface.

Upon binding, the phage punctures the cell wall and one or two cell membranes (gram positive

or gram negative respectively) to deliver the genetic material through its tail into the host

(Hyman and Abedon 2010).

Siphoviridae Myoviridae Podoviridae

Figure 2: Caudovirales phages. Structural differences between the three different Caudovirales phages. Parts of this figure were modified from Servier Medical Art, licensed under a Creative Common Attribution 3.0 Generic License. http://smart.servier.com/.

Once the genetic material has entered the bacterial cytoplasm a lytic phage will begin replicating its genome and producing phage particles (Molineux and Panja 2013). These particles will form mature phages and will cause bacterial cell lysis and death (Figure 3). In contrast, if the infection is completed by a temperate phage a decision point is reached determining whether the phage

undergoes a lytic or lysogenic lifecycle (Wang et al. 2000). If the lytic cycle is chosen, the phage will cause cell lysis in a manner similar to its lytic counterpart. If the lysogenic cycle is chosen, the phage genome will integrate within the host and replicate with the host (Figure 3). In this state, the phage inhibits the production of phage structural genes and may express genes that alter the phenotype of the bacterial cell, such as virulence factors. A lysogenic phage can become lytic under certain stresses such as DNA damage and can therefore return to lytic replication. A temperate phage present within the genome of a bacterial cell is called a prophage and bacteria carrying a prophage are referred to as lysogens (Figure 3).

Figure 3: Lytic and lysogenic cycles. The phage initially binds to its surface receptor and injects its DNA into the cytoplasm. If lytic infection occurs, the phage will immediately begin replicating its genome and making phage particles, resulting in eventual bacterial cell lysis. If the lysogenic cycle occurs, the phage will initially integrate into the host genome and replicate with the host. Lysogenic phages can enter the lytic cycle under certain stress conditions and cause bacterial cell death. This figure

was modified from Servier Medical Art, licensed under a Creative Common Attribution 3.0 Generic License. http://smart.servier.com/.

1.4 Co-evolution of bacteria and phages

Bacteriophages are ubiquitous across multiple natural ecosystems and have been shown to account for the turnover of 20 % of the living biomass in the sea (Koskella and Brockhurst

2014). This impact coupled with the reality that phages have been discovered for well over 140 bacterial genera makes phages a significant threat to bacterial survival (Ackermann 2003). To protect against this threat bacteria have evolved several mechanisms to defend against phage infection (Seed 2015). In return, phages are constantly evolving mechanisms to evade these bacterial defenses and infect their host (Samson et al. 2013). Collectively, this co-evolution is a perfect example of reciprocal adaptation and counter adaptation between two entities.

In order to defend against phage infection bacteria have evolved mechanisms to inhibit adsorption, kill the bacterial host prior to phage replication, and specifically target phage DNA for degradation, among others. In the first mechanism, bacteria alter the accessibility or distribution of a surface receptor required for infection and can therefore inhibit phage adsorption (Avrani et al. 2011). In the second mechanism, also called abortive infection, bacteria inhibit phage replication by inducing host cell death (Chopin et al. 2005). This mechanism is thought to be altruistic and inhibits propagation of the phage. In the third mechanism, bacteria can use restriction enzymes and/or CRISPR-Cas (Clustered Regularly Interspaced Short

Palindromic Repeats and CRISPR associated gene) systems to induce cleavage of phage DNA that has entered the bacterial cytoplasm (Barrangou et al. 2007). This cleavage results in an inactive phage. As expected, these bacterial defenses have been met by phage countermeasures,

including the mutation of phage receptor binding proteins to recognize modified bacterial surface receptors for adsorption and infection.

CRISPR-Cas systems represent one of the most recently discovered bacterial immune systems and are the only adaptive immune system characterized thus far. As such, the focus of this thesis is on CRISPR-Cas systems and their phage encoded inhibitors.

1.5 CRISPR-Cas systems (Class I and Class II)

Bacteria and phage have been undergoing co-evolution for billions of years (Serwer 2007; Keen

2015). Consequently, bacteria have evolved several defense mechanisms against phage predation. Some of these defenses include restriction modification systems, abortive infection, and the more recently discovered CRISPR-Cas immune defense. CRISPR-Cas systems represent a bacterial adaptive immune system that employs an RNA targeting complex to recognize, bind, and degrade a foreign phage genome. CRISPR-Cas systems are found in approximately 50 % of all sequenced Eubacteria and are classified into two classes, six types, and 24 subtypes (Koonin et al. 2017).

All class I systems are composed of multi-subunit RNA guided protein complexes that are responsible for immunity, all class II systems require only one RNA guided protein subunit for defense. Although each of these classes are genetically and structurally distinct, they all share a similar mechanism of action that is divided into three stages: adaptation, maturation, and target interference. For the purposes of this thesis, I will be focusing on the Class I Type I, and Class II

Type II CRISPR-Cas systems. During adaptation, Cas proteins acquire DNA fragments from a phage genome and integrate them into the CRISPR locus. Upon integration, these fragments are

called spacers and are flanked by semi-palindromic DNA repeats (Figure 4) (Levy et al. 2015;

Savitskaya et al. 2013; Wiedenheft et al. 2009; Barrangou et al. 2007; Nuñez et al. 2014). In maturation, the CRISPR array is transcribed as one large transcript. This transcript is cleaved at the repeat sequences (this step requires a Cas protein, and in some cases the host endoribonuclease III) to form small, mature CRISPR RNAs (crRNAs) that allow for the formation of a fully functional CRISPR-Cas targeting complex (Haurwitz et al. 2010; Brouns et al. 2008; R. Wang et al. 2011, 65; Przybilski et al. 2011; Carte et al. 2010) (Figure 4). During target interference, the crRNA guides the targeting complex to a foreign DNA sequence through sequence specific crRNA-DNA annealing, resulting in local dsDNA unwinding called R-loop formation (Xiao et al. 2017; Szczelkun et al. 2014) (Figure 4). If the DNA target sequence is correct, the CRISPR-Cas complex will induce DNA degradation (Zhang et al. 2012; Beloglazova et al. 2011; Hale et al. 2009; Marraffini and Sontheimer 2008; Garneau et al. 2010) (Figure 4).

Using this mechanism, CRISPR-Cas systems can theoretically target any phage that carries a sequence found in the CRISPR array, thereby inhibiting infection.

Figure 4: General representation of CRISPR-Cas function. The type I-E CRISPR-Cas system is used in this figure as a representative of the general function of CRISPR-Cas systems. Spacers are acquired 9

and integrated into the CRISPR-locus (colored boxes represent spacers; black boxes represent repeat regions). The CRISPR-locus produces one large transcript that requires a Cas protein for maturation. Upon maturation a functional targeting complex is formed. The targeting complex uses the spacer to anneal to a foreign DNA sequence that is a perfect match. Upon binding the type I-E CRISPR-Cas system recruits a nuclease/helicase protein called Cas3 to degrade the foreign DNA. In other systems, the targeting complex has native nuclease/helicase activity. (Red circle = Cas6. Purple circle = Cas8, Cas5, Cas7, Cse2. Yellow packman = Cas3).

This thesis is primarily focused on the Class I type I-E and Class II type II-A CRISPR-Cas

systems which will be discussed in further detail below.

1.5.1 Type I-E and Type II-A CRISPR-Cas mechanisms

Pseudomonas aeruginosa is a gram-negative bacterium that harbors two different class I-type I

CRISPR-Cas systems dubbed subtype I-E and I-F (Cady et al. 2012; Pawluk et al. 2014).

Although these two systems share no significant protein or nucleotide sequence similarity (Table

1) their overall structure and mechanisms are conserved.

Table 1: Type I-F and type I-E CRISPR-Cas sequence comparison.

Nucleotide sequence comparison Query Cover Identity Additional Information cas3e - cas3f 0 % 0 % cas3f and cas2f are fused cas8e - cas8f 0 % 0 % cas5e - cas5f 0 % 0 % cas7e - cas7f 0 % 0 % cas6e - cas6f 0 % 0 % cse2 NA NA cas1e - cas1f 0 % 0 % cas2e - cas2f 0 % 0 % cas3f and cas2f are fused Amino acid sequence comparison Cas3e - Cas3f 47 % 31 % Cas3f and Cas2f are fused Cas8e - Cas8f 13 % 34 % Cas5e - Cas5f 21 % 30 % Cas7e - Cas7f 22 % 40 % Cas6e - Cas6f 0 % 0 %

Cse2 NA NA Cas1 92 % 23 % Cas2 0 % 0 % Cas3f and Cas2f are fused

CRISPR loci contain semi-palindromic repeat sequences separated by spacer regions that can be identical to plasmid and phage DNA. These repeat spacer sequences form crRNAs through transcription and must be matured via Cas6 cleavage at the repeat regions (Gesner et al. 2011;

Haurwitz et al. 2010). A mature targeting complex for the type I-E system, also known as

CASCADE, is composed of one mature crRNA, one Cas8e, Cas5e, and Cas6e protein, two Cse2 proteins, and six subunits of Cas7e (Wiedenheft, Lander, et al. 2011; Jackson et al. 2014;

Mulepati et al. 2014) (Figure 5). For the type I-F system, a mature complex is composed of one mature crRNA, one Cas8f, Cas5f, and Cas6f protein, and six subunits of the Cas7f protein

(Richter and Fineran 2013; Wiedenheft, van Duijn, et al. 2011). A mature complex can interact with a DNA target or protospacer through crRNA sequence specific targeting and must recognize an adjacent DNA sequence (Protospacer adjacent motif) via Cas8 (Figure 5). For the

Type I-E system the PAM is 5’-CTT-3’ and for the type I-F system the PAM is 5’-GG-3’ found at the 3’ end of the complementary DNA strand (Gleditzsch et al. 2018). If the protospacer and

PAM sequence are correct, the CRISPR-Cas complex can recruit the nuclease-helicase protein

Cas3 and induce DNA degradation (Westra et al. 2012; Sinkunas et al. 2011; Mulepati and

Bailey 2011; Sinkunas et al. 2013) (Figure 5).

Figure 5: Type I-E and type I-F CRISPR-Cas targeting mechanism. For type I-E CRISPR-Cas targeting, a crRNA is matured through Cas6e cleavage. A matured crRNA allows for the nucleation of four more Cas proteins to form an active targeting complex. The complex locally unwinds foreign double stranded DNA and allows the spacer to anneal with the target. A perfect protospacer match and PAM allow for the recruitment of Cas3e and DNA degradation. For type I-F CRISPR-Cas targeting, a crRNA is matured through Cas6f cleavage. A matured crRNA allows for the nucleation of three more Cas proteins to form an active targeting complex. Local DNA unwinding and cleavage follows in a mechanism similar to type I-E unwinding and cleavage.

Streptococcus pyogenes is a gram-positive bacterium that harbors a Class II type II-A CRISPR-

Cas system. Although this system bears general similarities to the overall function of the type I systems as an adaptive immune mechanism, there are some important differences. Firstly, the type II-A CRISPR-Cas9 system does not use Cas6 to cleave and mature the crRNAs produced from its CRISPR array. Instead, maturation requires the Cas9 nuclease protein to be bound to a pre-crRNA that interacts with a tracrRNA to form a section of dsRNA. This allows for the subsequent maturation of the crRNA via cleavage by the hosts endoribonuclease III protein

(RNase III) and additional crRNA trimming (Barrangou and Marraffini 2014) (Figure 6). Once maturation has been achieved the crRNA-tracrRNA-Cas9 complex can be guided to a phage protospacer for complementary spacer annealing. Upon binding, Cas9 recognizes a three

nucleotide PAM (5’-NGG-3’) found at the 5’ end of the complementary DNA strand and uses its

RuvC and HNH nuclease domains to cleave both DNA strands (Nishimasu et al. 2014; Anders

et al. 2014) (Figure 6).

Figure 6: Type II CRISPR-Cas9 recognition and targeting. Cas9 initially loads a crRNA and a tracrRNA, RNAse III is then recruited and allows for cleavage and partial maturation of the crRNA. Full maturation occurs with additional 5’ trimming. Once a mature complex has formed Cas9 unwinds the DNA target and allows the spacer to anneal (the spacer is colored red, the repeat is colored blue). If the spacer match anneals perfectly with the target and the PAM is correct, Cas9 uses its HNH and RuvC nuclease domains to cut the DNA. The PAM motif is recognized by Cas9 through protein – nucleic acid interactions and is independent of the crRNA.

1.6 Discovery of CRISPR-Cas inhibitors

From the perspective of an infecting phage, CRISPR-Cas systems pose a great threat to infection.

An adaptive immune system, that can theoretically acquire crRNAs against any DNA target

would be able to stop infection at the point of DNA entry. So, the question arose, how do phages evade CRISPR-Cas systems, and how do they manage to infect. The first ideas of CRISPR-Cas evasion considered the possibility of phages mutating their DNA sequences to evade CRISPR-

Cas targeting. Experiments showed that phages which acquired point mutations within the PAM or protospacer regions were able to escape CRISPR-Cas targeting and infect the host, suggesting that mutation is a mechanism used to evade CRISPR targeting (Semenova et al. 2011;

Wiedenheft, van Duijn, et al. 2011; Barrangou et al. 2007). It was not until the discovery of phage encoded CRISPR-Cas inhibitors that we realized phages were also capable of actively inhibiting CRISPR-Cas systems.

The first discovery of a phage encoded CRISPR-Cas inhibitor was made in P. aeruginosa

DMS3-like phages (Bondy-Denomy et al. 2013). In this work five distinct anti-CRISPR (Acr) genes found in Siphophages coded for small protein molecules that could inhibit the type I-F

CRISPR-Cas system of P. aeruginosa . Further work showed that some of the Joe-Bondy

Denomy (JBD) Siphophages also coded Acr proteins that could inhibit the type I-E CRISRP-Cas system of P. aeruginosa. (Pawluk et al. 2014). As the field progressed 23 families of Acr proteins against a number of type I and type II CRISPR-Cas systems were found in a number of phages and mobile genetic elements (Y. Zhu, Zhang, and Huang 2018). Collectively, this suggests that Acr proteins may be a widely used mechanism of Acr dependent CRISPR-Cas inhibition.

1.6.1 Anti-CRISPR mechanisms

To date, for P. aeruginosa , four type I-F Acr proteins and one type I-E protein have been structurally and mechanistically characterized and published (Figure 7). Of the Acr proteins that

inhibit the I-F system, AcrIF1 was shown to interact with the Cas7f backbone resulting in a

CRISPR-Cas complex that cannot hybridize to its DNA target (Bondy-Denomy et al. 2015; Peng et al. 2017; Maxwell et al. 2016). AcrIF2 was shown to interact with the Cas8f and Cas7f subunits of the type I-F system, suggesting that it may block a critical DNA binding site (Bondy-

Denomy et al. 2015). In a similar mechanism to AcrIF2, AcrIF10 could also block a critical site for DNA binding by interacting with the Cas8f and Cas5f subunits (Bondy-Denomy et al. 2015;

Guo et al. 2017). Furthermore, AcrIF3 was shown to interact with the Cas3f subunit and allows for CRISPR-Cas DNA binding but inhibits DNA cleavage (Bondy-Denomy et al. 2015). For the

I-E system, AcrIE1 has been the first and only published I-E Acr protein mechanism thus far

(Pawluk et al. 2017). AcrIE1 was shown to interact with the Cas3e subunit and much like AcrIF3 allows for DNA binding but inhibits Cas3 dependent DNA degradation. The functioning of

AcrIE1 and AcrIF3 results in a CRISPR-Cas complex that can bind DNA and inhibit the expression of a nearby gene by sterically blocking RNA polymerase dependent transcription

(Bondy-Denomy et al. 2015; Pawluk et al. 2017). This transcriptional repression mechanism can be used as a biotechnological tool to repress targeted genes. Based on these findings, additional research into Acr proteins will allow us to discover new mechanisms of anti-CRISPR activity, develop new biotechnological tools, and gain insight into the intricate evolutionary interactions between CRISPR-Cas and anti-CRISPR proteins.

Figure 7: Mechanisms of CRISPR-Cas inhibition. Schematic of previously published anti-CRISPR dependent CRISPR-Cas inhibition for the type I-F and type I-E CRISPR-Cas systems. (1) AcrIF3 interacts with the Cas3f protein of the type I-F system and inhibits its recruitment to the targeting complex. (2) AcrIE1 uses a mechanism similar to AcrIF3 but interacts with the Cas3e protein of the type I-E system. (3) AcrIF1 interacts with the Cas7f backbone of the type I-F targeting complex. (4) AcrIF2 interacts with Cas8f and Cas7f of the type I-F system. (5) AcrIF10 interacts with Cas8f and Cas5f. AcrIF1, IF2, and IF10 all inhibit DNA annealing.

1.7 CRISPR-Cas: As a biotechnological tool

Following the discovery of CRISPR-Cas systems, the CRISPR field moved in two directions.

The first, was focused on understanding the evolution and mechanism of CRISPR-Cas systems and CRISPR-Cas inhibitors. This focus was summarized above and reflects the first portion of my thesis. The second, focused on using CRISPR-Cas systems as a biotechnological tool for the purposes of genome editing. This will represent the second part of my thesis and is discussed below.

