Examining the Role of Non-Canonical NOD-Like Receptors and Inflammasomes in Inflammation and Disease

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2018-03-21 Examining the Role of Non-Canonical NOD-like Receptors and Inflammasomes in Inflammation and Disease

Platnich, Jaye Matthew

Platnich, J. M. (2018). Examining the Role of Non-Canonical NOD-like Receptors and Inflammasomes in Inflammation and Disease (Unpublished doctoral thesis). University of Calgary, Calgary, AB. doi:10.11575/PRISM/31757 http://hdl.handle.net/1880/106465 doctoral thesis

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Examining the Role of Non-Canonical NOD-like Receptors and Inflammasomes in

Inflammation and Disease

Jaye Matthew Platnich

A THESIS

SUBMITTED TO THE FACULTY OF GRADUATE STUDIES

IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE

DEGREE OF DOCTOR OF PHILOSOPHY

GRADUATE PROGRAM IN IMMUNOLOGY

CALGARY, ALBERTA

MARCH, 2018

The NOD-like Receptors (NLRs) are a family of pattern recognition receptors known to regulate a variety of immune signaling pathways. A substantial portion of NLR research focuses on the pyrin domain-containing NLRP subfamily. The canonical NLRPs are inflammasome-forming proteins responsible for the activation of caspase-1 and the maturation and secretion of the pro-inflammatory cytokines IL-1β and IL-18. In contrast, the non-canonical inflammasome-independent NLRPs regulate a variety of other pathways, including MAPK and NF-κB, through the formation of non-inflammasome complexes.

Interestingly, not all inflammasomes are nucleated by NLRPs. The recently characterized non-canonical caspase-4 (caspase-11 in mice) inflammasome is known to be a key driver of the innate immune response to intracellular pathogens (and the molecules associated with them), by triggering both inflammatory cell death and the activation of canonical inflammasomes.

At the outset of this PhD work, the understanding of both non-inflammasome- forming NLRPs and the non-canonical caspase-4 inflammasome was poor and the studies were sparse. It was the goal of this thesis to characterize the expression, gene regulation, and function of the non-inflammasome-forming NLR protein NLRP6, both at the cellular and biochemical level. Furthermore, using a pathogen-associated molecular pattern

(PAMP)-driven model of inflammation, we sought to elucidate the function of the non- canonical caspase-4 inflammasome, particularly as it pertains to the regulation of the canonical inflammasome and cell death.

ii By studying the fundamental biology underlying these lesser-known mediators of the innate immune system, we hoped to better understand their contribution to the early immune response and their role in driving inflammatory disease with a view to, one day, ameliorating the condition of patients suffering from these afflictions through the development of targeted therapeutics.

iii Acknowledgements

They say it takes a village to raise a child, and a PhD project is akin to the most squalling and unruly child imaginable. I could not have completed this work without the intellectual, technical, and emotional support of many individuals.

First, I’d like to sincerely thank my supervisor, Dr. Dan Muruve. You are among the cleverest people I’ve had the pleasure to meet and you have been a constant source of inspiration to me. Your steadfast commitment to my development as a scientist has been invaluable in terms of time, effort, and financial support.

I would also like to thank the other professors that served as mentors to me, namely Drs. May Ho, Justin MacDonald, and Hank Duff.

Second, I’d like to extend my thanks to my current and former labmates Arthur

Lau, Adom Bondzi-Simpson, Christie Sandall, Nathan Bracey, Sharon Clark, Hyunjae

“JC” Chung, Kosh Vilaysane, Justin Chun, and Takanori “Tak” Komada. I’d name you all as friends, but over the past four years we truly have become family. I can’t imagine a better group to have spent my time with and I wish you all the best in your future endeavors.

Third, I want to express my deepest gratitude to my family. My parents, Tim and

Tracy, and my sister Casey have been my bedrock of support throughout this project and

I would not have been able to complete it without you. I love you all beyond my ability to convey with words.

Last, but certainly not least, I want to offer my heartfelt thanks to my friends.

2017-2018 was a very challenging year for me, both academically and personally. Many

iv of you saw me at my emotional nadir and rose to the occasion magnificently. I cannot even begin to repay the kindness and patience you showed me. In particular, I would like to express my profound gratitude to Matt and Maryna Szojka, Alec Campbell, Alex Pynn,

David Guzzardi, Arthur Lau, Christie Sandall, Adom Bondzi-Simpson, and Richard

Magbojos for the special efforts you all made on my behalf.

v Table of Contents

Abstract ...... ii

Acknowledgements ...... iv

Table of Contents ...... vi

List of Tables ...... x

List of Illustrations and Figures ...... xi

List of Abbreviations ...... xiv

Chapter One: Introduction ...... 1

1.1 Introduction ...... 2 1.2 Pattern Recognition Receptors (PRRs) and the Innate Immune System ...... 3 1.2.1 Innate Immunity and Pattern Recognition Receptors (PRRs) ...... 3 1.2.2 NOD-like Receptors (NLRs) ...... 4 1.3 Canonical NLRs and Inflammasomes ...... 9 1.3.1 Canonical Inflammasomes ...... 9 1.3.2 NLRP1 Inflammasome ...... 10 1.3.3 NLRC4 Inflammasome ...... 11 1.3.4 AIM2 Inflammasome ...... 12 1.3.5 Pyrin Inflammasome ...... 13 1.3.6 NLRP3 Inflammasome ...... 13 1.4 Non-Inflammasome-Forming NLRs (Including Non-Canonical NLRP Proteins) and Their Functions ...... 16 1.4.1 NOD1/2 ...... 17 1.4.2 NLRP3 ...... 17 1.4.3 NLRP6 and NLRP12 ...... 18 1.5 NLRP6 ...... 19 1.5.1 Structure ...... 19 1.5.2 Expression ...... 24 1.5.3 Function ...... 25 1.6 Non-Canonical Inflammasome and Pyroptotic Cell Death ...... 28 1.6.1 Non-Canonical Inflammasome ...... 28 1.6.2 Pyroptosis ...... 29 1.6.3 NLRP3 Activation by the Non-Canonical Caspase-4 Inflammasome ...... 35 1.7 Normal Kidney Physiology and Disease States ...... 37 1.7.1 Kidney Physiology ...... 37 1.7.2 Pathogenesis of Kidney Disease ...... 37 1.8 Summary, Rationale, and Hypotheses ...... 39 1.8.1 NLRP6 Project ...... 40 1.8.2 Non-Canonical Caspase-4 Inflammasome Project ...... 41