1.8 CRISPR-Cas used for genome editing

The bacterial chromosome is composed of millions of DNA bases. The ability to alter these nucleotides at precise locations has important implications for molecular biology, as precise edits are essential for understanding the function of specific genes. In light of this, scientists have spent years developing new technologies to alter and manipulate the bacterial genome.

Traditional methods of genome editing in bacteria have focused primarily on using homologous recombination (HR) of a drug cassette, vector based genomic transmission, and transposon-based mutagenesis (Hmelo et al. 2015; Hoang et al. 1998; Kulasekara 2014). These methods, although effective, are often laborious and come with their own caveats. One recent technology that has significantly improved the efficiency of genomic mutations in bacteria as well as eukaryotes is

CRISPR-Cas9.

There are two mechanisms of creating mutations in the genome of a bacterium using CRISPR-

Cas. The first method is HR independent and the second method requires HR (Figure 8). HR independent mutagenesis requires the use of a plasmid encoding the Cas protein(s) along with a

crRNA that can target a specific region of the host genome (Su et al. 2016). This plasmid is transformed into the bacterial cell, and a CRISPR-Cas targeting complex forms, anneals to the sequence of interest, and cleaves the genome, resulting in a significant number of cells dying.

Following cleavage, the remaining bacterial organism can use a naturally expressed nonhomologous end joining (NHEJ) system that allows for reattachment of the two DNA strands (it should be noted that not all bacterial organisms have an NHEJ system) (Zhu and Shuman 2008;

Shuman and Glickman 2007; Chayot et al. 2010). The resulting colonies have repaired the damage and contain a random mutation that could be anything from a silent mutation to a large deletion. This method of mutagenesis is random and can sometimes have negative effects on the expression of surrounding genes. In order to make more specific mutations a method that uses

HR in conjunction with CRISPR-Cas was developed.

In HR dependent mutagenesis, the cells are co-transformed with a plasmid carrying the mutant version of a gene of interest as well as the CRISPR-Cas targeting plasmid (Figure 8) (Selle and

Barrangou 2015). Upon transformation, cells cleave the WT gene but not the mutant and a small proportion of them will recombine the mutant gene into their chromosome. This targeting results in a mixture of cells that carry either a random mutation due to NHEJ (described above) or the specific mutation selected for via recombination. With sequence verification this method permits the selective mutagenesis of specific regions anywhere on the bacterial genome for research purposes.

Figure 8: CRISPR-Cas mechanisms for genome editing. In the absence of HR, a crRNA and appropriate Cas gene(s) are transformed into a bacterial cell. A targeting complex forms, anneals to the appropriate DNA region, and cleaves. NHEJ is then used to randomly repair the damage resulting in a mutation (depicted as a red segment). Genome editing in the presence of HR, requires a plasmid to be transformed into the cells that will recombine a mutant gene into the chromosome. A plasmid carrying the

appropriate Cas gene(s) and a crRNA targeting the WT gene is also transformed. Surviving colonies either have a random mutation, or a specific recombined DNA segment (sequencing is usually necessary to differentiate between random mutants and the recombined gene). This figure was modified from Servier Medical Art, licensed under a Creative Common Attribution 3.0 Generic License. http://smart.servier.com/.

1.8.1 Type I-E CRISPR-Cas mechanism for biotechnology

One of the CRISPR-Cas systems used for genome editing is the Type I-E CRISPR-Cas system

(Kiro, Shitrit, and Qimron 2014). This system was used in Escherichia coli to make a mutation in the T7 phage genome. In work done by Kiro et al, the researchers used HR to create a mixture of WT and mutant T7 phages and subsequently selected against WT phage using the type I-E

CRISPR Cas system. The result was a clean deletion of a nucleotide kinase gene from the phage genome. Noting that P. aeruginosa strain SMC 4386 endogenously expresses an active type I-E

CRISPR-Cas system we wanted to determine whether we could use this type I-E system for genome editing in P. aeruginosa .

One of the first attempts at using the type I-E CRISPR-Cas system for genome editing in P. aeruginosa was conducted in our lab (Pawluk et al. 2016a). In this experiment, the cells were transformed with a plasmid carrying a type I-E crRNA targeting the Cas8 gene of P. aeruginosa .

The result was a reduction in transformation efficiency and one Pseudomonas mutant containing a 125kb genomic deletion that included both P. aeruginosa CRISPR arrays and all the Cas genes. Unfortunately, this mutant also included the loss of numerous other genes in the surrounding region, making it a relatively poor control for downstream experiments. To improve the efficacy of type I-E CRISPR-Cas genome editing I used HR in conjunction with type I-E

CRISPR-Cas targeting to make selective mutations which will be discussed in chapter 3 (Pawluk et al. 2017).

1.8.2 Type II-A CRISPR-Cas mechanism for biotechnology

The primary system used for genome editing today is the Streptococcus pyogenes CRISPR-Cas9.

Unlike the type I-E CRISPR-Cas system which requires the expression of multiple Cas genes for targeting and nuclease activity, the CRISPR-Cas9 system requires only one approximately 4 kb gene and a crRNA to be cloned into a plasmid and expressed (Jiang et al. 2013). This makes

Cas9 a simpler and more versatile genome editing tool to use. As expected, CRISPR-Cas9 has been used for genome editing in a variety of eukaryotic model systems and bacteria (Jiang et al.

2015; Ng and Dean 2017). Much like the type I-E CRISPR-Cas system, Cas9 allows for recombination dependent and independent mechanisms of genome editing in bacteria and can be used to edit the genome of different P. aeruginosa strains (discussed in chapter 3).

1.9 Thesis Objectives

In the first section of this work, the structure, function, and mechanism of action for the type I-E

Acr protein AcrIE2 is determined. Here, AcrIE2 is shown to interact with the I-E CRISPR-Cas complex while allowing CRISPR-Cas DNA binding, but inhibiting CRISPR-Cas DNA cleavage.

Furthermore, exploiting the ability of AcrIE2 to interact with the CRISPR-Cas complex I show that AcrIE2 allows us to turn the type I-E CRISPR-Cas system into a transcriptional activator and repressor. In the second section of this thesis, genome editing technologies are developed for

P. aeruginosa using the type I-E CRISPR-Cas system of P. aeruginosa strain SMC 4386 and type II-A CRISPR-Cas9 system of S. pyogenes .

1.10 Thesis Outline

In chapter 2, I focus on my work in characterizing the type I-E CRISPR-Cas inhibitor called

AcrIE2. Noting that AcrIE2 was previously shown to inhibit the type I-E CRISPR-Cas system of

P. aeruginosa , I hypothesized that AcrIE2 was likely interacting with one or more subunits of the CRISPR-Cas system to inhibit DNA degradation. To that end, I showed that AcrIE2 interacts with the CASCADE complex and does not simply inhibit CRISPR-Cas targeting but also alters the function of CRISPR-Cas from DNA degradation to transcriptional regulation. This work suggests that there may be more to the development and evolution of Acr proteins than simply inhibiting CRISPR-Cas nuclease function and enabling phage infection.

In chapter 3, I focus on my work in developing genome editing tools to study P. aeruginosa in a laboratory setting. With the success of CRISPR genome editing in a variety of cell lines and bacteria, I hypothesized that a CRISPR mediated genome editing tool could be developed for P. aeruginosa . In this work, I have shown that the endogenously expressed type I-E CRISPR-Cas system of P. aeruginosa SMC 4386 can be used to make specific mutations in the host genome.

Moreover, I developed a versatile CRISPR-Cas9 system that can be used in P. aeruginosa to edit the genome. I and others in the lab have shown that this system allows us to specifically edit the genome of different P. aeruginosa strains as well as edit the genome of P. aeruginosa phages.

Collectively, this work provides new tools for the efficient genetic engineering of P. aeruginosa strains for the purposes of research. This work may have implications in understanding P. aeruginosa physiology and pathogenesis with regards to infection.

Chapter 2

Anti-CRISPR AcrIE2: A novel mechanism of CRISPR-Cas inhibition

2.1 Acknowledgements

The type I-E anti-CRISPR AcrIE2 was originally cloned by Joe Bondy-Denomy. All JBD phages were isolated by Joe Bondy-Denomy. P. aeruginosa SMC strains were provided by the George

O’Toole Lab at Dartmouth College. The NMR structure of AcrIE2 was solved by April Pawluk and Karen Maxwell. This research was funded with support from the Canada Institutes of Health

Research (CIHR).

2.2 Abstract

Anti-CRISPRs are protein inhibitors of CRISPR-Cas systems and are produced by phages to evade CRISPR-Cas-mediated destruction. Anti-CRISPRs function through a variety of mechanisms which allow us to gain further insight into CRISPR-Cas function, and develop new anti-CRISPR-based biotechnological tools. In this study, I present the protein structure, define the functional pocket, and describe the mechanism of action for the type I-E anti-CRISPR

AcrIE2. We determine the structure of AcrIE2 using nuclear magnetic resonance (NMR) spectroscopy and we use mutagenesis to identify the residues responsible for AcrIE2 function. I then use affinity co-purification to determine that AcrIE2 interacts with proteins found in the type I-E CRISPR-Cas surveillance complex (CASCADE). Furthermore, I use a transcriptional reporter assay to determine the mechanism of action for this anti-CRISPR. Using this assay, I show that AcrIE2 allows the CASCADE complex to bind its genomic target while inhibiting

CRISPR-Cas DNA degradation. Moreover, my data show that AcrIE2 can turn the CASCADE

complex into a transcriptional regulator, displaying both anti-CRISPR dependent transcriptional repression and, for the first time, anti-CRISPR dependent transcriptional activation. Taken together, this data suggests that AcrIE2 may have two functions – the first being CRISPR-Cas inactivation, and the second being CRISPR-Cas dependent transcriptional regulation.

2.3 Anti-CRISPR AcrIE2

Previously completed work showed that gene 32 (coding for the AcrIE2 protein) of the P. aeruginosa phage JBD88a could inhibit the type I-E CRISPR-Cas system of the P. aeruginosa strain SMC 4386 (Pawluk et al. 2014). In a phage spotting assay, AcrIE2 expression was able to inhibit CRISPR-Cas dependent targeting of a CRISPR sensitive phage, resulting in successful phage infection. To understand more about the activity and mechanism of AcrIE2, the structure of this protein was solved by April Pawluk using nuclear magnetic resonance (NMR) spectroscopy with an RMSD of 1.6 Å.

The structure determined for AcrIE2 is shown in Figure 9a. The structure of AcrIE2 is an 84- residue monomer with a central α-helix and a five-strand antiparallel β-sheet (Figure 9a, b). The

α-helix packs against one side of the β-sheet, forming a tight hydrophobic core with 10 residues

>90 % buried. The structure of AcrIE2 is not similar to any previously determined Acr protein, and searches using the Dali server did not reveal structural similarity to any other protein (Holm and Rosenström 2010).

Figure 9: NMR structure AcrIE2. (a) The NMR solution structure of AcrIE2 showing the backbone atoms in ribbon representation (N, C α and C’). (b) Secondary structure of AcrIE2 with relative residue accessibility displayed in different shades of blue. (c) Amino acid alignment of the JBD88a-AcrIE2 protein using Blastp.

2.4 Results

2.4.1 AcrIE2 functional residues

To identify the AcrIE2 residue(s) responsible for inhibiting the CRISPR-Cas system, I wanted to combine structural and sequence analysis with site-directed mutagenesis. Unfortunately, amino acid sequence analysis using Blastp showed that all AcrIE2 homologues were found in P. aeruginosa and shared more than 88 % identity across a 73 % query coverage, making it difficult to predict functional residues (Figure 9c). As such, 26 site-directed substitutions were made by

April Pawluk and myself on AcrIE2 solvent exposed residues, as determined by the NMR structure (Figure 10). The function of these point mutations was tested in a phage spotting assay.

None of the mutations resulted in a significant loss of AcrIE2 activity Table 2.

Figure 10: Point mutations of AcrIE2. All AcrIE2 alanine scanning point mutations created to determine functional residues.

Table 2: Point mutations of AcrIE2. Protein Name Mutation Fold reduction in phage titre AcrIE2 WT 0 AcrIE2 R9A 10 AcrIE2 K10A 0 AcrIE2 K10E 0 26

AcrIE2 N11A 0 AcrIE2 D13A 0 AcrIE2 S15A 0 AcrIE2 E17K 0 AcrIE2 R18A 0 AcrIE2 F19A 10 AcrIE2 V24A 0 AcrIE2 K31A 0 AcrIE2 S32A 0 AcrIE2 Q35A 10 AcrIE2 Q36A 0 AcrIE2 E40A 0 AcrIE2 E43A 0 AcrIE2 L45A 0 AcrIE2 Y47A 0 AcrIE2 R48A 10 AcrIE2 M58A 0 AcrIE2 Y62A 0 AcrIE2 H64A 0 AcrIE2 L65A 0 AcrIE2 H66A 0 AcrIE2 R70A 10 AcrIE2 Y77A 0 AcrIE2 Y77S 0 AcrIE2 H80A 0 AcrIE2 M82A 0

This resilience in AcrIE2 activity pushed me to perform additional mutagenesis experiments.

The first approach used site-directed mutagenesis to make AcrIE2 double mutants, with a focus on residues that were in close proximity to each other in the NMR structure. The second approach made multiple mutations on the surface of AcrIE2 in a more random fashion (see methods). I made more than 40 combinatorial mutations and the majority of these mutants were found to be inactive via phage spotting (Table 3). To narrow the search for the functional pocket of AcrIE2 I looked for the smallest number of mutated residues that were in close structural proximity and resulted in a loss of activity. Using these parameters, the AcrIE2 A10 and AcrIE2

R9A/H80A mutants were good candidates (Table 3). AcrIE2 A10 had mutations in residues 27

M58, H80, and M82, these residues were in close structural proximity to each other and showed a 10 4-fold reduction in activity during phage spotting (Table 3, Figure 11a). AcrIE2 R9A/H80A was an interesting double mutant as it contained both an H80 mutation (the most common mutation found in Table 3 and a mutation that does not reduce AcrIE2 activity in Table 2) as well as an R9 mutation which was shown to reduce AcrIE2 activity 10-fold in Table 2. The R9 residue was also in close structural proximity to H80 and a double mutant of these two residues resulted in a 10 5-fold reduction in activity in phage spotting (Figure 11a). Phage spotting experiments for both AcrIE2 inactive mutants were controlled using AcrIE2 H80A, R9A, and

M58V/M82R mutants, all of which showed close to wild type levels of AcrIE2 activity (Figure

11a). Figures 11b and c display the residues mutated in AcrIE2 A10 and AcrIE2 R9A/H80A, respectively. Looking at the location of all four residues, it became clear that R9, M58, H80, and

M82 are in close proximity to each other, suggesting that these residues represent a functional pocket on the surface of AcrIE2 (Figure 11d).

Table 3: Mutagenesis of multiple AcrIE2 residues. Protein Name Mutation Fold reduction in phage titer AcrIE2-R9A-H80A R9A/H80A 10^5 AcrIE2-B10 aa19-23 Truncation/Y62S/H64L/H66A/E72A/H80A 10^7 AcrIE2-B11 A30X/K31X/Y62S/H64A/H66A/N69S/Y77S/H80A 10^7 AcrIE2-B12 H64A/H66A/E72A/T74A/Y77S/H80A 10^7 AcrIE2-B13 Y47S/Y62S/H64A/H66A/T74A/H80A 10^7 AcrIE2-C11 D22A/K31T/Q35L/Q36L/E40A/E43A/L65R 10^3 AcrIE2-B16 H64A/ H66A/ E72A/ T74A/ H80A 10^7 AcrIE2-B17 Y62S/H64A/H66A/N69S/T74A/H80A 10^7 AcrIE2-B18 Y62S/H64A/H66A/E72A/H80A 10^7 AcrIE2-B19 No E43 AND G44... 10^7 Y62S/H64A/H66A/E72A/T74A/H80A AcrIE2-B20 Y62S/H64A/H66A/N69S/H80A 10^7 AcrIE2-B21 H64A/ H66A/ T74A/ Y77S/ H80A 10^7

AcrIE2-B23 Y47S/Y62S/H64A/H66A/N69S/T74A/H80A 10^7 AcrIE2-B24 Y47S/Y62S/H64A/H66A/H80A 10^7 AcrIE2-B27 Y47S/Y62S/H64A/H66A/T74A/Y77S/H80A 10^7 AcrIE2-B28 Y62S/H64A/H66A/E72A/T74A/ Y77S/ H80A 10^7 AcrIE2-B30 H64A/ H66A/ E72A/ T74A/ Y77S/ H80A 10^7 AcrIE2-B32 H64A/ H66A/ E72A/Y77S/ H80A 10^7 AcrIE2-B34 Y47S/Y62S/H64A/H66A/N69S/T74A/Y77S/H80A 10^7 AcrIE2-A14 R9S/N14S/E17A/E52A/M58V/H80A 10^7 AcrIE2-A15 R9S/N12S/N14S/E17A/H80A/M82R 10^7 AcrIE2-A16 R9S/N12S/N14S/E52A/M58V/H80A 10^7 AcrIE2-C13 D13A/S15A/D22A/E40A/E41A/E43A/L65R 10^4 AcrIE2-C15 D13A/Q35L/Q36L/L45V/L65R 10^2 AcrIE2-C17 N11S/D13A/S15A/T27A/K31T/E41A/L45V 10^3 AcrIE2-A10 M58V/H80A/M82R 10^4 AcrIE2-A10-A80D1 M58V/H80D/M82R 10^ 4 AcrIE2-A10-A80H1 M58V/M82R 10^ 1 AcrIE2-A10-R82D1 M58V/H80A/M82D 10^ 4 AcrIE2-A10-R82L1 M58V/H80A/M82L 10^ 4 AcrIE2-A10-R82M1 M58V/H80A 10^ 2 AcrIE2-A10-V58A2 M58A/H80A/M82R 10^ 4 AcrIE2-A10-V58D1 M58D/H80A/M82R 10^ 4 AcrIE2-A10-V58L2 M58L/H80A/M82R 10^ 4 AcrIE2-A10-V58M1 H80A/M82R 10^ 3 AcrIE2-A10-(All alanine)- M58A/H80A/M82A 10^7 AcrIE2-A10-(All M58M/H80A/M82A 10^3 alanine/M58) AcrIE2-A10-(All M58A/H80A/M82M 10^2 alanine/M82) AcrIE2-B12-(H80)-2 H64A/H66A/E72A/T74A/Y77S/H80H 10^4 AcrIE2-B13-(H80)-2 Y47S/Y62S/H64A/H66A/T74A/H80H 10^7 AcrIE2-B23-(H80)-1 Y47S/Y62S/H64A/H66A/N69S/T74A/H80H 10^7 AcrIE2-B27-(H80)-1 Y47S/Y62S/H64A/H66A/T74A/Y77S/H80H 10^7 AcrIE2-B34-(H80)-1 Y47S/Y62S/H64A/H66A/N69S/T74A/Y77S/H80H 10^7 AcrIE2-A16-(H80)-1 R9S/N12S/N14S/E52A/M58V/H80H 10^7

Figure 11: Functional residues of AcrIE2. (a) Tenfold dilutions of a CRISPR-sensitive phage (JBD8) spotted on bacterial lawns of P. aeruginosa 4386 expressing AcrIE2 WT or an AcrIE2 mutant from a plasmid. Zones of clearing represent phage replication and AcrIE2 activity. The replication efficiency of a CRISPR-insensitive phage, JBD93a, was constant (data not shown). (b) AcrIE2 structure showing the positions of two residues (in blue) that are critical for AcrIE2 function. (c) AcrIE2 structure showing the positions of three residues (in blue) that are critical for AcrIE2 function. (d) AcrIE2 structure showing the positions of all five residues (in blue) that likely form the AcrIE2 functional pocket.

To determine if AcrIE2 R9A/H80A and AcrIE2 M58V/H80A/M82R were still expressed, protein purification out of P. aeruginosa was performed. 6XHis-AcrIE2 WT, 6XHis-AcrIE2

R9A/H80A, and 6XHis-AcrIE2 M58V/H80A/M82R were induced and purified out of P. aeruginosa SMC 4386, all proteins were expressed (Figure 12).

Figure 12: Protein expression of AcrIE2 WT and AcrIE2 inactive mutants from P. aeruginosa . Western blot of 6XHis tagged AcrIE2 WT, and AcrIE2 mutants purified by nickel affinity purification.

Noting that AcrIE2 R9A/H80A and AcrIE2 M58V/H80A/M82R were expressed in P. aeruginosa, I wanted to purify these inactive mutants and subject them to structural analysis. To do this, I used circular dichroism (CD). CD uses circularly polarized light to attain information about the overall secondary structure and stability of a protein. To conduct CD both inactive mutants and AcrIE2 WT were expressed and purified out of Escherichia coli using nickel affinity purification (Figure 13a). The purifications were then run through a size exclusion column via FPLC (Figure 13b). As expected, all three proteins eluted at the same volume

suggesting they were similar in size. Following size exclusion chromatography (SEC), the proteins were structurally compared using CD. All three proteins had similar CD spectra, suggesting that they were structurally similar and that the inactive mutants were properly folded

(Figure 14a). To assess the stability of the inactive AcrIE2 mutants a heat-induced protein unfolding experiment was conducted using the CD machine. The results showed similar midpoint melting temperatures (TM) for all three proteins, namely, 42.5 °C for AcrIE2 WT, and

39.5 °C for both the inactive mutants, suggesting that all three proteins had similar stabilities

(Figure 14b). These data suggest that both AcrIE2 mutants were expressed in Pseudomonas , had no gross structural defects, and were similar in stability to AcrIE2 WT. Collectively, these results show that the loss of function observed in the two inactive AcrIE2 mutants was due to mutations in residues important for the interaction with the CRISPR-Cas system.

Figure 13: Protein purification and SEC of AcrIE2 and AcrIE2 mutants for circular dichroism. a, SDS-PAGE gel stained with Coomassie Blue showing purified AcrIE2 and mutant AcrIE2 proteins prior to FPLC. b, SEC chromatogram of AcrIE2 WT and mutant AcrIE2 proteins.

Figure 14: Circular dichroism analysis of AcrIE2 mutants. a, Circular dichroism (CD) spectral overlay of WT AcrIE2 and mutant AcrIE2 proteins conducted at 25 °C with a 60 μM protein concentration in Tris-buffered NaCl. b, Heat-induced protein unfolding experiment monitored by CD. Overlay of WT AcrIE2 and mutant AcrIE2 proteins measured at 210 nm ranging from a temperature of 10 °C to 80 °C using the same concentration and buffer as in a.

2.4.2 AcrIE2 interacting partners

Noting that AcrIE2 could inhibit the Type I-E CRISPR-Cas system of P. aeruginosa and that previous Acr proteins were able to interact with CRISPR-Cas subunits, I hypothesized that

AcrIE2 was likely inhibiting the CRISPR-Cas complex by interacting with one or more

CRISPR-Cas subunits. To determine the interacting partner(s) of AcrIE2, a co-purification experiment was performed. P. aeruginosa SMC 4386 cells natively expressing the CRISPR-Cas system were transformed with a plasmid expressing either 6XHis-tagged AcrIE1 (control) or

AcrIE2 WT. These proteins were then purified through nickel affinity chromatography, and the eluted proteins were analyzed by mass spectrometry (Table 3). As expected, the AcrIE1 control co-purified with Cas3 consistent with our previously published work (Pawluk et al. 2017) (Table

3). AcrIE2 co-eluted with Cas7e with no Cas3 present (n=3) (Table 3). Notably, Cas7e was a top hit in the mass spectrometry analysis for AcrIE2 and could also be observed on an SDS-PAGE gel (Figure 15). Cas7e is part of the CASCADE targeting complex that is responsible for binding

DNA. Additional mass spectrometry analysis of the AcrIE2 elution revealed that AcrIE2 was also capable of pulling down Cas6e, Cas5e, and Cse2, suggesting that AcrIE2 may be interacting with an intact Type I-E CASCADE complex (Table 3). These data suggest that AcrIE1 binds to

Cas3, while AcrIE2 likely interacts with Cas7e and potentially the CASCADE complex

(although it should be noted that Cas8e was not present). It is possible that AcrIE2 interacts with the CASCADE targeting complex and Cas7e, being the most abundant protein in the complex, shows up as a top hit in mass spectrometry and can be seen on a gel. Irrespective of whether

AcrIE2 interacts specifically with Cas7e or the entire CASCADE complex, these data suggest that AcrIE2 can interact with at least one Cas protein, suggesting that this interaction is likely important for AcrIE2 dependent CRISPR-Cas inhibition. 35

Table 4: Mass spectrometry results summary.

2.4.3 AcrIE2 mutants

The observation that AcrIE2 WT and Cas7e were pulled down together pushed me to conduct the same affinity purification experiments found in section 2.4.2 with both AcrIE2

M58V/H80A/M82R and AcrIE2 R9AH80A. I was interested in seeing whether the residues mutated on the surface of these inactive proteins were responsible for pulling down Cas7e (as observed on a protein gel), which would lend further support to the idea that AcrIE2 needs to interact with either Cas7e or the targeting complex in order to be an effective Acr protein. As such, a nickel affinity co-purification experiment was conducted in P. aeruginosa natively expressing the type I-E CRISPR-Cas system as well as, either 6XHis-tagged AcrIE1, AcrIE2

WT, AcrIE2 R9A/H80A, or AcrIE2 M58V/H80A/M82R (Figure 15). The goal of this experiment was to investigate whether the inactive mutants could still pulldown the natively expressed Cas7e protein. Upon purification, AcrIE1 co-eluted with Cas3 and AcrIE2 WT co- eluted with Cas7e as expected (Figure 15). Importantly, both AcrIE2 R9A/H80A and AcrIE2

M58V/H80A/M82R did not co-elute with Cas7e, suggesting that the interaction required to pulldown Cas7e was abrogated (Figure 15). This data suggests that our inactive mutants cannot inhibit the CRISPR-Cas system because they lose the ability to interact with their Cas protein target.

Figure 15: AcrIE2 affinity purification and interactors. (a) Silver stained gel of 6Xhis tagged AcrIE1, AcrIE2, and AcrIE2 mutants purified by nickel affinity purification. AcrIE1 interacts with Cas3, and AcrIE2 interacts with Cas7. No Cas7 is apparent for either AcrIE2 mutant. (n=3)

2.4.4 AcrIE2 inhibits DNA degradation but permits DNA binding

To determine the mechanism of AcrIE2 activity, a transcriptional reporter assay was used. In this assay, the CASCADE complex was targeted to the promoter region of phzM , a gene responsible for producing the blue green pigment pyocyanin. phzM expression is determined by the LasR transcription factor (a transcriptional regulator of quorum sensing and virulence genes) which is found approximately 35 to 40 nucleotides upstream of the phzM transcriptional start site (Huang et al. 2009; Wurtzel et al. 2012). The -35 and -10 regions of the phzM gene are predicted to be found between the LasR binding site and the transcriptional start site (Figure 16a). When targeting the CRISPR-Cas complex to the phzM promoter region in the absence of Cas3 activity, the CASCADE complex is expected to bind the phzM promoter region and sterically inhibit

RNA polymerase from accessing and transcribing phzM . Using a P. aeruginosa Cas3 nuclease deficient strain, the CASCADE complex was targeted to two regions of the phzM promoter and two regions downstream of the phzM transcriptional start site (Figure 16a). For every crRNA the result was decreased phzM mRNA production as seen through RT-qPCR, and a complete loss of green pigment in culture (Figure 16b, 16c).

a phzM Promoter Targets LasR Binding Site 5’ ATAAAACACAGAACGCTCGTACCGGCTC AACTACAAGATCTGGTAGGT GCCAGACAGGGTATGCGGG

crRNA 1: -47/ -15 crRNA 2: -8/+23 crRNA 4: +27/+59

ATTGCTAAGCTGATGCTTCCTGCAATGCCGGAGGTTGTAGCCAAGTTGTAATTTTATTCTTTCGGTACGC TSS crRNA 3: +8/+40 3’ 206 nt AGGAAAAGG………………ATG phzM Start Codon

b c 120 4386 Cas3 (Nu-)

100

60 rRNA/ no crRNAnorRNA/ %) 40 Pyocyanin Expression phzM c phzM ( 20

2+ 1+ A 3- 4+ tor N c rR crRNA crRNA c crRNA Empty Ve

Figure 16: CRISPR-Cas dependent transcriptional repression of phzM . a, Schematic of the phzM promoter sequences targeted by different crRNAs in the transcriptional reporter assay. (TSS represents the transcriptional start site). b, Pyocyanin expression levels measured out of P. aeruginosa cells carrying phzM promoter targeting crRNAs in the absence of both AcrIE2 and an active Cas3 protein. The + sign represents targeting the non-coding strand, the - sign represents targeting the coding strand, Cas3 (Nu-) means Cas3 nuclease deficient. c, RT-qPCR results reflecting mRNA reduction in the presence of phzM 40

promoter targeting crRNAs and in the absence of both AcrIE2 and an active Cas3 protein (statistics are defined by One-way ANOVA, P < 0.01, n=2). Error bars represent standard deviation.

To assay the mechanism of AcrIE2 activity in vivo , AcrIE2 was expressed in the presence of a nuclease deficient Cas3, an intact CASCADE complex and each of the four crRNA targets. If

AcrIE2 inhibits CASCADE from interacting with its phzM target, pyocyanin will be produced at

WT levels. If AcrIE2 allows the CASCADE complex to bind its phzM target, then phzM will be repressed. Expression of AcrIE2 in the presence of a CASCADE complex targeting the phzM promoter resulted in significant phzM repression, suggesting that AcrIE2 allows the complex to bind its target while interacting with the CASCADE complex (Figure 17a, 17c). Expression of

AcrIE2 in the presence of a CASCADE complex targeting regions downstream of the phzM transcriptional start site resulted in phzM overexpression (Figure 17a, 17c). The overexpression of phzM mediated by a CRISPR-Cas complex has never been seen before and suggests that RNA polymerase can knock off the DNA bound CASCADE-AcrIE2 complex to transcribe the gene.

This phenotype is not observed in the absence of AcrIE2 (Figure 16b, c). Therefore, in the presence of AcrIE2, the CASCADE complex has a lower affinity for its DNA target when compared to CASCADE targeting alone. This is likely attributed to a conformational change in the CRISPR-Cas complex when AcrIE2 interacts with CASCADE. However, this does not explain how an overexpression phenotype is observed. When focusing on the overexpression phenotype, in the presence of AcrIE2 the CASCADE complex interacts with its target and unwinds the DNA helix directly downstream of the transcriptional start site (R-loop formation).

This unwinding leaves the promoter of phzM relatively unperturbed allowing for RNA polymerase binding. Moreover, because the R-loop has already unwound the downstream DNA, it is possible that RNA polymerase can more easily initiate successful transcription. These processes collectively, result in an RNA polymerase that binds the phzM promoter, is more likely 41

to successfully initiate transcription and can knock off CASCADE, resulting in phzM overexpression.

All the above transcriptional reporter assays were also conducted in the presence of an active

Cas3 protein and the results remained the same (Figure 17b). Collectively, these data coupled with the in vivo pulldown assays suggest that AcrIE2 interacts with the CRISPR-Cas complex and allows for DNA binding while inhibiting DNA cleavage. Although these assays shed light on the mechanism of AcrIE2 activity, the question arose, can AcrIE2 through its interaction with

CASCADE cause the activation and repression of any targeted promoter region in P. aeruginosa ? To answer this question, the CASCADE complex was targeted to another P. aeruginosa gene called pilA , a gene responsible for twitching motility.

a 500 b 400 4386 Cas3 (Nu -) 4386 WT

400 + AcrIE2 300 + AcrIE2

300 200

200 crRNA/ crRNA/ no crRNA%) crRNA/ crRNA no %) PyocyaninExpression

Pyocyanin Pyocyanin Expression 100 phzM

100 phzM ( (

0 0

+ + - + r 3- 1 2 3 4 o A A A A A ct A 1+ N A 4+ N N N N e N NA 2+ R N R R rR R v rR r cr cr c cr ty c c p 2 E2 m E2 crR E E2 crR Empty vector E

Figure 17: AcrIE2 dependent transcriptional regulation of phzM . a, Pyocyanin expression levels measured out of P. aeruginosa cells carrying phzM promoter targeting crRNAs in the presence of AcrIE2 and the absence of an active Cas3 protein. b, Pyocyanin expression levels measured out of P. aeruginosa cells carrying phzM promoter targeting crRNAs in the presence of AcrIE2 and Cas3. c, RT-qPCR results reflecting both mRNA overexpression and repression in the context of crRNA targeting and the presence 43

of AcrIE2. E2 = AcrIE2, Cas3 (Nu-) = Cas3 nuclease knockout cells (statistics are defined by One-way ANOVA, P < 0.01, n=2). Error bars represent standard deviation.

2.4.5 AcrIE2 is responsible for CRISPR-Cas dependent transcriptional repression

and activation

To determine if AcrIE2 could alter the expression level of another gene, the transcriptional reporter assay was used on pilA , a gene responsible for twitching motility in P. aeruginosa . It is important to note that pilA contains a σ54 promoter that has a -12 and -24 segment rather than the traditional -10 and -35. As mentioned above, in the absence of Cas3 activity, the CASCADE complex is expected to bind the pilA promoter region and sterically inhibit RNA polymerase from accessing and transcribing pilA . Using a Cas3 nuclease deficient strain, the CASCADE complex was targeted to two regions of the pilA gene, one targeting the σ54 promoter (cRNA1) and the other targeting downstream of the pilA transcriptional start site (crRNA2) (Figure 18a).