vi 1.8.3 Contextual Importance of this Work ...... 42

Chapter Two: Materials and Methods ...... 43

2.1 Materials ...... 44 2.1.1 Reagents, Kits, and Buffers ...... 44 2.1.2 Antibodies ...... 46 2.1.3 Oligonucleotides ...... 48 2.2 Methods ...... 50 2.2.1 Animal Studies ...... 50 2.2.1.1 Anti-Glomerular Basement Membrane Nephrotoxic Serum Nephritis (NTS) ...... 50 2.2.1.2 Unilateral Ureteral Obstruction (UUO) ...... 50 2.2.1.3 Tissue Preparation ...... 51 2.2.1.4 Mechanical Glomerular Isolation ...... 51 2.2.1.5 Laser-Capture Microdissection ...... 52 2.2.2 Assays ...... 52 2.2.2.1 Bradford Assay ...... 52 2.2.2.2 Enzyme-Linked Immunosorbent Assay (ELISA) ...... 53 2.2.2.3 Immunoblotting ...... 54 2.2.2.4 Lactate Dehydrogenase Assay ...... 55 2.2.2.5 Reconstituted Inflammasome Expression ...... 55 2.2.2.6 NLRP Oligomerization Assay ...... 56 2.2.3 Cell Culture ...... 57 2.2.3.1 Cell Line Culture ...... 57 2.2.3.2 Primary Cell Culture ...... 58 2.2.3.2 Generation of CRISPR THP-1 Cell Lines ...... 60 2.2.4 Cloning and Polymerase Chain Reaction (PCR) ...... 62 2.2.4.1 Cloning ...... 62 2.2.4.2 Quantitative Real Time PCR (qRT-PCR) ...... 62 2.2.4.3 Site-Directed Mutagenesis ...... 63 2.2.4.4 XBP-1 Splicing Assay ...... 64 2.2.5 Flow Cytometry ...... 64 2.2.5.1 Flow Cytometric Analysis of CD77 Expression ...... 64 2.2.6 Imaging of Cells and Tissues ...... 65 2.2.6.1 Histology ...... 65 2.2.6.2 Immunocytochemistry ...... 65 2.2.6.3 Immunohistochemistry ...... 66 2.2.6.4 Live Cell Imaging with MitoSOX ...... 67 2.2.7 RNASeq ...... 67 2.2.7.1 RNASeq Analysis ...... 67 2.2.8 Shiga Toxin Preparation and Labeling ...... 68 2.2.8.1 Shiga Toxin Preparation ...... 68 2.2.7.2 Shiga Toxin Labeling with AlexaFluor 594 ...... 69 2.2.9 Subcellular Fractionation ...... 70 2.2.9.1 Subcellular Fractionation of Crude Membranes ...... 70 2.2.9.2 Subcellular Fractionation of Mitochondria ...... 70

vii 2.2.10 Statistical Analysis ...... 71

Chapter Three: Characterization of the Expression and Role of NLRP6 in the Kidney Under Both Normal And Disease States ...... 72

3.1 Rationale ...... 73 3.2 Results ...... 75 3.2.1 Nlrp6 mRNA, But Not Protein, is Expressed in Murine Glomerular Tissue 75 3.2.2 NLRP6 mRNA, But Not Protein, is Expressed in Human Glomerular Tissue ...... 79 3.2.3 NLRP6 Protein Remains Undetectable Under Various Protein Extraction Conditions ...... 81 3.2.4 The Commercially Available Antibodies Against Human NLRP6 Bind Different Domains of NLRP6 ...... 83 3.2.5 Nlrp6 +/+ and Nlrp6 -/- Mice Have Indistinguishable Phenotypes in Murine Models of Kidney Disease ...... 86 3.2.6 RNASeq Identifies a Novel Fusion Transcript and Reveals Select Differences in Gene Expression Between Nlrp6 -/- and Wild Type Mice at Baseline ...... 98 3.3 Summary ...... 107

Chapter 4: Characterization of the Fundamental Molecular Biology Underlying NOD-like Receptor Activation ...... 109

4.1 Rationale ...... 110 4.2 Results ...... 114 4.2.1 ATP-Binding in NLRs is Mediated by Residues in the Walker A, Walker B, and Lid Regions of the ATP-Binding Pocket...... 114 4.2.2 Mutation of Either the Walker A Motif or GxP Motif-Containing Lid Region Attenuates NLR Oligomerization...... 121 4.2.3 Mutation of Either the Walker A Motif or GxP Motif-Containing Lid Region Attenuates NLRP3 Inflammasome Activation...... 125 4.2.4 Abrogation of ATPase Activity in NLRs Results in Hyperactivity. 128 4.3 Summary ...... 134

Chapter 5: Caspase-4 and Gasdermin D Trigger the NLRP3 Inflammasome Through the Regulation of Mitochondrial Reactive Oxygen Species ...... 136

5.1 Rationale ...... 137 5.2 Results ...... 140 5.2.1 Shiga Toxins Activate the Canonical and Non-Canonical Caspase- 4 Inflammasome in Human Macrophages...... 140 5.2.2 Canonical and Non-Canonical Inflammasome Activation by Stx Requires NLRP3...... 147

viii 5.2.3 Shiga Toxin-Mediated Inflammasome Activation is Dependent on Co-Transported LPS...... 155 5.2.4 Shiga Toxin-LPS-Mediated Non-Canonical Inflammasome Activation Induces Mitochondrial ROS Upstream of NLRP3...... 159 5.3 Summary ...... 171

Chapter 6: Discussion and Future Directions ...... 176

6.1 Discussion and Future Directions ...... 177 6.1.1 Overview ...... 177 6.1.2 NLRP6 Project: Expression and Gene Regulation ...... 178 6.1.3 NLRP6 Project: Biochemistry ...... 183 6.1.4 Non-Canonical Caspase-4 Inflammasome Project ...... 187 6.1.5 Summary ...... 191

References ...... 192

Appendix A: List of Publications ...... 209

ix List of Tables

Table 2.1 Reagents ...... 44

Table 2.2 Commercial Kits ...... 44

Table 2.3 Cell Culture Media/Buffer ...... 45

Table 2.4 Buffers, Solutions, and Supplemented Media ...... 45

Table 2.5 Primary Antibodies for Immunoblotting and Flow Cytometry ...... 46

Table 2.6 Primary Antibodies for Immunohistochemistry/Immunocytochemistry ...... 47

Table 2.7 Secondary Antibodies ...... 47

Table 2.8 Oligonucleotides used for Cloning and Site-Directed Mutagenesis ...... 48

Table 2.9 Oligonucleotides used for PCR, RT-PCR, and qRT-PCR ...... 49

x List of Illustrations and Figures

Figure 1.1 Domain Architecture of NOD-like Receptors, AIM2, and the Adaptor Protein ASC...... 7

Figure 1.2 Proposed Mechanism of AAA+ (or STAND) ATPase Activity...... 21

Figure 1.3 Sequence Alignments of the Conserved Motifs of 17 Members of the NLR Family of Proteins...... 23

Figure 1.4 Domain Architecture of the Caspase Family of Proteases...... 32

Figure 1.5 Activation of the Canonical and Non-Canonical Inflammasome Mediates Both Cytokine Maturation and Pyroptotic Cell Death...... 33