As expected, CASCADE complexes targeted using both crRNA1 and crRNA2 resulted in decreased pilA mRNA production as seen through RT-qPCR, and a significant loss of twitching motility (Figure 18b, 18c).

a pilA Promoter Targets crRNA 1: -16/16 5’ CCT CTCTGTGGACT TGGCATGCAAGATGCTT AAAGAGGGCA GGCGTTAGGCCTATTC

σ54 promoter TSS pilA Start Codon crRNA 2: +41/73 3’ ATATCAACGGAGAGACACATGAAAGCTCAAAAAGGCTTCACTTTGATCGAGCTGATGA

b 10

4 4386 Cas3 (Nu -) Diameter (mm)

Figure 18: CRISPR-Cas dependent transcriptional repression of pilA . a, Schematic of the pilA promoter sequences targeted by different crRNAs in the transcriptional reporter assay. (TSS represents

the transcriptional start site). b, Twitching motility measured from P. aeruginosa cells carrying pilA promoter targeting crRNAs in the absence of both AcrIE2 and an active Cas3 protein. All targets bind the non-coding DNA strand, Cas3 (Nu-) means Cas3 nuclease deficient. c, RT-qPCR results reflecting mRNA reduction in the presence of pilA promoter targeting crRNAs and in the absence of both AcrIE2 and an active Cas3 protein. (statistics are defined by One-way ANOVA P < 0.01, n=2). Error bars represent standard deviation.

Next, AcrIE2 was expressed in the presence of a nuclease deficient Cas3, an intact CASCADE complex and each of the two crRNA targets. Expression of AcrIE2 in the presence of a

CASCADE complex targeting the pilA promoter resulted in significant pilA repression (Figure

19a, c). Expression of AcrIE2 in the presence of a CASCADE complex targeting a region downstream of the pilA transcriptional start site resulted in a recovery of pilA mRNA expression and a recovery in twitching motility (Figure 19a, c). Note, that in the Cas3 nuclease deficient strain, pilA overexpression was not observed. Interestingly, when the same experiment was conducted in the presence of a functional Cas3, a targeting complex, and AcrIE2, the crRNA targeting the promoter resulted in repression of pilA whereas the crRNA targeting downstream of the transcriptional start site resulted in robust overexpression of pilA mRNA and a recovery in twitching motility (Figure 19b, c). These data are consistent with the phzM targeting results above and display only one difference. pilA is only overexpressed when an active Cas3 is present, whereas phzM is overexpressed in the presence or absence of a functional Cas3 protein.

10 10 4386 Cas3 (Nu -) b a 4386 WT + AcrIE2 8 8 + AcrIE2

6 6

4 4 Diameter(mm) Diameter (mm)

2 2

0 0

+ 1 or ct NA e 2+ R RNA 2+ v A ctor r r ve ty E2 c E2 c Empty crRNA 1+ crRN mp E

Figure 19: AcrIE2 dependent transcriptional regulation of pilA . a, Twitching motility measured from P. aeruginosa cells carrying pilA promoter targeting crRNAs in the presence of AcrIE2 and the absence of an active Cas3 protein. b, Twitching motility measured from P. aeruginosa cells carrying pilA promoter targeting crRNAs in the presence of AcrIE2 and Cas3. All targets bind the non-coding DNA strand. c, RT-qPCR results reflecting both mRNA overexpression and repression in the context of crRNA targeting and AcrIE2 (presented in the first four bars of the graph). E2 = AcrIE2. (statistics are defined by One-way ANOVA, P < 0.01, n=2). Error bars represent standard deviation.

2.5 Discussion

Bacteria and phages have been undergoing co-evolution for billions of years. The interactions between these entities is responsible for influencing everything from environmental nutrient turnover, to the human microbiome (Koskella and Brockhurst 2014; Scanlan 2017). Therefore, understanding the individual mechanisms that allow for the interactions between phages and bacteria is essential. Unfortunately, billions of years in evolutionary time provides a plethora of interactions to study. As such, it is important to focus on individual mechanisms first, to get a clearer understanding of the full picture.

One of the most important interactions between phage and bacteria is focused on how phages infect and kill their host, and in turn, how bacteria defend against infection (Seed 2015). A large body of work has shown that to inhibit phage infection bacteria have developed several different defense mechanisms (Samson et al. 2013b). CRISPR-Cas is the only known adaptive immune system found in bacteria and archaea. There are several different CRISPR-Cas systems that differ both in sequence and in structure, but they all share a similar overall function (Koonin,

Makarova, and Zhang 2017). That is, all CRISPR-Cas systems are adaptive immune systems that use an RNA guided complex to degrade foreign DNA or RNA.

The discovery of these systems brought an interesting question to light. If these systems are adaptive, meaning they are theoretically capable of targeting any sequence in a phage genome, then we would predict that bacteria have won the evolutionary arms race against phages. An elegant answer to this question came from the discovery of phage encoded Acr proteins that could inhibit the type I-F CRISPR-Cas system of P. aeruginosa (Bondy-Denomy et al. 2013).

These relatively small proteins had the capability of inhibiting CRISPR-Cas activity and

enabling phage infection. Additional work showed that there were Acr proteins which could inhibit several different CRISPR-Cas systems, suggesting that the use of Acrs for CRISPR-Cas inhibition was widespread (Pawluk et al. 2016). Taken together, it was becoming obvious, that although CRISPR-Cas systems were a very elegant mechanism for inhibiting phage infection,

Acr proteins were an effective means to inhibit CRISPR-Cas activity. Moreover, it was also becoming clear that anti-CRISPRs in general were structurally very different with differing mechanisms of action.

AcrIE2 is one of the first anti-CRISPR proteins found to inhibit the type I-E CRISPR-Cas system of P. aeruginosa strain SMC 4386. Noting the differences between the mechanisms of action for type I-F and type I-E anti-CRISPR proteins, I wanted to know more about AcrIE2. In this work, the structure of AcrIE2 is determined, adding to the type I Acr structural database which includes structures of AcrIF1, AcrIF2, AcrIF3, AcrIF10 and AcrIE1. It was shown that the AcrIE2 structure differs significantly from every Acr protein making it a unique structural family.

Moreover, my work shows that AcrIE2 contains a functional pocket composed of residues R9,

M58, H80, and M82 that are responsible for allowing AcrIE2 to interact with the CASCADE targeting complex. The ability of AcrIE2 to interact with the complex is in stark contrast to the mechanism of CRISPR-Cas activity for AcrIE1 which interacts with the Cas3e nuclease/helicase protein. These results showed that AcrIE2 is a structurally distinct Acr protein that functions through a different mechanism of action to inhibit the type I-E system.

To understand more about the mechanism of AcrIE2 I used a transcriptional reporter assay. In this assay, I targeted the CRISPR-Cas system to promoter regions of two different genes in the presence of AcrIE2. The results suggested that promoter targeting complexes could inhibit the

transcription of a downstream gene, suggesting that the targeting complex could bind the DNA but was unable to induce DNA degradation. Complexes that targeted regions downstream of the transcriptional start site resulted in transcriptional expression and in some cases overexpression, suggesting that the complex was in some way becoming displaced from the target and allowing transcription.

It is interesting to note that the effects observed above are based on the position of CASCADE targeting. That is, targeting the promoter of a gene results in repression, and targeting downstream of the transcriptional start site results in increased transcription. Focusing on repression, it is likely that CASCADE binding deforms the promoter and inhibits promoter recognition by RNA polymerase, or simply blocks access to the promoter, resulting in gene repression (Figure 20). When focusing on the overexpression/expression phenotype, in the presence of AcrIE2 the CASCADE complex interacts with its target and unwinds the DNA helix directly downstream of the transcriptional start site (R-loop formation). This unwinding leaves the promoter of the gene relatively unperturbed resulting in regular RNA polymerase binding.

An important component of successful transcriptional initiation is the ability of RNA polymerase to unwind DNA downstream of the promoter, also called scrunching. In scrunching, RNA polymerase (RNAP) remains stationary and pulls downstream DNA into itself, this energy dependent process is followed by the transition from an RNAP initiation complex to an elongation complex (Revyakin et al. 2006) . It is possible, that because the R-loop has already unwound the downstream DNA, RNA polymerase can more easily initiate successful transcription (Figure 20). This idea, coupled with the ability of RNAP to knock off an AcrIE2 bound CASCADE complex (as shown by the reporter assay) suggests that RNA polymerase can more easily initiate successful transcription (Figure 20). These processes collectively, result in an

RNA polymerase that binds the promoter, is more likely to successfully initiate transcription and can knock off CASCADE, resulting in gene overexpression/expression.

Figure 20: AcrIE2 model of activity. In the presence of AcrIE2 a type I-E CRISPR-Cas complex that is targeted to a gene promoter can bind the promoter but is unable to recruit or permit Cas3 nuclease activity. The bound complex deforms the promoter region by unwinding the dsDNA and inhibits the recruitment of RNA polymerase, resulting in the reduction of transcription (displayed by the red x). When the type I-E CRISPR-Cas complex is targeted downstream of a transcriptional start site in the presence of AcrIE2, RNA polymerase can interact with a nearby promoter and knock the complex off during transcription, resulting in observed transcriptional expression/overexpression of the targeted gene.

The observed ability of Acr dependent CRISPR-Cas transcriptional regulation suggests that Acr proteins do more than simply inhibit CRISPR-Cas activity. This concept is further corroborated by two other Acr proteins AcrIE1 and AcrIF3 which can both induce Acr dependent CRISPR-

Cas transcriptional inhibition through their interaction with the Cas3e and Cas3f proteins respectively (Pawluk et al. 2017; Joseph Bondy-Denomy et al. 2015). It is possible that Acr proteins can be used for the transcriptional control of targeted genes, which may be beneficial for either the phage, the bacterium, or both. For example, it is possible that following DNA injection, an AcrIE2 producing phage can induce the overexpression of a CRISPR targeted phage gene required for infection. This would allow for phage survival and may improve the efficiency of infection. More research will be required to determine whether an Acr protein has been co- opted for such uses in natural systems.

Collectively, several different Acr protein families have been found to inhibit a variety of

CRISPR-Cas systems, but only a small number have been studied (Pawluk et al. 2016). Thus far, all studied Acr proteins have different mechanisms of activity. Therefore, more structural and biochemical work must be conducted to uncover other mechanisms of Acr dependent CRISPR-

Cas inhibition, providing deeper insight into the function of CRISPR-Cas systems, the function of anti-CRISPR proteins, and their evolutionary interactions.

2.6 Materials and Methods

2.6.1 Phage propagation and bacterial growth

Pseudomonas aeruginosa and Escherichia coli were propagated in lysogeny broth (LB) media or agar, unless otherwise stated. Electroporation was used to introduce the stock pHERD30t

plasmid or pHERD30t expressing AcrIE2 and its mutants into P. aeruginosa . Calcium competent cells were used for transforming E. coli. P. aeruginosa with pHERD30t was grown on LB supplemented with gentamycin 50 μg/mL and E. coli with pHERD30t was grown in 30 μg/mL. 1 mM arabinose was used to induce AcrIE2 expression from pHERD30t, unless otherwise stated.

To prepare phage stocks for JBD93a, 3 mL of 0.7 % LB top agar was inoculated with 150 μL of

O/N P. aeruginosa SMC 4386 WT cultures and 10 μL of JBD93a phage. The mixture was vortexed and plated on 40 mL of LB agar supplemented with 10 mM MgSO 4 and incubated overnight at 30 °C. Plates were then soaked with SM buffer to extract phage; the contents were centrifuged at 14,000 RPM removing cellular and agar debris. Finally, chloroform was added to the SM buffer and the phage were stored at 4 °C. To prepare JBD8 stocks, SMC 4386 Cas3 knockout cells were used instead of SMC 4386 WT.

2.6.2 Phage spotting assays

To conduct phage spotting assays 3 mL of 0.7 % LB top agar was inoculated with 150 μL of overnight P. aeruginosa SMC 4386 WT or SMC 4386 Cas3 nuclease deficient cells. The contents were then plated on 40 mL of LB agar supplemented with 10 mM MgSO 4. The plates were left to dry, and 1.5 µl aliquots of phage were spotted on top in tenfold dilutions. Observed plaques and spots were indicative of successful phage replication and bacterial lysis.

2.6.3 Lysogen formation

P. aeruginosa cells from the center of a phage infected plaque were streaked on LB agar to individual colonies. The presence of prophage was confirmed through resistance to superinfection and the subsequent production of phage from the lysogen.

2.6.4 Site-directed mutagenesis and splicing by overlap extension - polymerase

chain reaction (SOE-PCR)

To create selected mutations in AcrIE2 complementary oligonucleotides containing a mutated codon flanked by five additional codons were synthesized by Eurofins Genomics. PCR was conducted using Pfu DNA polymerase on the template pHERD30t vector containing the acrIE2 gene . The template was then digested using DpnI (New England BioLabs [NEB]). Digestion was followed by ethanol precipitation to purify and concentrate the DNA. Calcium competent E. coli

DH5α was transformed with the plasmid resuspended in H 2O and plated on gentamycin 30

μg/mL plates. acrIE2 mutations were confirmed by sequencing and plasmids were electroporated into P. aeruginosa SMC 4386 or SMC 4386 Cas3 nuclease deficient cells for phage spotting assays.

To create multiple mutations for AcrIE2, three primers were used with overhangs to each other.

These primers, when amplified using Pfu DNA polymerase resulted in successful overlap extension amplification of the full AcrIE2 gene (see below).

Each of the three primers had M (codes for an A/C nucleotide), K (codes for a G/T nucleotide),

R (codes for an A/G nucleotide), and W (codes for an A/T nucleotide) nucleotide codes placed at regions that would mutate surface exposed residues on the AcrIE2 protein. Upon overlap extension amplification, each AcrIE2 PCR fragment had a different set of surface exposed 54

residues mutated at random. The result, was a library of acrIE2 mutants that were ligated into pHERD30t, transformed into E. coli, sequenced, and finally electroporated into P. aeruginosa for the downstream phage spotting assay.

2.6.5 AcrIE2 expression in E. coli and protein purification

The AcrIE2 open reading frame was cloned by ligation into pET-21d with a C-terminal non- cleavable 6xHis tag. E. coli BL21 DE3 cells carrying the plasmid maintained in 100 µg/mL ampicillin were grown overnight at 37 °C in 10 mL LB medium and then sub-cultured into 1 L

LB medium at 37 °C. After the cells reached an OD 600 of 0.5-0.6, IPTG was added to a final concentration of 0.8 mM to induce plasmid-based protein expression. After four hours of shaking incubation at 37 °C, cells were pelleted and stored at -80 °C. Cells were resuspended in binding buffer (20 mM Tris-HCl pH 7.5, 200 mM NaCl, 5 mM β-mercaptoethanol) with 5 mM imidazole added. Cells were lysed by sonication and lysates were cleared by centrifugation for 20 min at

17000 RPM. 1 mL Qiagen Ni-NTA bead suspension was added and mixed with the cell lysate for 30 min at 4 °C. Column purification at room temperature was performed with washes in binding buffer + 30 mM imidazole, and the protein was eluted in binding buffer + 300 mM imidazole. Binding buffer was used for overnight dialysis at room temperature. The protein was further purified by size exclusion chromatography (SEC) using a Superdex 200 16/60 column

(large-scale purification) column in binding buffer.

2.6.6 Circular dichroism spectroscopy

Circular dichroism (CD) was conducted using a 0.1 cm quartz cuvette at 25 °C with a measurement range of 260 to 200 nm. The scanning speed was 60 nm/min with a bandwidth of

1nm, response time of 5 s, and data pitch of 1.0 nm. Protein concentrations for both WT and mutant His-tagged AcrIE2 were 60 µM in buffer containing 20 mM Tris-HCl, pH 7.5, 200 mM

NaCl. Triplicate measurements were recorded and averaged for each CD run. Three biological replicates were collected.

2.6.7 P. aeruginosa AcrIE2 affinity purification

Five-hundred-milliliter cultures of P. aeruginosa strain SMC 4386 containing pHERD30t expressing 6XHis-tagged AcrIE1, AcrIE2, or AcrIE2 mutants were grown at 37 °C in LB medium to an OD 600 of 0.6. At this OD, 0.8 mM arabinose was added to induce Acr expression from the plasmid. Following an additional 8 h of incubation the cells were spun down at 7500

RPM for 20 min. Cells were resuspended in binding buffer, lysed by sonication, and centrifuged with the same method described in “expression and purification in E. coli ” above. 500 µL of

Qiagen Ni-NTA bead suspension was added and mixed with the cell lysate for 30 min at 4 °C.

Column purification, and elution was the same as described above. 2 mL of the elution was trypsin digested and analyzed by mass spectrometry. The remaining material was used to analyze the elution contents via SDS-PAGE and silver stain.

2.6.8 Silver Staining

The SDS-PAGE gel was placed in overnight solution (110 mL H 2O, 80 mL ethanol 95 %, 10 mL acetic acid) for 16 h. Oxidizing solution was added (110 mL H 2O, 80 mL ethanol 95 %, 10 mL acetic acid, and 1.4 g periodic acid) for 20 min with shaking at 250 to 300 RPM. The gel was washed 5-6 times (7 min per wash) with H 2O while shaking. Staining solution was added (113 mL H 2O, 30 mL ammonium hydroxide, 560 µL of 5 N NaOH, and 1 g of silver nitrate in 5 mL

of H 2O) for 15 min with shaking. The gel was washed for 4-5 times, 5 min each, with H 2O and

finally developed with 200 mL H 2O, 0.01 g citric acid, and 130 µL of formaldehyde for 5-10 minutes. Stop solution (7.5 mL of 5 % acetic acid in 150 mL of H 2O) was added to end the

reaction.