Figure 1.6 Potential Signaling Cascades that Activate the Canonical NLRP3 Inflammasome Downstream of the Non-Canonical Inflammasome...... 36

Figure 3.1 Nlrp6 mRNA, But Not Protein, is Expressed in Murine Glomerular Tissue. . 77

Figure 3.2 NLRP6 mRNA, But Not Protein, is Expressed in Human Glomerular Tissue.80

Figure 3.3 NLRP6 Protein Remains Undetectable Under Various Protein Extraction Conditions...... 82

Figure 3.4 The Commercially Available Antibodies Against Human NLRP6 Bind Different Domains of NLRP6...... 84

Figure 3.5 NLRP6 Expression in the Unilateral Ureteral Obstruction (UUO) Model of Murine Kidney Disease...... 88

Figure 3.6 Nlrp6 +/+ and Nlrp6 -/- Mice Have Undistinguishable Histological and Biochemical Phenotypes in the Unilateral Ureteral Obstruction (UUO) Model of Murine Kidney Disease...... 90

Figure 3.7 NLRP6 Expression in the Nephrotoxic Serum Nephritis (NTS) Model of Murine Kidney Disease...... 94

Figure 3.8 Nlrp6 +/+ and Nlrp6 -/- Mice Have Undistinguishable Histological and Biochemical Phenotypes in the Nephrotoxic Serum Nephritis (NTS) Model of Murine Kidney Disease...... 96

Figure 3.9 Unique Fusion Splicing of BC024386 to Nlrp6 As Determined by RNAseq Analysis...... 101

Figure 3.10 Differential Expression of Genes Between Nlrp6 +/+ and Nlrp6 -/- Mice in Colon and Kidney Tissue as Determined by RNASeq...... 105

xi Figure 4.1 Structural Similarity and Phyre2 Modeling of NLRP3 and NLRP6...... 118

Figure 4.2 Model of the ATP-Binding Pocket of NLRP3...... 120

Figure 4.3 Mutation of the Putative Lid Region Impairs NLR Protein Oligomerization...... 123

Figure 4.4 NLRP3 is Functionally Impaired Following Mutation of the Putative Lid Region...... 127

Figure 4.5 Proposed Mechanism of NLRP3 ATPase Activity...... 130

Figure 4.6 NLRP3 is Functionally Hyperactive Following Mutation of Amino Acids Responsible for ATP Hydrolysis...... 131

Figure 4.7 Model for the Activation of NLR Proteins...... 133

Figure 5.1 Stx2 Activates the Canonical Inflammasome in Human Macrophages...... 142

Figure 5.2 Shiga Toxins Induce NLRP3, ASC, and Caspase-1 Containing Inflammasomes...... 144

Figure 5.3 Stx2 Triggers Non-Canonical Caspase-4 Inflammasome Activation and Pyroptosis in Human Macrophages...... 145

Figure 5.4 Murine Macrophages are Resistant to Shiga Toxins...... 150

Figure 5.5 Canonical and Non-Canonical Inflammasome Activation by Stx Requires NLRP3...... 152

Figure 5.6 Validation of the NLRP3 -/- (CRISPR) THP-1 Cell Line...... 154

Figure 5.7 Stx-Mediated Inflammasome Activation is Dependent on Co-Transported LPS...... 157

Figure 5.8 Shiga Toxin/LPS-Induced Inflammasome Activation is Independent of the ER Stress Response...... 160

Figure 5.9 GSDMD p30 Localizes to Membrane and Mitochondrial Fractions of Human Macrophages Following Stx/LPS Treatment...... 164

Figure 5.10 Stx/LPS-Mediated Non-Canonical Inflammasome Activation Induces Mitochondrial ROS Production To Drive Activation of the Canonical Inflammasome Upstream of NLRP3...... 165

Figure 5.11 Inhibitor Effects on Stx2/LPS Mediated ROS Production...... 167

Figure 5.12 Gene Knockout Effects on Stx2/LPS Mediated ROS Production...... 168

xii Figure 5.13 NLRP3 and GSDMD Are Required For Shiga Toxin/LPS-Induced ROS Generation...... 169

Figure 5.14 Potassium Efflux is Required for Stx-Mediated NLRP3 Inflammasome Activation...... 174

xiii List of Abbreviations

AAA+ ATPases ATPases-associated with diverse cellular activities

ADP adenosine diphosphate

AF-567 AlexaFluor 567

AIM2 absent in melanoma 2

AKI acute kidney injury

Anti-GBM anti-glomerular basement membrane

ASC apoptosis-associated speck-like protein containing a CARD

domain

ATF6 activating transcription factor 6

ATP adenosine triphosphate

A.U. arbitrary units

BMDM bone marrow-derived macrophage

BRCC3 breast cancer 1 (BRCA1)-breast cancer 2 (BRCA2)-containing

complex 3

CAPS cryopyrin-associated periodic syndromes

CARD caspase recruitment domain

Cas9 CRISPR (clustered regularly interspaced short palindromic

repeats)-associated protein 9

CD11b cluster of designation 11b, also known as ITGAM (integrin alpha

CD77 cluster of designation 77

xiv CKD chronic kidney disease

COL1A1 collagen, type 1, alpha 1 chain

CRISPR clustered regularly interspaced short palindromic repeats

CTB Cholera toxin subunit B

CTGF connective tissue growth factor

CTLR C-type lectin receptor

DAMP danger/damage-associated molecular pattern

DAPI 4',6-diamidino-2-phenylindole

DMEC dermal microvascular endothelial cell

DMEM Dulbecco’s modified eagle medium dsDNA double-stranded deoxyribonucleic acid

DSS dextran sulfate sodium

EHEC enterohemorrhagic Escherichia coli

ELISA enzyme-linked immunosorbent assay

ER endoplasmic reticulum

FBS fetal bovine serum

FCAS familiar cold autoinflammatory syndrome

FITC fluorescein isothiocyanate

GAPDH glyceraldehyde-3-phosphate dehydrogenase

GEC glomerular endothelial cell

GFP green fluorescent protein

GFR glomerular filtration rate

Gliben. glibenclamide, also known as glyburide

xv GM130 Golgi matrix protein 130 kiloDaltons

GSDMD gasdermin D

H&E hematoxylin and eosin stain

HRP horseradish peroxidase

HUS hemolytic uremic syndrome

IB immunoblot, also known as a western blot iE-DAP glutamyl-meso-diaminopimelic acid

IFNγ interferon gamma

IL-1α interleukin 1 alpha

IL-1β interleukin 1 beta

IL-18 interleukin 18

Inflammasome the inflammasome is defined in this thesis as any complex of

proteins that acts as a caspase-1 activating platform.

Inflammasomes generally consist of an NLRP protein (or AIM2),

the adaptor protein ASC, and the effector molecule caspase-1.

IP immunoprecipitation

IRE1α inositol-requiring enzyme 1 alpha kDa kiloDalton

KIM-1 kidney injury molecule 1

KO knock-out

LDH lactate dehydrogenase

LPS lipopolysaccharide

LRR leucine-rich repeat

xvi LYS cell lysate

MAPK mitogen-activated protein kinase

MDP muramyl dipeptide

MitoROS mitochondrial reactive oxygen species

MLKL mixed lineage kinase domain-like protein

MWS Muckle-Wells syndrome

NAC N-acetyl-L-cysteine

NACHT domain present in NAIP, CIITA, HET-E and TP1

NAIP NOD-like receptor family apoptosis inhibitory protein

NBD nucleotide-binding domain

NEK7 NIMA (never in mitosis gene A)-related kinase 7

NETs neutrophil extracellular traps

NF-κB nuclear factor kappa-light-chain-enhancer of activated B cells

NGC nigericin

NK cells natural killer cells

NLR NOD-like receptor or nucleotide-binding domain leucine-rich

repeat containing genes

NLRC4 NLR family card domain-containing 4

NLRP NLR family pyrin domain-containing (subgroup of the NLRs)

NLRP1 NLR family pyrin domain-containing 1, also known as NACHT,

LRR and PYD domains-containing protein-1

NLRP3 NLR family pyrin domain-containing 3, also known as NACHT,

LRR and PYD domains-containing protein-3

xvii NLRP6 NLR family pyrin domain-containing 6, also known as NACHT,

LRR and PYD domains-containing protein-6

NOD1 nucleotide-binding oligomerization domain-containing protein 1

NOD2 nucleotide-binding oligomerization domain-containing protein 2

NOMID neonatal-onset multisystem inflammatory disorder

NPHS1 nephrotic syndrome type 1, commonly known as nephrin

NPHS2 nephrotic syndrome type 2, commonly known as podocin

N.S. not significant

NT no treatment

NTS nephrotoxic serum

PAMP pathogen-associated molecular pattern

PBMC peripheral blood mononuclear cell

PBS phosphate-buffered saline qRT-PCR quantitative real time polymerase chain reaction

PERK PKR (protein kinase R)-like endoplasmic reticulum kinase

PMA phorbol 12-myristate 13-acetate

PMN polymorphonuclear cell

PolyB polymyxin b

PRR pattern recognition receptor

PYD pyrin domain

RIG-I retinoic acid-inducible gene I

RIPA buffer radioimmunoprecipitation assay buffer

RLR RIG (retinoic acid-inducible gene)-like receptor

xviii mRNA messenger ribonucleic acid

RNS reactive nitrogen species

ROS reactive oxygen species

RPMI 1640 Roswell Park Memorial Institution medium 1640

SD standard deviation

SDS sodium dodecyl sulfate

SEM standard error of the mean

αSMA alpha smooth muscle actin

STAND ATPases signal transduction ATPases with numerous domains

Stx1 shiga toxin 1

Stx2 shiga toxin 2

Stx2B shiga toxin 2, subunit b rStx2B recombinant shiga toxin 2, subunit b

SUP supernatant

T3SS type III secretion system

T4SS type IV secretion system

TBS TRIS-buffered saline

TC tunicamycin mTEC mouse renal tubular epithelial cell

TGFβ transforming growth factor beta

TLR toll-like receptor

TLR4 toll-like receptor 4

UUO unilateral ureteral obstruction

xix VDAC voltage-dependent anion channel

WA Walker A motif

WB Walker B motif

WT wild type

XBP-1 X-box-binding protein

xx Chapter One: Introduction

1 1.1 Introduction

Broadly speaking, this work focuses on the response of the innate immune system to inflammatory stimuli and pathology. Specifically, it investigates the role of select pattern recognition receptor pathways that do not conform to the conventional or canonical rules of their ascribed families. These receptors and pathways will be termed

“non-canonical” for the duration of this thesis.

Two distinct projects were completed over the course of this PhD work, related to inflammasome-independent pattern recognition receptor signaling and non-canonical inflammasome complexes.

The first project examined the biology of NLRP6, a poorly characterized NOD- like receptor (NLR), which functions quite differently from related members of the NLRP family of proteins. Chapters 3 and 4 will discuss the structure, expression, and function of

NLRP6 in detail, with a focus on its role in the kidney. It is worth noting that a basic assumption underlying the entire NLR field is that all members of the NLRP family can form caspase-1-activating inflammasomes and that this is their “canonical function’. No such function has been demonstrated for select family members, including NLRP6. These shall be referred to as “non-canonical NLRPs” or “non-inflammasome-forming NLRs” for the purposes of this thesis. This concept will be discussed at length later in this introduction.

The second project examined the activation of the caspase-4 inflammasome: an

NLR-independent inflammasome complex, commonly referred to as the “non-canonical inflammasome” because it does not directly activate caspase-1. Chapter 5 will discuss the

2 activation of the non-canonical inflammasome in a Shiga toxin model of inflammation in macrophages, focusing on the key proteins that regulate it and the downstream activation of the canonical NLRP3 inflammasome.

This introduction (Chapter 1) will provide a thorough review of the relevant literature, including a detailed overview of pattern recognition receptors, NLR biology, and inflammasomes, both canonical and non-canonical.

1.2 Pattern Recognition Receptors (PRRs) and the Innate Immune System

1.2.1 Innate Immunity and Pattern Recognition Receptors (PRRs)

The immune system consists of two main branches: the innate and adaptive immune systems. In contrast to the adaptive immune system, which mounts clonal, epitope-specific responses to encountered threats through the generation of B and T cells, the role of the innate immune system is to immediately respond to all threats in a specific, but non-clonal fashion. To accomplish this, the innate immune system makes use of a restricted set of germ-line-encoded receptors known as pattern recognition receptors

(PRRs) (1). PRRs are known to respond to a variety of pathogen- and danger-associated molecular patterns (PAMPs and DAMPs respectively), which arise during microbial infections and sterile injury respectively. Ligands known to stimulate PRRs include microbial molecules such as LPS and flagellin, as well as endogenous danger signals such as uric acid, ATP, and HMGB1 (1).

There are several families of PRRs. These can be categorized based on their structure, ligands, and the cellular compartments that they survey. The Toll-like receptors

3 (TLRs) and C-type lectin receptors (CLRs) are membrane-bound receptors that bind a variety of microbial ligands (e.g. lipopolysaccharide, peptidoglycan, and flagellin in bacteria; β-glucans in yeast, etc.) in both the extracellular and endosomal compartments

(1). Retinoic acid-inducible gene I (RIG-I) and absent in melanoma 2 (AIM2) are cytosolic PRRs that detect exogenous nucleic acids, predominantly in the context of viral infection. Finally the NOD-like receptors (NLRs), a fourth type of PRR, detect a wide range of both pathogen and danger-associated molecular patterns in the cytoplasm (1).