2.6.9 Transcriptional reporter assay

As a control for crRNA activity, WT P. aeruginosa SMC 4386 or Cas3 nuclease inactive SMC

4386 cells were transformed with pHERD30t expressing a 4386 type I-E crRNA targeting either phzM or pilA . The transformation efficiency of WT P. aeruginosa SMC 4386 was significantly

decreased in the presence of each crRNA, whereas in the cas3 nuclease deficient strain

transformation efficiency was un-affected when compared to a pHERD30t transformation alone.

crRNA targets were as follows:

phzM target crRNAs

crRNA1 5’cggttcatccccacgcatgtggggaacacATCTGGTAGGTGCCAGACAGGGTATG CGGGATcggttcatccccacgcatgtggggaacac3’ crRNA2 5’cggttcatccccacgcatgtggggaacacCTGATGCTTCCTGCAATGCCGGAGGTT GTAGCcggttcatccccacgcatgtggggaacac3’ crRNA3 5’cggttcatccccacgcatgtggggaacacAATAAAATTACAACTTGGCTACAACC TCCGGCcggttcatccccacgcatgtggggaacac3’ crRNA4 5’cggttcatccccacgcatgtggggaacacTTGTAATTTTATTCTTTCGGTACGCAG GAAAAcggttcatccccacgcatgtggggaacac3’ pilA target crRNAs

crRNA1 5’cggttcatccccacgcatgtggggaacacATGCTTAAAGAGGGCAGGCGTTAGGC CTATTCcggttcatccccacgcatgtggggaacac3’ crRNA2 5’cggttcatccccacgcatgtggggaacacCTCAAAAAGGCTTCACTTTGATCGAG CTGATGcggttcatccccacgcatgtggggaacac3’ Vector

pHERD-30t Pbad promoter, Gentamycin resistance, E.coli origin, P. aeruginosa origin.

AcrIE2 amino acid MANTYLIDPRKNNDNSGERFTVDAVDITAAAKSAAQQILGEEFEGLV sequence YRETGESNGSGMFQAYHHLHGTNRTETTVGYPFHVMELAHHHHHH*

Strains used

E. coli top ten Used for cloning

P. aeruginosa SMC Transformed with sequenced plasmids for experiments 4386 P. aeruginosa SMC Transformed with sequenced plasmids for experiments 4386 (Cas3 Nu-)

To conduct the transcriptional reporter assay WT P. aeruginosa , or Cas3 (Nu-) cells were transformed with a pHERD-30t carrying plasmid expressing both AcrIE2 and a crRNA targeting different promoter regions of either the phzM gene or pilA . To assess pyocyanin production cells were plated on gentamycin 50 μg/mL plates and placed overnight at 37 °C. Individual colonies were picked and made into an overnight in King’s A media supplemented with gentamycin 50

μg/mL, cells were grown for 16 h. The next day a 1:100 dilution of cells was placed in a new 5 mL overnight of King’s A media and grown for 5 h at 37 °C. At 5 h the expression of AcrIE2 and the crRNA were induced using 5 mM arabinose. The cells were then grown for 20-24 h overnight in a 37 °C shaker.

2.6.10 Twitching motility assays

To assess twitching motility, WT P. aeruginosa SMC 4386 or Cas3 (Nu-) cells were transformed with pHERD-30t expressing both AcrIE2 and a crRNA targeting pilA. Cells were plated on gentamycin (50 μg/mL) supplemented with 5 mM arabinose and grown overnight at 37 °C.

Individual colonies were picked using a toothpick and were stabbed into the plastic base of a gentamycin 50 μg/mL LB 1 % agar thin plate supplemented with 5 mM arabinose. The plates were placed upside down in a humid container in a 37 °C incubator for 36 – 48 h. The following day the plates were soaked with 6 mL of developer solution (400 mL water, 100 mL glacial acetic acid, 500 mL methanol, store at 4 °C) for 30 min. The liquid was dispensed, and the agar

was carefully removed. The plates were dried, revealing a cloudy zone on the plastic plate representing twitching motility.

2.6.11 RT-qPCR experiments

Phenol chloroform extraction followed by gDNA degradation (1 μL of 10X ezDNase Buffer, 1

μL ezDNase enzyme, 2 μg of template RNA, topped to 10 μL with H 2O, the contents were mixed and incubated at 37 °C for 4 min) and cDNA synthesis (4 μL SuperScript IV VILO Master Mix with 6 μL of water added to the gDNA degradation mix above , incubated for 10 min at 25 °C,

10 min at 50 °C and 5 min at 85 °C) using the Thermo Scientific SuperScript IV VILO Master

Mix. The final concentration of cDNA in the RT-qPCR was 10 ng/μL, the RT-qPCR was conducted using the Thermo Fisher SYBR Green Real-Time PCR Master Mix (5 μL PowerUp

SYBR Green Master Mix with 10 ng/μL cDNA topped to 10 μL of water). The annealing temperature for all primers was 60 °C. All RT-qPCR efficiency values exceeded 85 %. The primer sets used to amplify each gene can be found below:

Housekeeping genes: rpoD : 5’ – GGGCGAAGAAGGAAATGGTC / 5’ – CAGGTGGCGTAGGTAGAGAA clpX : 5’ – CGCTTGTAGTGGTTGTATACCG / 5’ – AAAGTAGTGGGCACAAACTTCC

Target genes: pilA : 5’ – GATCGAGCTGATGATCGTTGTAG / 5’ – ACGAACGGTGTAGTCCTGATA phzM : 5’ – CTGCTGCGCGTAATTTGATAC / 5’ – TCGATCCCGCTCTCGAT

Chapter 3

CRISPR-Cas genome editing in P. aeruginosa

3.1 Acknowledgments

I would like to thank Kristina Sztanko for helping with the development of a homologous recombination dependent Cas9 genome editing method for P. aeruginosa . I would also like to thank the Marraffini lab for providing the pCas9 construct. A special thanks to Kristina, Eric, and

Veronique for testing and showing that the CRISPR-Cas9 genome editing technique developed below works on different P. aeruginosa genes and strains. This research was funded with support from the Canada Institutes of Health Research (CIHR).

3.2 Abstract

P. aeruginosa is a gram negative multidrug resistant, opportunistic pathogen that is responsible for approximately 15000 infections and 400 deaths every year (CDC 2018). In light of this, the ability to manipulate the genome of P. aeruginosa is necessary in order to understand this bacterium in the context of infection. Unfortunately, most methods used today for the manipulation of the P. aeruginosa genome tend to be laborious. In this section, I describe the development and use of a type I-E and type II-A CRISPR-Cas system for genome editing in P. aeruginosa. These methods allow for the development of random or scarless genetic modifications in the P. aeruginosa chromosome and can be used on a variety of P. aeruginosa strains. Use of these genome editing techniques will improve our ability to investigate P. aeruginosa physiology in the context of infection.

3.3 Results

3.3.1 Type I-E CRISPR-Cas genome editing methods for P. aeruginosa

As previously discussed, P. aeruginosa strain SMC 4386 harbors a naturally expressed and active type I-E CRISPR-Cas system. I wanted to determine if this system could be used for creating specific scarless mutations in the P. aeruginosa genome. Previous work with this system in our lab had shown that the type I-E CRISPR-Cas system could target the genome of SMC

4386 and cause large deletions in the genome. To be specific, targeting the type I-E CRISPR-Cas system to the Cas8 gene of P. aeruginosa resulted in a strain containing a 125 kb genomic deletion that included both P. aeruginosa CRISPR arrays and all the Cas genes. The reason for this large deletion is expected to be a function of how the type I-E system targets and cleaves

DNA. That is, upon annealing to an appropriate target the nuclease Cas3 does not form a dsDNA nick but rather nicks and continues to unwind and cleave downstream DNA using its helicase and nuclease domains respectively (Gong et al. 2014). The result is a deletion that is significantly larger than the protospacer target. To circumvent this issue, I used homologous recombination

(HR) followed by type I-E CRISPR-Cas targeting to make specific scarless mutations in the

SMC 4386 genome (Figure 21).

Figure 21: HR dependent type I-E CRISPR-Cas genome editing. Experimental layout of genome editing. The genome of 4386 is extracted. PCR is used to make a cas3 DNA fragment that is digested and ligated into the pHERD-20t plasmid. pHERD-20t undergoes site-directed mutagenesis of cas3 through PCR. The plasmid is then transformed into 4386 and HR allows for recombination of the cas3 mutation into the bacterial genome. To select for cas3 mutant colonies, the cells are transformed with a second plasmid pHERD-30t expressing a type I-E crRNA that targets the host genome. Cells with a mutant cas3 gene are not targeted. This figure was modified from Servier Medical Art, licensed under a Creative Common Attribution 3.0 Generic License. http://smart.servier.com/.

To do this, I chose to mutate the cas3 nuclease/helicase gene. Firstly, the genome of SMC 4386 was extracted, and a PCR reaction was used to amplify the entirety of the cas3 gene. Next, the cas3 gene was cloned into the pHERD-20t vector through restriction digest and ligation.

Following ligation and sequencing, the plasmid was subjected to site directed mutagenesis in order to create two different mutations, the first mutation would result in a Cas3 nuclease deficient protein (cas3 H74A/D75A), and the second would result in a Cas3 helicase deficient protein (cas3 D434A) (Gong et al. 2014; Beloglazova et al. 2011, 3; Sinkunas et al. 2011;

Pawluk et al. 2017). Once the deletions were sequenced, the plasmids were transformed into

SMC 4386 independently and the cells were plated on carbenicillin 300 μg/mL. Following the transformation, a low naturally occurring level of HR allowed some SMC 4386 cells to recombine the cas3 nuclease or helicase mutant genes into their genome. The cells were then collected and sub-cultured for three days in antibiotic free LB media to help in curing the pHERD-20t plasmid. Finally, cells were transformed with a plasmid carrying a crRNA that would target the type I-E CRISPR-Cas system to the DNA polymerase III chi subunit which is essential for P. aeruginosa survival. If the cas3 gene was active, the crRNA would target the host DNA polymerase gene resulting in cell death. If the cas3 gene coded for a nuclease or helicase deficient Cas3 protein the type I-E CRISPR-Cas system would be unable to target the host DNA polymerase and the cells would survive. As expected, upon transformation with the crRNA there was a significant reduction in colony count on a plate when compared to empty vector alone (Figure 22a). Moreover, each mutant had 50 % (2/4) of the colonies sequenced correctly. The correct mutants contained a scarless mutation resulting in two new SMC 4386 mutants, SMC 4386 ( cas3 H74A/D75A) and SMC 4386 ( cas3 D434A) (Figure 22b). Both mutants were then shown to have active CRISPR-Cas binding through the transcriptional

reporter assay and no nuclease activity through phage spotting assays. Briefly, in the reporter assay, cells are transformed with a crRNA targeting the phzM gene, this gene is responsible for the production of the blue green pigment pyocyanin. When pyocyanin is produced P. aeruginosa cultures turn green, when pyocyanin is not produced the cultures are yellow. WT P. aeruginosa cells transformed with a self-targeting crRNA against phzM target their own genome and cause cell death via Cas3 nuclease/helicase activity, but, if the nuclease or helicase domain of Cas3 is inactivated (via mutagenesis) the CASCADE complex can still bind the target but cannot induce

DNA degradation. The outcome, is a CASCADE complex that binds to phzM and sterically inhibits RNA polymerase transcription, resulting in no pyocyanin production and yellow cultures as seen in Figure 22c and d (Bondy-Denomy et al. 2015; Pawluk et al. 2017). These results suggest that although an inactive Cas3 protein was produced, all other components of the

CRISPR-Cas targeting complex were expressed and functional. Although the type I-E CRISPR-

Cas system of SMC 4386 could be used for both random and specific genome editing, it was not the best system to use for genome editing in other P. aeruginosa strains. This caveat is primarily due to the type I-E CRISPR-Cas system being composed of multiple genes. Such a system would be difficult to clone and may suffer from various expression defects. To develop a more versatile

CRISPR-Cas genome editing method for P. aeruginosa I changed my focus to the use of

CRISPR-Cas9.

Figure 22: P. aeruginosa 4386 knockout strains. a, Transformation efficiency of 4386 cells that have undergone transformation with a type I-E crRNA targeting self vs 4386 cells transformed with pHERD- 30t vector alone (EV). b, Representation of cas3 nuclease and helicase mutations sequenced from the chromosome of P. aeruginosa 4386. c, Transcriptional reporter assay of 4386 Cas3 (Nu-) (nuclease inactive) and 4386 Cas3 (He-) (helicase mutant) in the presence or absence of a crRNA targeting the promoter of phzM . In the presence of a crRNA, both Cas3 mutant proteins are inactive, but the type I-E CRISPR-Cas targeting complex is still able to bind the promoter target and inhibit RNA polymerase recruitment. The result is phzM repression and yellow cultures. d, Phage spotting assays with ten-fold dilutions of CRISPR-sensitive phage spotted on a lawn of 4386 WT, 4386 Cas3 (Nu-), and 4386 Cas3 (He-) cells. Error bars represent standard deviation.

3.4 P. aeruginosa tCas9 plasmid

Cas9 has been shown to be a versatile genome editing tool for bacteria. This is primarily because

CRISPR-Cas9 requires only one gene and one sgRNA (a crRNA-tracrRNA variant) to be expressed for efficient targeting and cleavage of a DNA sequence. Moreover, Cas9 specifically nicks dsDNA to form a blunt end double stranded break, this is in stark contrast to the Cas3 nuclease/helicase protein of the type I-E system which is known to degrade a large proportion of the DNA target. These factors make Cas9 easy to clone (one gene) and very specific in targeting

(blunt end cut). To this end, I worked to develop a Cas9 expression vector that could be used in

P. aeruginosa for the purpose of gene editing (dubbed tCas9).

To develop the tCas9 plasmid, a PCR reaction was used to amplify the type II-A S. pyogenes cas9 gene from the pCas9 plasmid designed by the Marraffini lab. This amplicon was then inserted into the multiple cloning site of pHERD-30t through ligation independent cloning. The sequenced plasmid contained an arabinose inducible promoter upstream of the cas9 gene. Site directed mutagenesis was then used to remove an NheI site found within the Cas9 gene by creating a silent mutation. This mutation was necessary as the NheI site found on the plasmid

backbone was required for ligating the sgRNA sequence. A plasmid map can be found in figure

23.

Figure 23: Plasmid map of tCas9. Cas9 is under the control of the arabinose inducible araBAD promoter. sgRNAs can be ligated into the NheI site in either direction as NheI is flanked by promoters on either side. The plasmid is compatible with E. coli (pBR322 origin) and P. aeruginosa (pRO1600 origin). Image adapted from plasmapper (Dong et al. 2004).

3.4.1 HR independent Cas9 genome editing method for P. aeruginosa

The tCas9 plasmid was used in two methods, the first was HR independent and the second was

HR dependent. Both methods mirror those previously used for the type I-E system above. For

HR independent tCas9 genome editing, the tCas9 plasmid containing a sgRNA targeting the type

I-E cas3 or cas6 genes was transformed into SMC 4386 cells, the tCas9 plasmid alone was

transformed as a control. Cells transformed with the tCas9+sgRNA plasmid targeting either cas3 or cas6 had a more than 1000-fold reduction in transformation efficiency, suggesting that Cas9 targeted the cells (Figure 24). Amplification and sequencing of 10 cas3 targeted colonies showed that 1:10 transformants contained a point mutation in the start codon of cas3, resulting in a Cas3

M1I mutation (Figure 25b), all other cells were WT. In this targeting assay I expected to create a mutation close to the start codon of cas3 as the sgRNA used in this assay targeted the RuvC and

HNH domains to the start codon of cas3 (Figure 25a). Phage spotting assays showed that this mutant was unable to target phage DNA suggesting the cas3 gene was no longer expressed correctly or that the transcript produced an inactive Cas3 protein (Figure 25b). Amplification of

10 cas6 targeted colonies showed that every colony contained a genomic deletion much larger than the cas6 gene alone (in this case the cas6 target was 45 nucleotides downstream of the cas6 start codon). Although these knockouts were not pursued further each of the cas6 knockout colonies had no detectable CRISPR-Cas activity as shown via phage spotting (Figure 26). This work showed that tCas9 was active in strain SMC 4386 and could target the SMC 4386 genome.

Unfortunately, as expected, the wide range of mutations that were observed, from point mutations to deletions suggested that HR independent tCas9 mutagenesis was not the best approach.

Figure 24: tCas9 + sgRNA transformation efficiency in SMC 4386 . Transformation efficiency of 4386 cells transformed with a type I-E crRNA against either the cas3 or cas6 gene. Error bars represent standard deviation.

Figure 25: HR independent tCas9 mutation. a, Schematic of the spacer used for Cas9 targeting of P. aeruginosa type I-E cas3 . b, Sequence result of Cas3 M1I mutation formed through direct tCas9 targeting of the cas3 gene followed by NHEJ. c, Cas3 M1I and Cas3 WT cells spotted in ten-fold dilutions with a CRISPR sensitive phage.

Figure 26: HR independent tCas9 knockout. Multiple Cas6 genomic deletions spotted in ten-fold dilutions with a CRISPR sensitive phage.

3.4.2 P. aeruginosa Cas9 genome editing with homologous recombination

To improve the specificity of mutations, HR dependent tCas9 mutagenesis was used. In this case,

P. aeruginosa strain PAO1-JBD68 lysogen was required. JBD68 is a phage that requires an interaction with the PAO1 type IV pilus (a receptor on the surface of PAO1) for adsorption.