1.2.2 NOD-like Receptors (NLRs)

The NOD-like Receptor (NLR) family of proteins is a group of pattern recognition receptors (PRRs) known to mediate the initial innate immune response to cellular injury and stress (2-4). Members of the NLR family of proteins share many common structural features. They all possess C-terminal leucine-rich repeat (LRR) domains and central nucleotide binding (NBD) domains. The NLR family is divided into a variety of subfamilies, which differ significantly at their N-termini (3). The two most prominent of these subfamilies are the NLRC and NLRP groups of proteins. The NLRC proteins have one or more caspase recruitment (CARD) domains at their N-termini, whereas the NLRP proteins have pyrin domains (PYD) instead. The latter group, which consists of 14 distinct family members, is by far the best-characterized subfamily of

NLRs.

The NBD domain of all NLR proteins contains an ATP-binding pocket. This pocket has both Walker A and Walker B motifs, which mediate ATP-binding and hydrolysis respectively (5-7). ATP-binding is known to be essential for the

4 oligomerization and downstream functions of several NLRs (e.g. NOD2, NLRP3,

NLRP7, NLRP12) (6-9). Several of these family members are also known to have intrinsic ATPase activity (5-7). The specific roles of ATP-binding and hydrolysis in NLR activation have not been parsed. A detailed analysis of the ATP-binding and hydrolysis activities of these proteins will be presented in the specific context of NLRP6 and NLRP3 in Chapter 4.

Certain NLRP proteins are functionally related in their ability to form inflammasomes. Inflammasomes are multi-protein complexes consisting of an NLR, the adaptor protein apoptosis-associated speck-like protein containing a CARD domain

(ASC), and the effector protein pro-caspase-1 (3, 4). ASC has two domains: a pyrin domain and a CARD domain. This enables it to bridge interactions between NLRPs and pro-caspase-1. When activated, the NLRP recruits ASC via homotypic interactions between their pyrin domains, catalyzing a prion-like polymerization of the ASC molecules to enhance downstream signaling (10-12). Pro-caspase-1, a member of the cysteine-aspartic acid protease family, is subsequently recruited to the complex via

CARD-CARD interactions with the oligomerized ASC. The close association of many pro-caspase-1 molecules results in autocatalytic cleavage and the generation of p10 and p20 subunits. These assemble into the two heterodimers that constitute the activated form of caspase-1 (3, 4). Once activated, caspase-1 cleaves both pro-IL-1β and pro-IL-18 into their mature and secreted forms. IL-1β and IL-18 are both potent pro-inflammatory cytokines, with a wide variety of effects on the innate and adaptive immune systems.

Caspase-1 activation in leukocytes (in conjunction with caspase-4/5 in human and caspase-11 in mice) also results in a pro-inflammatory and non-apoptotic form of

5 programmed necrosis known as pyroptosis (4). This form of cell death will be discussed at length at a later point in this introduction.

Figure 1.1 Domain Architecture of NOD-like Receptors, AIM2, and the Adaptor

Protein ASC.

Figure legend on next page.

7 A colour-coded schematic diagram detailing the domain architecture of the 14 NLRP family members, NLRC4, NOD1 and 2, AIM2, and the inflammasome adaptor protein

ASC. Domain abbreviations are as follows: pyrin domain (PYD); domain present in

NAIP, CIITA, HET-E, and TP-1 (NACHT); leucine-rich repeat domain (LRR); caspase- recruitment domain (CARD); function to find domain (FIIND); B-box-type zinc finger domain (BB); coiled coil domain (CC); exon B30-2 domain (B30.2); and hematopoietic expression, interferon-inducible nature, and nuclear localization domain containing a 200 amino acid repeat (HIN-200).

8 1.3 Canonical NLRs and Inflammasomes

In the early days of the NLR field, it was assumed that all NLRP proteins formed caspase-1-activating inflammasomes. This became known as their “canonical function”.

Despite this assumption, certain NLRPs have not been biochemically demonstrated to form inflammasomes and activate caspase-1 (e.g. NLRP6, NLRP12). As such, NLRPs can be broadly subdivided into two categories: 1) the canonical NLRPs, which signal through the formation of caspase-1-activating inflammasomes and 2) the non-canonical

NLRPs, which function through inflammasome-independent mechanisms. Some NLRPs, such as NLRP3, have both canonical and non-canonical functions depending on the stimulus and the tissue-type in question. Furthermore, other NLRs and non-NLR proteins are also capable of assembling canonical (caspase-1) inflammasomes. This section will discuss the biology of canonical, caspase-1-activating inflammasomes. The non- inflammasome-forming NLRs (including both NOD1/2 and the non-canonical NLRPs) and the functions ascribed to them will be discussed in the subsequent section.

1.3.1 Canonical Inflammasomes

The term “inflammasome” was first coined in 2002 by Martinon et al. to describe an NLRP1-containing caspase-1/caspase-5-activating complex present in leukocytes treated with inflammatory stimuli (13). Since then, six widely accepted inflammasome complexes have been identified. These are the NLRP1 inflammasome, the NLRC4 inflammasome, the AIM2 inflammasome, the Pyrin inflammasome, the NLRP3

9 inflammasome, and the non-canonical caspase-4 (caspase-11 in mice) inflammasome (4).

The non-canonical inflammasome will be discussed at length later in this introduction.

1.3.2 NLRP1 Inflammasome

The NLRP1 protein differs from its relatives in the NLRP family of proteins in that it possesses a C-terminal CARD domain as well as an N-terminal pyrin domain

(Figure 1.1) (14). This enables it to directly recruit caspase-1 in the absence of the ASC adaptor protein. Notwithstanding this fact, ASC is thought to stabilize the CARD-CARD interactions between NLRP1 and caspase-1 in order to drive the immune response (15,

16).

The mouse orthologs of NLRP1 have been reasonably well characterized. Mouse

NLRP1b is important for coordinating the host immune response to the lethal toxin of

Bacillus anthracis – mice deficient in NLRP1b are impaired in clearing B. anthracis infections (17-19). Specifically, lethal toxin is responsible for inducing pyroptosis in mouse macrophages via NLRP1 activation (15). Interestingly, lethal toxin is not an agonist of human NLRP1, whose only known ligand is muramyl dipeptide – a microbial cell wall product (20).

In terms of human disease, the NLRP1 inflammasome is genetically associated with vitiligo, Addison’s disease, autoimmune thyroid disease, type I diabetes, and a number of other autoimmune diseases (21, 22). The mechanisms underlying NLRP1 involvement in these diseases remain unresolved and are in need of further investigation.

10 1.3.3 NLRC4 Inflammasome

As previously mentioned, members of the NLRC family of proteins have an N- terminal CARD domain through which they can directly interact with RIP2 or pro- caspase-1 (Figure 1.1) (23). One family member, NLRC4, interacts directly with caspase-

1 to form an inflammasome. Consequently, pyroptosis downstream of NLRC4 activation should theoretically be independent of ASC involvement. Notwithstanding this fact, ASC is necessary for both caspase-1 autocatalytic cleavage and the maturation of both IL-1β and IL-18 downstream of NLRC4 activation (24, 25). This partial requirement for ASC- mediated interactions mirrors the findings noted with NLRP1 (16).

The NLRC4 inflammasome plays a significant role in the immune response to intracellular bacteria such as Salmonella typhimurium, Shigella flexneri, Pseudomonas aeruginosa, and Legionella pneumophila (2). Activation is mediated through a number of

PAMPs produced by these pathogens. These include flagellin and the components of both type III (T3SS) and type IV (T4SS) bacterial secretion systems (26-29). In mice, multiple

NOD-like receptor family apoptosis inhibitory protein (NAIP) proteins operate upstream of NLRC4 to mediate the activation of the inflammasome in response to these stimuli

(30, 31). Humans only possess a single NAIP protein, which has been implicated in binding proteins from the T3SS of various bacteria, including Chromobacterium violaceum (31). A recent study has identified a second, extended isoform of human NAIP that confers responsiveness to Salmonella flagellin (32).

These findings have led to a model of NLRC4 inflammasome activation wherein

NLRC4 serves as a caspase-1-recruiting adaptor protein downstream of the PAMP- sensing NAIP proteins.

11 1.3.4 AIM2 Inflammasome

The AIM2 protein is not a member of the NLR family of proteins - it falls into a separate category of PRRs. Structurally, it consists of two domains: a HIN200 domain and a pyrin domain (Figure 1.1). The HIN-200 domain allows for the detection of double- stranded DNA (dsDNA) in the cytoplasm, whereas the pyrin domain allows for the recruitment of ASC and the formation of an inflammasome complex (33-35). Indeed, stimulation of macrophages and dendritic cells with cytosolic dsDNA triggers a potent activation of caspase-1, IL-1β cleavage, and the initiation of pyroptotic cell death (33,

36).

The AIM2 inflammasome is involved in the response to a number of intracellular bacterial and viral infections. Specifically, caspase-1 cleavage and IL-1β secretion is potently induced in macrophages infected with Francisella tularensis, vaccinia virus, and mouse cytomegalovirus (mCMV) in an AIM2-dependent manner (36, 37). Additionally,

AIM2-deficient mice infected with mCMV have higher viral titers compared to wild-type mice as a result of impaired IFNγ production from NK cells. This impairment is linked to the absence of AIM2 inflammasome activity via the inability to generate mature IL-18 – a key cytokine in the induction of IFNγ secretion from NK cells (36).

Given its ability to sense dsDNA, it is no great surprise that AIM2 also has a role in the pathophysiology of systemic lupus erythematosus. Specifically, patients with the disease have been found to express higher levels of AIM2, contributing to enhanced IL-

1β-mediated inflammation (38, 39).

On a completely unrelated note, it is worth mentioning that the AIM2 protein also has important inflammasome-independent functions as a tumor suppressor protein – its

12 absence has been noted in a number of cancers, specifically melanoma and colorectal cancer (40, 41). These effects are mediated through the suppression of cellular proliferation and enhancement of apoptosis (42). Specifically, AIM2 limits DNA-PK- mediated phosphorylation of Akt: a key driver of cell growth and survival (41, 43).

1.3.5 Pyrin Inflammasome

The human pyrin protein consists of an N-terminal pyrin domain, two B-box domains, a coil-coiled domain, and a C-terminal B30.2 domain (the mouse protein lacks the B30.2 domain) (Figure 1.1) (4). Given the early state of the field, its function has only begun to be characterized. In the context of various bacterial toxins, including

Clostridium difficile toxin B and Clostridium botulinum C3 toxin, pyrin can detect, become activated by, and form an inflammasome in response to the specific inactivation of RhoA and aberrations in cytoskeleton dynamics (44, 45). The importance of this finding remains to be elucidated, as C. difficile toxins also activate the NLRP3 inflammasome (4, 46).

1.3.6 NLRP3 Inflammasome

The NLRP3 inflammasome is the best studied and most possesses the most complex signaling of all inflammasomes. Structurally, it is a prototypical model of the

NLRP subset of NLR proteins. It has a C-terminal LRR domain, a central NACHT domain, and an N-terminal pyrin domain (Figure 1.1). When activated, the pyrin domain allows for the recruitment of ASC via homotypic PYD-PYD interactions. This allows for the recruitment and activation of pro-caspase-1, and ultimately the maturation of IL-1β

13 and IL-18 (2). The most recent model for the assembly of these inflammasome proteins involves a prion-like polymerization of the pyrin domains of ASC, nucleated by the pyrin domain of activated NLRP3 (10, 11).

There are two key signals allowing for the activation of the NLRP3 inflammasome: a priming signal and an activation/assembly signal. The priming signal, a unique requirement among NLRPs, is triggered through the stimulation of PRRs via

PAMPs and DAMPs and subsequent signaling through NF-κB (47, 48). This increases the expression of both NLRP3 and pro-IL-1β, although it is unclear whether or not the induction of NLRP3 expression is necessary for subsequent inflammasome activation

(49, 50). Priming also has a second consequence: the deubiquitination of NLRP3 by

BRCC3, which is a prerequisite to NLRP3 inflammasome activation (51). The mechanism through which deubiquitination contributes to NLRP3 inflammasome activation is incompletely characterized.

The second signal in the activation process is less well understood. There are a myriad of PAMPs and DAMPs that activate the NLRP3 inflammasome including ATP, uric acid, nigericin, alum, asbestos, amyloid aggregates, as well as a range of microbial toxins including Clostridium difficile toxin and Shiga toxins (2, 46, 52). None of these stimuli directly bind and activate NLRP3. Instead, these stimuli engage a number of common pathways that ultimately culminate in NLRP3 inflammasome activation. These pathways include potassium efflux from the cell, lysosomal rupture, calcium mobilization, the production of reactive oxygen species (ROS), NLRP3 translocation to the mitochondria, and the release of mitochondrial DAMPs, such as DNA and cardiolipin

(53-60). The current thought is that potassium efflux is the convergence point for all of

14 these pathways, as blockade of potassium efflux inhibits inflammasome activation downstream of all of the aforementioned pathways and low potassium alone is sufficient to trigger NLRP3 inflammasome activation (56). In this model, NLRP3 serves as a general sensor for disturbances in cellular homeostasis.

Apart from BRCC3-mediated deubiquitination, there are a number of post- translational modifications and protein-protein interactions that mediate NLRP3 inflammasome assembly and activation (51). These include negative regulation by phosphorylation and the interaction with PYD-only and CARD-only proteins (POPs and

COPs respectively) (4, 61). Most recently, a key role has been found for NEK7 – a kinase known to be involved in mitosis. NEK7 binds to NLRP3 and is required for its oligomerization, in a manner independent of its kinase activity (62-64). Interestingly, given the role of NEK7 in mitosis, NLRP3 inflammasome assembly is far more efficient in interphase than in cells undergoing cell division.