Gene 24 encodes a protein found in the JBD68 phage particle and shares sequence similarity to a protein called Gp49, which is a pilus binding protein found in another phage (DMS3). The goal of this experiment was to make a mutation in gene 24 and assess the ability of this JBD68 mutant to infect PAO1. Although additional details of this gene are the purview of another student’s work, the knockout experiment was conducted using HR dependent Cas9 mutagenesis by myself and Kristina Sztanko (summarized in Figure 27).

Figure 27: HR dependent type II-A CRISPR-Cas genome editing. Experimental layout of genome editing. Two PCR reactions of JBD68 gp24 followed by splicing by overlap extension PCR produced a Δ1-150 gene 24 with 300 nucleotide adjacent homology arms to the genome of JBD68. This region was ligated into pHERD-20t and transformed into a PAO1-JBD68 lysogen where HR allowed for recombination of the mutant gene. To select for Δ1-150 gene 24 PAO1-JBD68 lysogens, a second plasmid, tCas9, containing an sgRNA that targets WT gene 24 was transformed. Cells with a mutant gene 24 are not targeted by tCas9. This figure was modified from Servier Medical Art, licensed under a Creative Common Attribution 3.0 Generic License. http://smart.servier.com/.

The first step required the production of two PCR amplifications of gene 24 and adjacent homology arms from JBD68. Through splicing by overlap extension (SOE) PCR, the two PCR fragments could produce a Δ1-150 gene 24 with 300 nucleotide adjacent homology arms to the genome of JBD68. The Δ1-150 gene 24 was then ligated into the pHERD-20t plasmid and transformed into a PAO1-JBD68 lysogen, the cells were plated on carbenicillin 300 μg/mL.

Following the transformation, a low naturally occurring level of HR allowed some PAO1-JBD68 cells to recombine Δ1-150 gene 24 into the prophage genome of JBD68. The cells were then collected and sub-cultured for three days in antibiotic free LB media to help in curing the pHERD-20t plasmid. Finally, cells were transformed with the tCas9 plasmid carrying a sgRNA that would only target the WT gene 24 . Upon Cas9 cleavage, only cells containing a mutation should survive when plated on gentamycin 50 μg/mL.

As expected, upon transformation with tCas9+sgRNA there was a significant decrease in transformation efficiency for PAO1 -JBD68 when compared to tCas9 alone, suggesting that tCas9 targeting was successful (Figure 28). To determine if the knockouts were correct, a DNA gel of

PCR fragments from PAO1-JBD68 on the plate was conducted. As expected, at least 1 in 10 colonies amplified fragments that were approximately 150 bp shorter than WT (Figure 29a).

Every one of the three amplicons with a perceived 150 bp deletion was sequenced correctly

(Figure 29b). To determine whether JBD68 phages carrying the Δ1-150 gene 24 could infect

PAO1, phages were extracted from PAO1-JBD68 Δ1-150 gene 24 and spotted in ten-fold dilutions on PAO1-WT cells. All mutant phages were still able to infect PAO1 , although to a lower plaquing efficiency. This data suggests that Gp24 is not solely responsible for P. aeruginosa PAO1 adsorption (Figure 29c). This work showed that the tCas9 plasmid

accompanied with HR could be used to make specific mutations in the genome of P. aeruginosa , and moreover, that it could be used to make mutations in a phage genome.

Figure 28: tCas9 + sgRNA transformation efficiency in PAO1 . Transformation efficiency of PAO1- JBD68 cells transformed with an sgRNA targeting WT gene 24 of JBD68a but not Δ1-150 gene 24 . Error bars represent standard deviation.

Figure 29: HR dependent type II-A CRISPR-Cas production of JBD68 gp24 Δ1-150 phage. a, Agarose gel of gene 24 PCR fragments from tCas9 targeted PAO1-JBD68 cells. b, Sequence result of gene 24 Δ1-150 following HR and tCas9 targeting. c, Phage spotting assay of WT-JBD68 phage and three different JBD68-Gp24 Δ1-150 phages on a lawn of PAO1 cells.

In the discussion section I will delve into the implications of these genome editing tools.

Moreover, I will also discuss the types of mutations other individuals in the lab have created using this system. Their work shows that tCas9 can be used in several different P. aeruginosa strains, to make mutations in various genes.

3.5 Discussion

One of the foundations for basic biology research is the ability to make mutations in the genome of an organism. Mutant genes, and their protein products can provide insight into the physiological activities of an organism. In addition, genetic mutations can provide insights into how organisms interact both in a symbiotic and disease-causing state. In this work, an efficient and dependable CRISPR-Cas genome editing tool for P. aeruginosa was developed.

Using the endogenous type I-E CRISPR-Cas system of P. aeruginosa strain SMC 4386 I showed that mutations in the genome of this strain can be made using either HR independent or HR dependent methods. As expected, HR independent methods that only require CRISPR-Cas targeting and NHEJ repair tend to result in random mutations that vary from point mutations to full deletions. With the use of HR and the type I-E system, I have shown that precise and scarless mutations can be made in the genome of SMC 4386 . To expand our genome editing abilities to other P. aeruginosa strains, I have also shown that the type II-A CRISPR-Cas system of S. pyogenes (called Cas9) can be used in HR independent and HR dependent methods for editing the P. aeruginosa genome. Much like the type I-E system, the results of HR independent Cas9 76

genome editing showed variability in the mutations produced, making this method of mutagenesis random. Yet, HR coupled with Cas9 selection resulted in precise scarless mutations in the P. aeruginosa genome.

The variability in mutations observed through HR independent methods is likely due to the native P. aeruginosa NHEJ system. In NHEJ a double stranded break (DSB) in P. aeruginosa causes the Ku heterodimer protein to bind two broken DNA ends and form a ring like structure.

Ku can then recruit a ligase (LigD) in order to ligate the ends together (Zhu and Shuman 2010;

Bowater and Doherty 2006). Because most DSB’s have non-complementary ends during damage, DSBs usually require additional processing from nucleases and DNA polymerase resulting in insertions and/or deletions (indel) at the cleavage site upon repair (Su et al. 2016).

Based on the mechanism of NHEJ, HR independent CRISPR-Cas mutagenesis was expected to result in a variety of different mutations. This is likely the explanation behind the large deletion observed for the cas6 gene during Cas9 targeting and the contrasting point mutation formed during cas3 gene targeting. This, in part, is the reason behind why HR dependent CRISPR-Cas mutagenesis was pursued.

Work done in the laboratory using the Cas9 genome editing tool has allowed for the specific and random mutagenesis of multiple P. aeruginosa genes in different P. aeruginosa strains (Table 5).

Table 5: P. aeruginosa Cas9 genome editing. Strain HR independent Cas9 Mutation P. aeruginosa SMC 4386 Cas3 M1I P. aeruginosa PAO1 PilZ N285fs P. aeruginosa PA14 ΔpilA Strain HR dependent Cas9 Mutation P. aeruginosa PAO1-JBD68 Gp18 Δ1-50 lysogen

P. aeruginosa PAO1-JBD68 Gp18 Δ31-100 lysogen P. aeruginosa PAO1-JBD68 Gp19 Δ1-50 lysogen P. aeruginosa PAO1-JBD68 Gp19 Δ81-381 lysogen P. aeruginosa PAO1-JBD68 Gp24 Δ1-50 lysogen P. aeruginosa PAO1-DMS3 Gp49 Δ59-222 lysogen P. aeruginosa PA14 Cas7f S70E P. aeruginosa PA14 Cas7f S73K P. aeruginosa PA14 Cas7f G236K P. aeruginosa PA14 Cas7f K238R P. aeruginosa PA14 Cas7f K239N

Collectively, this work shows that CRISPR-Cas can be used as an efficient mechanism for making mutations in P. aeruginosa .

The idea behind this genome editing technique is to improve our ability to study P. aeruginosa physiology both in nature and in disease. Editing the genome is an important step in understanding the genetic interactions within a host organism. The hope is that this work will enable the field to attain a better resolution into the physiology of P. aeruginosa .

3.6 Materials and Methods

3.6.1 P. aeruginosa competence and transformation

To make SMC 4386 competent cells, 1 mL of SMC 4386 overnight is spun down at 10000 RPM and resuspended in 1 mL of 300 mM sucrose, following resuspension the cells are spun down once more at 10000 RPM and resuspended in 300 mM sucrose X2. 100 μL of competent cells are transformed with 300 ng/μL of plasmid (or equimolar concentrations in the case of transformation efficiency assays). Cells are plated on LB agar with appropriate antibiotic. To make PAO1 or PAO1-JBD68 competent cells, 1 mL of PAO1 is spun down at 10000 RPM and

resuspended in 1 mL of 300 mM sucrose, following resuspension the cells are spun down once more at 10000 RPM and resuspended in 300 mM sucrose X2. During the final resuspension, cells are resuspended in 300 μL of sucrose. 100 μL of competent cells are transformed with 300 ng/μL of plasmid (or equimolar concentrations in the case of transformation efficiency assays).

3.6.2 Type I-E crRNAs and type II-A sgRNAs used in this work

In green are the spacer sequences. The remaining nucleotides represent repeats/ sgRNA ends/ and restriction sites.

Type I-E crRNA used for the transcriptional reporter assay

5’CGGCCCGGTTCATCCCCACGCATGTGGGGAACACCTGATGCTTCCTGCAATGCCGG

AGGTTGTAGCCGGTTCATCCCCACGCATGTGGGGAACACGGCCACCA

Type I-E CRISPR-Cas crRNA targeting DNA polymerase III chi subunit:

5’CGGCCCGGTTCATCCCCACGCATGTGGGGAACACGCCTGGCGGCAGGGCATGCAG

GTCTACCTGCACGGTTCATCCCCACGCATGTGGGGAACACGGCCACCA

Type II-A sgRNA used to produce a cas3 (M1I) mutant

5’CCCCGCTAGCTGGAAGGTGGAAGGAATGCTGTTTTAGAGCTAGAAATAGCAAGTT

AAAATAAGGCTAGTCCGTTATCAACTTGAAAAAGTGGCACCGAGTCGGTGCTTTTTT

TGCTAGCCCCC

Type II-A sgRNA used to produce a cas6 knockout

5’CCCCGCTAGCCTCCGGCATCGCGCTTCCTCGTTTTAGAGCTAGAAATAGCAAGTTA

AAATAAGGCTAGTCCGTTATCAACTTGAAAAAGTGGCACCGAGTCGGTGCTTTTTTT

GCTAGCCCCC

Type II-A sgRNA used to produce a gp24 mutant

5’CCCCGCTAGCCCGGTTGCGCTTGGCGGAACGTTTTAGAGCTAGAAATAGCAAGTT

AAAATAAGGCTAGTCCGTTATCAACTTGAAAAAGTGGCACCGAGTCGGTGCTTTTTT

TGCTAGCCCCC

3.6.3 SOE-PCR protocol

To create a gene 24 Δ1-150 gene with 300 nt of homology arms two sets of primers were used to produce two fragments – the first fragment contained 300 nt of adjacent homology arms upstream of the start codon for gene 24 , the second fragment started 150 nt downstream of the gene 24 start codon and went an additional 303 nt into gene 24 . Both fragments had overhangs that through SOE-PCR would result in a dsDNA sequence that contained the Δ1-150 gene 24 with 300 nt adjacent homology arms on either side.

3.6.4 Additional protocols

Site-directed mutagenesis, the transcriptional reporter assay, lysogen formation, and phage propagation can all be found under the materials and methods section of chapter 2.

Chapter 4 General Discussion 4.1 Summary

In summary, the first section of my work has focused on expanding our knowledge of Acr function and its interactions with the CRISPR-Cas system. My work on AcrIE2 has reasserted the idea that Acr proteins work through a variety of different mechanisms. Unlike, AcrIE1,

AcrIF1, AcrIF2, and AcrIF3, AcrIE2 is the first Acr protein capable of interacting with a DNA bound CASCADE targeting complex. This work has also shown the first occurrence of Acr dependent CRISPR-Cas transcriptional regulation, displaying both mRNA overexpression and repression phenotypes. Prior to this, only Acr dependent CRISPR-Cas transcriptional repression was observed for AcrIF3 and AcrIE1(Bondy-Denomy et al. 2015; Pawluk et al. 2017) in P. aeruginosa . Taken together this evidence suggests that Acr proteins may play a role in both

CRISPR-Cas inhibition and gene regulation.

The second section of my work was focused on developing a CRISPR-Cas genome editing tool for P. aeruginosa . With this focus, I have shown that the type I-E CRISPR-Cas system of P. aeruginosa and the type II-A CRISPR-Cas system of S. pyogenes can both be used to make genomic mutations in P. aeruginosa . I have shown that through HR independent CRISPR-Cas mechanisms, the resulting mutations are random whereas the use of HR dependent CRISPR-Cas mutagenesis allows the production of specific scar-less mutations. My work on CRISPR-Cas can be used in research to mutate P. aeruginosa genes and understand more about P. aeruginosa physiology both in nature and in the context of infection.

Noting that P. aeruginosa is a model organism other research groups have also co-opted Cas9 for genome editing and gene silencing techniques. One group has used an HR dependent Cas9 method similar to the one discussed in this thesis to make scarless mutations in the genome of P. aeruginosa (Chen et al. 2018). In this work the group first transforms P. aeruginosa cells with a plasmid expressing Cas9 and the λ-Red system (a system that helps improve the efficiency of recombination), they then induce the expression of Cas9 and the λ-Red proteins. Following induction, they co-transform a second plasmid that contains both a recombination template and an sgRNA (the sgRNA targets Cas9 to the WT gene but not the recombination template). The result was an efficient scarless mutation in the targeted gene. Much like the system developed in this thesis, two plasmids are required to provide a recombination template and Cas9 targeting with the main difference being the expression of a λ-Red system.

Another group has used a nuclease inactive Cas9 to selectively target and inhibit the expression of P. aeruginosa genes (Tan, Reisch, and Prather 2018). In this case the group integrated a nuclease deficient (dCas9) from S. pasteurianus into the genome of P. aeruginosa . Subsequently, they would transform cells with a plasmid expressing an sgRNA for a target gene, guiding dCas9 to the gene and resulting in transcriptional repression. Collectively, these works provide new tools for genome editing and gene silencing in P. aeruginosa . Tools that are essential in understanding basic P. aeruginosa biology and P. aeruginosa infection.

4.2 Future directions

4.2.1 Anti-CRISPR characterization and biotechnological tools

The continued discovery of new Acr proteins for CRISPR-Cas systems will be essential regarding the future of the CRISPR field. With the knowledge acquired thus far, Acr proteins interact with CRISPR-Cas systems in a variety of different mechanisms. Although it is likely that the initial evolutionary purpose of Acr proteins was to be simple inhibitors of CRISPR-Cas systems, it is interesting to note that some Acrs seem to have secondary functions (functions that are likely a byproduct of their mechanism for CRISPR-Cas inhibition). For example, the type I-F

Acr protein AcrIF3 and type I-E Acr protein AcrIE1 can inhibit CRISPR-Cas function but can also enable CRISPR-Cas dependent repression of a nearby targeted gene. Moreover, the type I-E

AcrIE2 protein inhibits CRISPR-Cas dependent degradation and can turn the CRISPR-Cas complex into a transcriptional regulator. With the ability of the Acr proteins to cause CRISPR-

Cas dependent transcriptional regulation there is the potential of creating new biotechnological tools. For example, genetic association studies can be conducted using AcrIE2 and the type I-E

CRISPR-Cas system of P. aeruginosa by selectively overexpressing and repressing genes expected to be in similar pathways. A more simplified biotechnological tool could employ the use of AcrIF1 or AcrIE1 to repress the expression of a targeted gene in the presence of a type I-F or type I-E system prior to making a full gene knockout. The repression could give phenotypic insights that reflect those observed in a full knockout. Collectively, these data and hypothesis suggest that the discovery of new Acr proteins targeting different CRISPR-Cas systems will likely lead to the emergence of new Acr mechanisms of action, and potentially new secondary functions that can be used for biotechnological purposes.

4.2.2 P. aeruginosa genome editing

Genome editing is an important method in molecular biology and is primarily used to understand genetic interactions within and between organisms. Therefore, it is important to develop gene editing methods that are simple and effective when studying different organisms in a research setting. Traditional methods of genome editing in bacteria have focused primarily on using HR of a drug cassette, vector based genomic transmission, and transposon-based mutagenesis. All these methods require the insertion of a cassette (in most cases an antibiotic resistance gene) into the gene of interest. The resulting insertion leads to a loss of function. Unfortunately, the presence of an antibiotic marker may impact host physiology and may alter the expression level of surrounding genes (polar effects). These caveats represent some of the main reasons for the development of new genome editing methods, methods that can result in scarless mutations and thereby minimize unwanted side effects. Noting that P. aeruginosa is a pathogenic model organism, I wanted to develop a tool to improve P. aeruginosa gene editing with the goal of efficiently creating scarless mutations in the genome. To do this, I used the CRISPR-Cas system.