To offer some clinical correlation to these findings, there are a few diseases arising directly from NLRP3 inflammasome dysregulation. A number of mutations

(which will be discussed at length in subsequent sections) have been described in the

NLRP3 gene that result in the constitutive activation of the NLRP3 inflammasome. These conditions are collectively referred to as cryopyrin-associated periodic syndromes

(CAPS). There are three distinct subtypes of CAPS: familial cold autoinflammatory syndrome (FCAS), Muckle-Wells syndrome (MWS), and neonatal-onset multisystem inflammatory disease (NOMID). These syndromes share a common pathophysiology: the dysregulated production of active IL-1β leading to pathologic inflammation in multiple organ systems including skin, mucous membranes, joints, ears, and nervous system,

15 depending on the subtype of the disease (65). Evidence for the direct involvement of IL-

1β in the pathology of these diseases is evident in the effectiveness of canukinumab, an

IL-1β neutralizing antibody, in the treatment of these disorders (66).

Apart from the CAPS syndromes, the NLRP3 inflammasome has been implicated in a large number of other human diseases including gout, inflammatory bowel disease, atherosclerosis, neurodegenerative disease, arthritis, diabetes, and cancer (67). The mechanism of NLRP3 inflammasome involvement in some of these diseases is very well characterized. In gout, uric acid crystals accumulate in the joints resulting in caspase-1 activation and IL-1β secretion in macrophages. The direct role for IL-1β in gout is clearly evidenced by the effectiveness of canakinumab treatment in clinical trials for gouty arthritis (68). The pathophysiologic mechanisms are less well defined in the other aforementioned diseases. For the neurodegenerative diseases, the current understanding suggests that aggregated or misfolded proteins (e.g. β-amyloid) serve as the activating signal for NLRP3 (69). In cancer the paradigm is that NLRP3 inflammasome dysregulation leads to chronic inflammation, cellular dysplasia, and the subsequent development of neoplasms (67).

1.4 Non-Inflammasome-Forming NLRs (Including Non-Canonical NLRP

Proteins) and Their Functions

As a family, NLRs are linked by their common protein domains/motifs and their need to oligomerize in order to function. Functionally, they are quite diverse. Select family members form caspase-1 activating inflammasomes in order to signal – this is the

16 assumed canonical function of NLRP family members in the literature. Others, such as

NOD1 and 2, operate entirely independent of inflammasomes. Whilst some of these inflammasome-independent signaling pathways are well understood, others (including the inflammasome-independent functions of select NLRP proteins) remain enigmatic.

1.4.1 NOD1/2

NOD1 and NOD2 are two well-studied NLRs that signal via NF-κB and MAPK following stimulation with bacterial peptidoglycan derivatives (i.e. iE-DAP and MDP respectively), in an inflammasome-independent manner (70). In the case of NOD2, MDP binding triggers oligomerization of NOD2, the recruitment of RIPK2 via CARD-CARD interactions, and downstream activation of NF-κB and MAPK via the phosphorylation of

TAK1 and MAPK8 respectively (70).

1.4.2 NLRP3

NLRP3, apart from its well-characterized ability to form an inflammasome in leukocytes (mostly macrophages and dendritic cells), regulates several other cellular processes. These pathways vary substantially depending on tissue/cell-type and the expression of inflammasome components.

In the context of epithelial cells, NLRP3 positively regulates TNFα and CD95- mediated apoptosis through the formation of a non-canonical ASC-containing, caspase-8- activating inflammasome complex (71).

17 It also positively regulates TGF-β-mediated fibrosis and EMT through the regulation of mitochondrial ROS generation in both epithelial cells and fibroblasts (72,

73).

In differentiated T cells, NLRP3 is localized to the nucleus where it acts as a transcription factor in concert with IRF4 to drive Th2-type polarization (74).

1.4.3 NLRP6 and NLRP12

Apart from NLRP1 and NLRP3, little is known about the other 12 members of the

NLRP family of proteins. Of these, the most studied are NLRP6 and NLRP12. Neither of these proteins has been biochemically demonstrated to form an inflammasome and both are reported to negatively regulate NF-κB and MAPK signaling (75-78). Both of these proteins are anti-inflammatory and anti-tumorigenic in the context of inflammatory bowel disease (77, 79). The mechanisms underlying these processes remain unknown.

Given the paucity of knowledge surrounding NLRP6, its known inflammasome- independent functions in terms of signaling in disease processes, and its high level of expression in our model organ (i.e. kidney), this thesis sought to explore the biology of

NLRP6 in the kidney, both under normal homeostatic and disease states. The following section will review the field of NLRP6 biology and will outline the rationale for its study.

18 1.5 NLRP6

1.5.1 Structure

NLRP6 is a prototypical member of the NLRP family of proteins containing an N- terminal pyrin domain, central NACHT domain (containing a nucleotide-binding domain), and five C-terminal leucine-rich repeats (Figure 1.1). It consists of 892 amino acids and has an approximate size of 99 kDa. It was first identified as an APAF-1-like protein, originally designated PYPAF5 (80).

NLRP6, and the NLR proteins as a group, are members of the AAA+ (ATPases- associated with diverse cellular activities) or STAND (signal transduction ATPases with numerous domains) family of ATPases (5). These ATPases, which are highly conserved among prokaryotic and eukaryotic species, possess a number of common structural motifs involved in both ATP-binding and hydrolysis (8). These include, but are not limited to, the Walker A motif, the Walker B motif, and an arginine finger motif (8).

Classically, AAA+ ATPases are thought to oligomerize to form hexamers, through which

ATPase activity and downstream function can be coordinated (8).

The Walker A motif, possessing the consensus sequence GXXXXGKS/T, coordinates ATP-binding via electrostatic interactions in the ATP-binding pocket (8).

This is in contrast to the Walker B motif, which contributes the catalytic residues necessary for ATP hydrolysis. The consensus sequence of the Walker B motif ends DE

(81, 82).

The proposed unified model for ATP hydrolysis by AAA+ ATPases is as follows.

Once ATP is bound in the ATP-binding pocket, its negatively charged phosphate

19 moieties are stabilized by both interactions with a magnesium ion (itself coordinated by the polar serine/threonine residue of the Walker A site and the aspartic acid residue of the

Walker B site), the positive charges present in the Walker A site and the positively charged arginine finger motif (Figure 1.2) (81, 82). In most AAA+ ATPases, this arginine residue is contributed in trans from a neighboring subunit in the hexameric form of the protein (81). This occurs in a manner analogous to that seen in small GTPase activation following GTPase-activating protein (GAP) binding (83). The net effect is to render the gamma phosphate a better electrophile by drawing away electron density from the central phosphorus atom. In concert, a water molecule is activated through interactions with the glutamic acid residue of the Walker B site. By enhancing the nucleophilic character of the water molecule’s oxygen atom, the glutamate residue serves to catalyze the nucleophilic attack of the gamma phosphate by the water molecule, resulting in the breakdown of ATP into ADP and a free phosphate molecule (Figure 1.2) (81). Defects in any of these key catalytic residues abrogate ATP hydrolysis.