In this section of my thesis I have shown that both the type I-E and type II-A CRISPR-Cas systems of P. aeruginosa and S. pyogenes respectively can be used to make both random and specific mutations in the P. aeruginosa genome. Following my work, other groups have also developed similar methods to improve our ability to edit the genome of P. aeruginosa (Chen et al. 2018). Collectively these works will allow us to understand more about P. aeruginosa physiology and infection.

Interestingly, the function of Cas9 in P. aeruginosa suggests that other biotechnological mechanisms can be used in P. aeruginosa . More specifically, it is known that a nuclease

deficient Cas9 gene can interact with a gene target and repress gene expression (Gilbert et al.

2013). Alternatively, a nuclease deficient Cas9 gene can be fused to an activation domain in order to induce the overexpression of a targeted gene (Sander and Joung 2014). It would be interesting to alter the already existing Cas9 genome editing plasmids to use in repression and overexpression assays, giving us further insight into the genetic interactions of this organism.

4.3 Closing Remarks

I am truly grateful to Alan for giving me the opportunity to work on these projects. From the start of my Ph. D. I was intrigued and interested in understanding how anti-CRISPR proteins inhibit CRISPR-Cas. I never expected for the type I-E project to develop into a biotechnological tool. I am also thankful to have worked on creating a genome editing tool for P. aeruginosa as I have watched it develop into an essential aid for everyone in our lab. As I complete this PhD, I can honestly say this journey has thoroughly prepared me for the world.

References or Bibliography (if any)

Ackermann. 2001. “Bacteriophages: Tailed.” In eLS . American Cancer Society. https://doi.org/10.1038/npg.els.0000782.

Ackermann, H.-W. 2003. “Bacteriophage Observations and Evolution.” Research in Microbiology 154 (4): 245–51. https://doi.org/10.1016/S0923-2508(03)00067-6.

Ackermann, H.-W. 2007. “5500 Phages Examined in the Electron Microscope.” Archives of Virology 152 (2): 227–43. https://doi.org/10.1007/s00705-006-0849-1.

Anders, Carolin, Ole Niewoehner, Alessia Duerst, and Martin Jinek. 2014. “Structural Basis of PAM-Dependent Target DNA Recognition by the Cas9 Endonuclease.” Nature 513 (7519): 569–73. https://doi.org/10.1038/nature13579.

Avery, Oswald T., Colin M. MacLeod, and Maclyn McCarty. 1944. “STUDIES ON THE CHEMICAL NATURE OF THE SUBSTANCE INDUCING TRANSFORMATION OF PNEUMOCOCCAL TYPES.” The Journal of Experimental Medicine 79 (2): 137–58.

Avrani, Sarit, Omri Wurtzel, Itai Sharon, Rotem Sorek, and Debbie Lindell. 2011. “Genomic Island Variability Facilitates Prochlorococcus –virus Coexistence.” Nature 474 (7353): 604–8. https://doi.org/10.1038/nature10172.

Barrangou, Rodolphe, Christophe Fremaux, Hélène Deveau, Melissa Richards, Patrick Boyaval, Sylvain Moineau, Dennis A. Romero, and Philippe Horvath. 2007. “CRISPR Provides Acquired Resistance against Viruses in Prokaryotes.” Science (New York, N.Y.) 315 (5819): 1709–12. https://doi.org/10.1126/science.1138140.

Barrangou, Rodolphe, and Luciano A. Marraffini. 2014. “CRISPR-Cas Systems: Prokaryotes Upgrade to Adaptive Immunity.” Molecular Cell 54 (2): 234–44. https://doi.org/10.1016/j.molcel.2014.03.011.

Beloglazova, Natalia, Pierre Petit, Robert Flick, Greg Brown, Alexei Savchenko, and Alexander F Yakunin. 2011. “Structure and Activity of the Cas3 HD Nuclease MJ0384, an Effector Enzyme of the CRISPR Interference.” The EMBO Journal 30 (22): 4616–27. https://doi.org/10.1038/emboj.2011.377.

Bondy-Denomy, Joe, April Pawluk, Karen L. Maxwell, and Alan R. Davidson. 2013. “Bacteriophage Genes That Inactivate the CRISPR/Cas Bacterial Immune System.” Nature 493 (7432): 429. https://doi.org/10.1038/nature11723.

Bondy-Denomy, Joseph, Bianca Garcia, Scott Strum, Mingjian Du, MaryClare F. Rollins, Yurima Hidalgo-Reyes, Blake Wiedenheft, Karen L. Maxwell, and Alan R. Davidson. 2015. “Multiple Mechanisms for CRISPR–Cas Inhibition by Anti-CRISPR Proteins.” Nature 526 (7571): 136. https://doi.org/10.1038/nature15254.

Bowater, Richard, and Aidan J. Doherty. 2006. “Making Ends Meet: Repairing Breaks in Bacterial DNA by Non-Homologous End-Joining.” PLOS Genetics 2 (2): e8. https://doi.org/10.1371/journal.pgen.0020008.

Brouns, Stan J. J., Matthijs M. Jore, Magnus Lundgren, Edze R. Westra, Rik J. H. Slijkhuis, Ambrosius P. L. Snijders, Mark J. Dickman, Kira S. Makarova, Eugene V. Koonin, and John van der Oost. 2008. “Small CRISPR RNAs Guide Antiviral Defense in Prokaryotes.” Science (New York, N.Y.) 321 (5891): 960–64. https://doi.org/10.1126/science.1159689.

Cady, Kyle C., Joe Bondy-Denomy, Gary E. Heussler, Alan R. Davidson, and George A. O’Toole. 2012. “The CRISPR/Cas Adaptive Immune System of Pseudomonas Aeruginosa Mediates Resistance to Naturally Occurring and Engineered Phages.” Journal of Bacteriology 194 (21): 5728–38. https://doi.org/10.1128/JB.01184-12.

Carte, Jason, Neil T. Pfister, Mark M. Compton, Rebecca M. Terns, and Michael P. Terns. 2010. “Binding and Cleavage of CRISPR RNA by Cas6.” RNA (New York, N.Y.) 16 (11): 2181–88. https://doi.org/10.1261/rna.2230110.

CDC. 2018. “The Biggest Antibiotic-Resistant Threats in the U.S.” Centers for Disease Control and Prevention. November 26, 2018. https://www.cdc.gov/drugresistance/biggest_threats.html.

Chayot, Romain, Benjamin Montagne, Didier Mazel, and Miria Ricchetti. 2010. “An End- Joining Repair Mechanism in Escherichia Coli.” Proceedings of the National Academy of Sciences 107 (5): 2141–46. https://doi.org/10.1073/pnas.0906355107.

Chen, Weizhong, Ya Zhang, Yifei Zhang, Yishuang Pi, Tongnian Gu, Liqiang Song, Yu Wang, and Quanjiang Ji. 2018. “CRISPR/Cas9-Based Genome Editing in Pseudomonas Aeruginosa and Cytidine Deaminase-Mediated Base Editing in Pseudomonas Species.” iScience 6 (August): 222–31. https://doi.org/10.1016/j.isci.2018.07.024.

Chibani, Cynthia Maria, Anton Farr, Sandra Klama, Sascha Dietrich, and Heiko Liesegang. 2019. “Classifying the Unclassified: A Phage Classification Method.” Viruses 11 (2). https://doi.org/10.3390/v11020195.

Chopin, Marie-Christine, Alain Chopin, and Elena Bidnenko. 2005. “Phage Abortive Infection in Lactococci: Variations on a Theme.” Current Opinion in Microbiology , Host--microbe interactions: fungi / edited by Howard Bussey · Host--microbe interactions: parasites / edited by Artur Scherf · Host--microbe interactions: viruses / edited by Margaret CM Smith, 8 (4): 473–79. https://doi.org/10.1016/j.mib.2005.06.006.

Clokie, Martha RJ, Andrew D Millard, Andrey V Letarov, and Shaun Heaphy. 2011. “Phages in Nature.” Bacteriophage 1 (1): 31–45. https://doi.org/10.4161/bact.1.1.14942.

Daubin, Vincent, and Gergely J. Szöllősi. 2016. “Horizontal Gene Transfer and the History of Life.” Cold Spring Harbor Perspectives in Biology 8 (4). https://doi.org/10.1101/cshperspect.a018036. 87

Dong, Xiaoli, Paul Stothard, Ian J. Forsythe, and David S. Wishart. 2004. “PlasMapper: A Web Server for Drawing and Auto-Annotating Plasmid Maps.” Nucleic Acids Research 32 (Web Server issue): W660–64. https://doi.org/10.1093/nar/gkh410.

Doolittle, W. F. 1999. “Lateral Genomics.” Trends in Cell Biology 9 (12): M5-8.

Drake, J. W. 1991. “A Constant Rate of Spontaneous Mutation in DNA-Based Microbes.” Proceedings of the National Academy of Sciences 88 (16): 7160–64. https://doi.org/10.1073/pnas.88.16.7160.

Fokine, Andrei, and Michael G Rossmann. 2014. “Molecular Architecture of Tailed Double- Stranded DNA Phages.” Bacteriophage 4 (February). https://doi.org/10.4161/bact.28281.

Garcia-Vallvé, Santiago, Anton Romeu, and Jaume Palau. 2000. “Horizontal Gene Transfer in Bacterial and Archaeal Complete Genomes.” Genome Research 10 (11): 1719–25.

Garneau, Josiane E., Marie-Ève Dupuis, Manuela Villion, Dennis A. Romero, Rodolphe Barrangou, Patrick Boyaval, Christophe Fremaux, Philippe Horvath, Alfonso H. Magadán, and Sylvain Moineau. 2010. “The CRISPR/Cas Bacterial Immune System Cleaves Bacteriophage and Plasmid DNA.” Nature 468 (7320): 67–71. https://doi.org/10.1038/nature09523.

Gesner, Emily M., Matthew J. Schellenberg, Erin L. Garside, Mark M. George, and Andrew M. Macmillan. 2011. “Recognition and Maturation of Effector RNAs in a CRISPR Interference Pathway.” Nature Structural & Molecular Biology 18 (6): 688–92. https://doi.org/10.1038/nsmb.2042.

Gilbert, Luke A., Matthew H. Larson, Leonardo Morsut, Zairan Liu, Gloria A. Brar, Sandra E. Torres, Noam Stern-Ginossar, et al. 2013. “CRISPR-Mediated Modular RNA-Guided Regulation of Transcription in Eukaryotes.” Cell 154 (2): 442–51. https://doi.org/10.1016/j.cell.2013.06.044.

Gleditzsch, Daniel, Patrick Pausch, Hanna Müller-Esparza, Ahsen Özcan, Xiaohan Guo, Gert Bange, and Lennart Randau. 2018. “PAM Identification by CRISPR-Cas Effector Complexes: Diversified Mechanisms and Structures.” RNA Biology 0 (0): 1–14. https://doi.org/10.1080/15476286.2018.1504546.

Gong, Bei, Minsang Shin, Jiali Sun, Che-Hun Jung, Edward L. Bolt, John van der Oost, and Jeong-Sun Kim. 2014. “Molecular Insights into DNA Interference by CRISPR- Associated Nuclease-Helicase Cas3.” Proceedings of the National Academy of Sciences of the United States of America 111 (46): 16359–64. https://doi.org/10.1073/pnas.1410806111.

Griffith, Fred. 1928. “The Significance of Pneumococcal Types.” The Journal of Hygiene 27 (2): 113.

Guo, Tai Wei, Alberto Bartesaghi, Hui Yang, Veronica Falconieri, Prashant Rao, Alan Merk, Edward T. Eng, et al. 2017. “Cryo-EM Structures Reveal Mechanism and Inhibition of 88

DNA Targeting by a CRISPR-Cas Surveillance Complex.” Cell 171 (2): 414–426.e12. https://doi.org/10.1016/j.cell.2017.09.006.

Hale, Caryn R., Peng Zhao, Sara Olson, Michael O. Duff, Brenton R. Graveley, Lance Wells, Rebecca M. Terns, and Michael P. Terns. 2009. “RNA-Guided RNA Cleavage by a CRISPR RNA-Cas Protein Complex.” Cell 139 (5): 945–56. https://doi.org/10.1016/j.cell.2009.07.040.

Haurwitz, Rachel E., Martin Jinek, Blake Wiedenheft, Kaihong Zhou, and Jennifer A. Doudna. 2010. “Sequence- and Structure-Specific RNA Processing by a CRISPR Endonuclease.” Science (New York, N.Y.) 329 (5997): 1355–58. https://doi.org/10.1126/science.1192272.

Herelle, MF d’. 1917. “On an Invisible Microbe Antagonistic to Dysentery Bacilli.” Bacteriophage 1 (1): 3–5. https://doi.org/10.4161/bact.1.1.14941.

Hershey, A. D., and M. Chase. 1952. “Independent Functions of Viral Protein and Nucleic Acid in Growth of Bacteriophage.” The Journal of General Physiology 36 (1): 39–56.

Hmelo, Laura R., Bradley R. Borlee, Henrik Almblad, Michelle E. Love, Trevor E. Randall, Boo Shan Tseng, Chuyang Lin, et al. 2015. “Precision-Engineering the Pseudomonas Aeruginosa Genome with Two-Step Allelic Exchange.” Nature Protocols 10 (11): 1820– 41. https://doi.org/10.1038/nprot.2015.115.

Hoang, T. T., R. R. Karkhoff-Schweizer, A. J. Kutchma, and H. P. Schweizer. 1998. “A Broad- Host-Range Flp-FRT Recombination System for Site-Specific Excision of Chromosomally-Located DNA Sequences: Application for Isolation of Unmarked Pseudomonas Aeruginosa Mutants.” Gene 212 (1): 77–86.

Holm, Liisa, and Päivi Rosenström. 2010. “Dali Server: Conservation Mapping in 3D.” Nucleic Acids Research 38 (Web Server issue): W545–49. https://doi.org/10.1093/nar/gkq366.

Huang, Jiaofang, Yuquan Xu, Hongyan Zhang, Yaqian Li, Xianqing Huang, Bin Ren, and Xuehong Zhang. 2009. “Temperature-Dependent Expression of phzM and Its Regulatory Genes lasI and ptsP in Rhizosphere Isolate Pseudomonas Sp. Strain M18.” Applied and Environmental Microbiology 75 (20): 6568–80. https://doi.org/10.1128/AEM.01148-09.

Hyman, Paul, and Stephen T. Abedon. 2010. “Chapter 7 - Bacteriophage Host Range and Bacterial Resistance.” In Advances in Applied Microbiology , 70:217–48. Advances in Applied Microbiology. Academic Press. https://doi.org/10.1016/S0065-2164(10)70007-1.

Jackson, Ryan N., Sarah M. Golden, Paul B. G. van Erp, Joshua Carter, Edze R. Westra, Stan J. J. Brouns, John van der Oost, Thomas C. Terwilliger, Randy J. Read, and Blake Wiedenheft. 2014. “Structural Biology. Crystal Structure of the CRISPR RNA-Guided Surveillance Complex from Escherichia Coli.” Science (New York, N.Y.) 345 (6203): 1473–79. https://doi.org/10.1126/science.1256328.

Jain, R., M. C. Rivera, and J. A. Lake. 1999. “Horizontal Gene Transfer among Genomes: The Complexity Hypothesis.” Proceedings of the National Academy of Sciences of the United States of America 96 (7): 3801–6.

Jiang, Wenyan, David Bikard, David Cox, Feng Zhang, and Luciano A. Marraffini. 2013. “CRISPR-Assisted Editing of Bacterial Genomes.” Nature Biotechnology 31 (3): 233–39. https://doi.org/10.1038/nbt.2508.

Jiang, Yu, Biao Chen, Chunlan Duan, Bingbing Sun, Junjie Yang, and Sheng Yang. 2015. “Multigene Editing in the Escherichia Coli Genome via the CRISPR-Cas9 System.” Appl. Environ. Microbiol. 81 (7): 2506–14. https://doi.org/10.1128/AEM.04023-14.

Keen, Eric C. 2015. “A Century of Phage Research: Bacteriophages and the Shaping of Modern Biology.” BioEssays : News and Reviews in Molecular, Cellular and Developmental Biology 37 (1): 6. https://doi.org/10.1002/bies.201400152.

Kimura, Motoo, and Takeo Maruyama. 1969. “The Substitutional Load in a Finite population 1.” Heredity 24 (1): 101–14. https://doi.org/10.1038/hdy.1969.10.

Kiro, Ruth, Dror Shitrit, and Udi Qimron. 2014. “Efficient Engineering of a Bacteriophage Genome Using the Type I-E CRISPR-Cas System.” RNA Biology 11 (1): 42–44. https://doi.org/10.4161/rna.27766.

Koonin, Eugene V., Kira S. Makarova, and Feng Zhang. 2017. “Diversity, Classification and Evolution of CRISPR-Cas Systems.” Current Opinion in Microbiology 37 (June): 67–78. https://doi.org/10.1016/j.mib.2017.05.008.

Koskella, Britt, and Michael A Brockhurst. 2014. “Bacteria–phage Coevolution as a Driver of Ecological and Evolutionary Processes in Microbial Communities.” Fems Microbiology Reviews 38 (5): 916–31. https://doi.org/10.1111/1574-6976.12072.

Kulasekara, Hemantha D. 2014. “Transposon Mutagenesis.” Methods in Molecular Biology (Clifton, N.J.) 1149: 501–19. https://doi.org/10.1007/978-1-4939-0473-0_39.

Levy, Asaf, Moran G. Goren, Ido Yosef, Oren Auster, Miriam Manor, Gil Amitai, Rotem Edgar, Udi Qimron, and Rotem Sorek. 2015. “CRISPR Adaptation Biases Explain Preference for Acquisition of Foreign DNA.” Nature 520 (7548): 505–10. https://doi.org/10.1038/nature14302.

Luo, Michelle L., Adam S. Mullis, Ryan T. Leenay, and Chase L. Beisel. 2015. “Repurposing Endogenous Type I CRISPR-Cas Systems for Programmable Gene Repression.” Nucleic Acids Research 43 (1): 674–81. https://doi.org/10.1093/nar/gku971.

Lynch, Michael, and John S. Conery. 2000. “The Evolutionary Fate and Consequences of Duplicate Genes.” Science 290 (5494): 1151–55. https://doi.org/10.1126/science.290.5494.1151.

Lynch, Michael, and Allan Force. 2000. “The Probability of Duplicate Gene Preservation by Subfunctionalization.” Genetics 154 (1): 459–73.

Marraffini, Luciano A., and Erik J. Sontheimer. 2008. “CRISPR Interference Limits Horizontal Gene Transfer in Staphylococci by Targeting DNA.” Science (New York, N.Y.) 322 (5909): 1843–45. https://doi.org/10.1126/science.1165771.

Maxwell, Karen L., Bianca Garcia, Joseph Bondy-Denomy, Diane Bona, Yurima Hidalgo- Reyes, and Alan R. Davidson. 2016. “The Solution Structure of an Anti-CRISPR Protein.” Nature Communications 7 (October): 13134. https://doi.org/10.1038/ncomms13134.

Molineux, Ian J., and Debabrata Panja. 2013. “Popping the Cork: Mechanisms of Phage Genome Ejection.” Nature Reviews Microbiology 11 (3): 194–204. https://doi.org/10.1038/nrmicro2988.

Mulepati, Sabin, and Scott Bailey. 2011. “Structural and Biochemical Analysis of Nuclease Domain of Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)- Associated Protein 3 (Cas3).” The Journal of Biological Chemistry 286 (36): 31896–903. https://doi.org/10.1074/jbc.M111.270017.

Mulepati, Sabin, Annie Héroux, and Scott Bailey. 2014. “Structural Biology. Crystal Structure of a CRISPR RNA-Guided Surveillance Complex Bound to a ssDNA Target.” Science (New York, N.Y.) 345 (6203): 1479–84. https://doi.org/10.1126/science.1256996.

Ng, Henry, and Neta Dean. 2017. “Dramatic Improvement of CRISPR/Cas9 Editing in Candida Albicans by Increased Single Guide RNA Expression.” mSphere 2 (2): e00385-16. https://doi.org/10.1128/mSphere.00385-16.

Nishimasu, Hiroshi, F. Ann Ran, Patrick D. Hsu, Silvana Konermann, Soraya I. Shehata, Naoshi Dohmae, Ryuichiro Ishitani, Feng Zhang, and Osamu Nureki. 2014. “Crystal Structure of Cas9 in Complex with Guide RNA and Target DNA.” Cell 156 (5): 935–49. https://doi.org/10.1016/j.cell.2014.02.001.

Nuñez, James K., Philip J. Kranzusch, Jonas Noeske, Addison V. Wright, Christopher W. Davies, and Jennifer A. Doudna. 2014. “Cas1-Cas2 Complex Formation Mediates Spacer Acquisition during CRISPR-Cas Adaptive Immunity.” Nature Structural & Molecular Biology 21 (6): 528–34. https://doi.org/10.1038/nsmb.2820.

Pawluk, April, Joseph Bondy-Denomy, Vivian H. W. Cheung, Karen L. Maxwell, and Alan R. Davidson. 2014. “A New Group of Phage Anti-CRISPR Genes Inhibits the Type I-E CRISPR-Cas System of Pseudomonas Aeruginosa.” mBio 5 (2): e00896-14. https://doi.org/10.1128/mBio.00896-14.

Pawluk, April, Megha Shah, Marios Mejdani, Charles Calmettes, Trevor F. Moraes, Alan R. Davidson, and Karen L. Maxwell. 2017. “Disabling a Type I-E CRISPR-Cas Nuclease with a Bacteriophage-Encoded Anti-CRISPR Protein.” mBio 8 (6). https://doi.org/10.1128/mBio.01751-17. 91

Pawluk, April, Raymond H. J. Staals, Corinda Taylor, Bridget N. J. Watson, Senjuti Saha, Peter C. Fineran, Karen L. Maxwell, and Alan R. Davidson. 2016a. “Inactivation of CRISPR- Cas Systems by Anti-CRISPR Proteins in Diverse Bacterial Species.” Nature Microbiology 1 (8): 16085. https://doi.org/10.1038/nmicrobiol.2016.85.

———. 2016b. “Inactivation of CRISPR-Cas Systems by Anti-CRISPR Proteins in Diverse Bacterial Species.” Nature Microbiology 1 (8): 16085. https://doi.org/10.1038/nmicrobiol.2016.85.

Peng, Ruchao, Ying Xu, Tengfei Zhu, Ningning Li, Jianxun Qi, Yan Chai, Min Wu, et al. 2017. “Alternate Binding Modes of Anti-CRISPR Viral Suppressors AcrF1/2 to Csy Surveillance Complex Revealed by Cryo-EM Structures.” Cell Research 27 (7): 853–64. https://doi.org/10.1038/cr.2017.79.

Przybilski, Rita, Corinna Richter, Tamzin Gristwood, James S. Clulow, Reuben B. Vercoe, and Peter C. Fineran. 2011. “Csy4 Is Responsible for CRISPR RNA Processing in Pectobacterium Atrosepticum.” RNA Biology 8 (3): 517–28. https://doi.org/10.4161/rna.8.3.15190.

Revyakin, Andrey, Chenyu Liu, Richard H. Ebright, and Terence R. Strick. 2006. “Abortive Initiation and Productive Initiation by RNA Polymerase Involve DNA Scrunching.” Science 314 (5802): 1139–43. https://doi.org/10.1126/science.1131398.

Richter, Corinna, and Peter C. Fineran. 2013. “The Subtype I-F CRISPR-Cas System Influences Pathogenicity Island Retention in Pectobacterium Atrosepticum via crRNA Generation and Csy Complex Formation.” Biochemical Society Transactions 41 (6): 1468–74. https://doi.org/10.1042/BST20130151.

Rosche, W. A., and P. L. Foster. 2000. “Determining Mutation Rates in Bacterial Populations.” Methods (San Diego, Calif.) 20 (1): 4–17. https://doi.org/10.1006/meth.1999.0901.

Ryan, Gillian L., and Andrew D. Rutenberg. 2007. “Clocking Out: Modeling Phage-Induced Lysis of Escherichia Coli.” Journal of Bacteriology 189 (13): 4749–55. https://doi.org/10.1128/JB.00392-07.

Samson, Julie E., Alfonso H. Magadán, Mourad Sabri, and Sylvain Moineau. 2013a. “Revenge of the Phages: Defeating Bacterial Defences.” Nature Reviews. Microbiology 11 (10): 675–87. https://doi.org/10.1038/nrmicro3096.

———. 2013b. “Revenge of the Phages: Defeating Bacterial Defences.” Nature Reviews Microbiology 11 (10): 675–87. https://doi.org/10.1038/nrmicro3096.

Sander, Jeffry D., and J. Keith Joung. 2014. “CRISPR-Cas Systems for Genome Editing, Regulation and Targeting.” Nature Biotechnology 32 (4): 347–55. https://doi.org/10.1038/nbt.2842.

Savitskaya, Ekaterina, Ekaterina Semenova, Vladimir Dedkov, Anastasia Metlitskaya, and Konstantin Severinov. 2013. “High-Throughput Analysis of Type I-E CRISPR/Cas 92

Spacer Acquisition in E. Coli.” RNA Biology 10 (5): 716–25. https://doi.org/10.4161/rna.24325.

Scanlan, Pauline D. 2017. “Bacteria-Bacteriophage Coevolution in the Human Gut: Implications for Microbial Diversity and Functionality.” Trends in Microbiology 25 (8): 614–23. https://doi.org/10.1016/j.tim.2017.02.012.

Schneider, Christine L. 2017. “Bacteriophage-Mediated Horizontal Gene Transfer: Transduction.” In Bacteriophages: Biology, Technology, Therapy , edited by David Harper, Stephen Abedon, Benjamin Burrowes, and Malcolm McConville, 1–42. Cham: Springer International Publishing. https://doi.org/10.1007/978-3-319-40598-8_4-1.

Seed, Kimberley D. 2015. “Battling Phages: How Bacteria Defend against Viral Attack.” PLOS Pathogens 11 (6): e1004847. https://doi.org/10.1371/journal.ppat.1004847.

Selle, Kurt, and Rodolphe Barrangou. 2015. “Harnessing CRISPR-Cas Systems for Bacterial Genome Editing.” Trends in Microbiology 23 (4): 225–32. https://doi.org/10.1016/j.tim.2015.01.008.

Semenova, Ekaterina, Matthijs M. Jore, Kirill A. Datsenko, Anna Semenova, Edze R. Westra, Barry Wanner, John van der Oost, Stan J. J. Brouns, and Konstantin Severinov. 2011. “Interference by Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR) RNA Is Governed by a Seed Sequence.” Proceedings of the National Academy of Sciences of the United States of America 108 (25): 10098–103. https://doi.org/10.1073/pnas.1104144108.

Serwer, Philip. 2007. “Evolution and the Complexity of Bacteriophages.” Virology Journal 4: 30. https://doi.org/10.1186/1743-422X-4-30.

Shuman, Stewart, and Michael S. Glickman. 2007. “Bacterial DNA Repair by Non-Homologous End Joining.” Nature Reviews Microbiology 5 (11): 852. https://doi.org/10.1038/nrmicro1768.

Sime-Ngando, Télesphore. 2014. “Environmental Bacteriophages: Viruses of Microbes in Aquatic Ecosystems.” Frontiers in Microbiology 5 (July). https://doi.org/10.3389/fmicb.2014.00355.

Sinkunas, Tomas, Giedrius Gasiunas, Christophe Fremaux, Rodolphe Barrangou, Philippe Horvath, and Virginijus Siksnys. 2011. “Cas3 Is a Single-Stranded DNA Nuclease and ATP-Dependent Helicase in the CRISPR/Cas Immune System.” The EMBO Journal 30 (7): 1335–42. https://doi.org/10.1038/emboj.2011.41.

Sinkunas, Tomas, Giedrius Gasiunas, Sakharam P. Waghmare, Mark J. Dickman, Rodolphe Barrangou, Philippe Horvath, and Virginijus Siksnys. 2013. “ In Vitro Reconstitution of CASCADE-Mediated CRISPR Immunity in Streptococcus Thermophilus.” The EMBO Journal 32 (3): 385–94. https://doi.org/10.1038/emboj.2012.352.

Su, Tianyuan, Fapeng Liu, Pengfei Gu, Haiying Jin, Yizhao Chang, Qian Wang, Quanfeng Liang, and Qingsheng Qi. 2016. “A CRISPR-Cas9 Assisted Non-Homologous End- Joining Strategy for One-Step Engineering of Bacterial Genome.” Scientific Reports 6 (November): 37895. https://doi.org/10.1038/srep37895.

Szczelkun, Mark D., Maria S. Tikhomirova, Tomas Sinkunas, Giedrius Gasiunas, Tautvydas Karvelis, Patrizia Pschera, Virginijus Siksnys, and Ralf Seidel. 2014. “Direct Observation of R-Loop Formation by Single RNA-Guided Cas9 and CASCADE Effector Complexes.” Proceedings of the National Academy of Sciences of the United States of America 111 (27): 9798–9803. https://doi.org/10.1073/pnas.1402597111.

Tan, Sue Zanne, Christopher R. Reisch, and Kristala L. J. Prather. 2018. “A Robust CRISPR Interference Gene Repression System in Pseudomonas.” Journal of Bacteriology 200 (7). https://doi.org/10.1128/JB.00575-17.

Trieu-Cuot, P., C. Carlier, P. Martin, and P. Courvalin. 1987. “Plasmid Transfer by Conjugation from Escherichia Coli to Gram-Positive Bacteria.” FEMS Microbiology Letters 48 (1–2): 289–94. https://doi.org/10.1111/j.1574-6968.1987.tb02558.x.

Twort, FW. 1915. “An Investigation on the Nature of Ultra-Microscopic Viruses.” Bacteriophage 1 (3): 127–29. https://doi.org/10.4161/bact.1.3.16737.

Wang, I. N., D. L. Smith, and R. Young. 2000. “Holins: The Protein Clocks of Bacteriophage Infections.” Annual Review of Microbiology 54: 799–825. https://doi.org/10.1146/annurev.micro.54.1.799.

Wang, Ruiying, Gan Preamplume, Michael P. Terns, Rebecca M. Terns, and Hong Li. 2011. “Interaction of the Cas6 Riboendonuclease with CRISPR RNAs: Recognition and Cleavage.” Structure (London, England : 1993) 19 (2): 257–64. https://doi.org/10.1016/j.str.2010.11.014.

Westra, Edze R., Paul B. G. van Erp, Tim Künne, Shi Pey Wong, Raymond H. J. Staals, Christel L. C. Seegers, Sander Bollen, et al. 2012. “CRISPR Immunity Relies on the Consecutive Binding and Degradation of Negatively Supercoiled Invader DNA by CASCADE and Cas3.” Molecular Cell 46 (5): 595–605. https://doi.org/10.1016/j.molcel.2012.03.018.

Wiedenheft, Blake, Esther van Duijn, Jelle B. Bultema, Jelle Bultema, Sakharam P. Waghmare, Sakharam Waghmare, Kaihong Zhou, et al. 2011. “RNA-Guided Complex from a Bacterial Immune System Enhances Target Recognition through Seed Sequence Interactions.” Proceedings of the National Academy of Sciences of the United States of America 108 (25): 10092–97. https://doi.org/10.1073/pnas.1102716108.

Wiedenheft, Blake, Gabriel C. Lander, Kaihong Zhou, Matthijs M. Jore, Stan J. J. Brouns, John van der Oost, Jennifer A. Doudna, and Eva Nogales. 2011. “Structures of the RNA- Guided Surveillance Complex from a Bacterial Immune System.” Nature 477 (7365): 486–89. https://doi.org/10.1038/nature10402.

Wiedenheft, Blake, Kaihong Zhou, Martin Jinek, Scott M. Coyle, Wendy Ma, and Jennifer A. Doudna. 2009. “Structural Basis for DNase Activity of a Conserved Protein Implicated in CRISPR-Mediated Genome Defense.” Structure (London, England: 1993) 17 (6): 904– 12. https://doi.org/10.1016/j.str.2009.03.019.

Wurtzel, Omri, Deborah R. Yoder-Himes, Kook Han, Ajai A. Dandekar, Sarit Edelheit, E. Peter Greenberg, Rotem Sorek, and Stephen Lory. 2012. “The Single-Nucleotide Resolution Transcriptome of Pseudomonas Aeruginosa Grown in Body Temperature.” PLOS Pathogens 8 (9): e1002945. https://doi.org/10.1371/journal.ppat.1002945.

Xiao, Yibei, Min Luo, Robert P. Hayes, Jonathan Kim, Sherwin Ng, Fang Ding, Maofu Liao, and Ailong Ke. 2017. “Structure Basis for Directional R-Loop Formation and Substrate Handover Mechanisms in Type I CRISPR-Cas System.” Cell 170 (1): 48–60.e11. https://doi.org/10.1016/j.cell.2017.06.012.

Zhang, Jing, Christophe Rouillon, Melina Kerou, Judith Reeks, Kim Brugger, Shirley Graham, Julia Reimann, et al. 2012. “Structure and Mechanism of the CMR Complex for CRISPR-Mediated Antiviral Immunity.” Molecular Cell 45 (3): 303–13. https://doi.org/10.1016/j.molcel.2011.12.013.

Zhu, Hui, and Stewart Shuman. 2008. “Bacterial Nonhomologous End Joining Ligases Preferentially Seal Breaks with a 3′-OH Monoribonucleotide.” The Journal of Biological Chemistry 283 (13): 8331–39. https://doi.org/10.1074/jbc.M705476200.

———. 2010. “Gap Filling Activities of Pseudomonas DNA Ligase D (LigD) Polymerase and Functional Interactions of LigD with the DNA End-Binding Ku Protein.” The Journal of Biological Chemistry 285 (7): 4815–25. https://doi.org/10.1074/jbc.M109.073874.

Zhu, Yuwei, Fan Zhang, and Zhiwei Huang. 2018. “Structural Insights into the Inactivation of CRISPR-Cas Systems by Diverse Anti-CRISPR Proteins.” BMC Biology 16 (March). https://doi.org/10.1186/s12915-018-0504-9.

Zinder, Norton D., and Joshua Lederberg. 1952. “GENETIC EXCHANGE IN SALMONELLA1.” Journal of Bacteriology 64 (5): 679–99.

Appendices (if any)