Figure 1.2 Proposed Mechanism of AAA+ (or STAND) ATPase Activity.

A schematic diagram depicting the proposed mechanism of AAA+ ATPase activity in the literature. The arginine finger motif is highlighted in purple, the residues of the Walker B motif are highlighted in orange, and the relevant serine/threonine residue of the Walker A motif is highlighted in cyan. The curved arrows depict a nucleophilic attack on the γ- phosphate moiety of the ATP molecule, catalyzed by the aforementioned residues of the

Walker A, Walker B, and arginine finger motifs in concert with a magnesium ion (Mg2+).

21 NLRP6, and at least 16 other NLR family members, possess some or all of these conserved structural motifs (Figure 1.3). Of note, several hyperactive mutations of NLR family members occur at or near residues thought to be crucial for ATP hydrolysis (84).

For example, the R334W mutation in NOD2, which results in an auto-inflammatory disease known as Blau syndrome, is thought to abrogate the ATPase activity of the protein thereby locking NOD2 into an ATP-bound, active configuration that drives the disease (85). Interestingly, this arginine residue is conserved in the NLRP3 protein where the equivalent mutation (R262W) is responsible for a family of auto-inflammatory diseases known as cryopyrin-associated periodic syndromes or CAPS (5, 84). The precise mechanisms underlying the hyperactivity of NLRP3 in these conditions remain unknown.

Figure 1.3 Sequence Alignments of the Conserved Motifs of 17 Members of the NLR

Family of Proteins.

A sequence alignment of the Walker A, Walker B, and arginine finger motifs of 17 NLR family members. Bolded letters indicate a conserved residue consistent with the known consensus sequence of the motif. Lowercase letters in the consensus sequence indicate the most common amino acid residue present at that location.

23 1.5.2 Expression

The first expression studies conducted for the NLRP6 gene were performed in human leukocytes and revealed that human NLRP6 (hNLRP6) mRNA expression was restricted to granulocytes and T cells (80). Subsequent papers found murine Nlrp6

(mNLRP6) mRNA to be most highly expressed in the small intestine, colon, liver, and kidney under basal conditions (79, 86-88). Furthermore, murine Nlrp6 mRNA expression falls in the context of disease states and infection (87, 89, 90).

Until recently, little was known as to which signaling pathways regulate NLRP6 mRNA expression. The PPARγ agonist rosiglitazone modestly upregulates intestinal

NLRP6 expression in vitro and in vivo, but it remains to be seen whether this is a major regulator of NLRP6 expression (89, 91). One recent paper on the subject suggests that murine Nlrp6 may be an interferon-stimulated gene, upregulated in the context of infection with RNA viruses. Apart from active viral infection, murine Nlrp6 mRNA expression increases in epithelial cells treated with either type 1 or type 3 interferons or with the TLR3 agonist poly(I:C), in an NF-κB-independent manner (92).

NLRP6 protein has only been reliably detected in intestinal tissue (79, 87, 88).

The issue is not due to limitations of the available antibodies, as immunoblotting using a

FLAG antibody in tissues isolated from an NLRP6-FLAG knock-in transgenic mouse failed to detect NLRP6 in any other tissue type outside of the intestine (92).

24 1.5.3 Function

At the start of this thesis work, very little was known about the function of

NLRP6 and the findings to date remain highly controversial. Though possible under overexpression conditions in vitro, the ability of NLRP6 to form a functional inflammasome in vivo remains controversial and has not been demonstrated biochemically (80). As such, it can be designated as a “non-canonical” or non- inflammasome-forming NLRP.

Most studies of NLRP6 function have been conducted in intestinal tissue, where

NLRP6 has an anti-inflammatory and anti-mitogenic role in the context of DSS colitis models of inflammatory bowel disease (79, 87). Specifically, mice deficient in Nlrp6 experience a more severe disease phenotype following induction of colitis with either dextran sodium sulfate (DSS) or azoxymethane (AOM)/DSS in terms of shortened bowel length, weight loss, histological score, and tumor number/size. The latter finding is due to enhanced epithelial cell proliferation in Nlrp6 -/- mice (79, 87).

One of the proposed mechanisms underlying these observations is that deficiency in NLRP6 reduces the expression of IL-18 in the gut (both by the epithelial cells and recruited inflammatory monocytes), which leads to the dysregulation of the intestinal microbiota (in particular the expansion of the Bacteroidetes phyla of bacteria) and the dysregulation of the IL-22/IL-22BP axis: a key regulator of the wound healing and epithelial cell repair pathways of the gut (88, 93, 94). This perturbation of intestinal microbiota promotes increased CCL5 expression in the gut, resulting in enhanced leukocyte recruitment and inflammation in the DSS colitis model. This same group also demonstrated that this increased CCL5 expression drives tumorigenesis through the

25 regulation of the IL-6 signaling axis (88, 95). Additionally, there is interplay between microbial metabolites in the gut and the production of IL-18-regulated anti-microbial peptides downstream of NLRP6, explaining the transferability of the colitogenic phenotype from Nlrp6-deficient mice to wild type mice following fecal transfer (96).

These ascribed functions of NLRP6 remain hotly contested, with two recent studies concluding that NLRP6 and ASC play no role in the regulation of the commensal intestinal microbiota when the appropriate littermate and ex-germ-free control experiments are performed (97, 98).

A second proposed mechanism explaining the colitogenic effect of NLRP6 deficiency deals with mucus secretion (Muc2) by goblet cells. Nlrp6 -/- mice, through defects in autophagy and NLRP6 inflammasome activation, are unable to properly secrete mucous into the lumen of the gut (99, 100). This renders the mouse unable to effectively clear enteric pathogens, allowing for the development of persistent infections and an enhanced pro-inflammatory phenotype.

Despite suggesting the presence of an in vivo NLRP6 inflammasome in all of the aforementioned studies, no biochemical data (i.e. co-immunoprecipitation) has been presented to validate this supposition, apart from the original over-expression experiments performed in vitro (80).

Apart from the functions ascribed to an inflammasome-dependent mechanism, two inflammasome-independent functions for NLRP6 have been reported. First, consistent with the anti-inflammatory findings noted above, mice deficient in NLRP6 are more resistant to Salmonella and Listeria infection, where NLRP6 negatively regulates both NF-κB and ERK1/2-mediated MAPK signaling in leukocytes (76). Nlrp6 -/- mice

26 have increased circulating monocytes and neutrophils, as well as higher levels of NF-κB- dependent cytokines in the blood, explaining the resistance to the studied infections (76).

Second, it was demonstrated that NLRP6 is a viral RNA sensor (in association with the helicase Dhx15) that serves to upregulate interferon-stimulated genes, namely type 1 and type 3 interferons, and that NLRP6 is upregulated by these signaling molecules (92). NLRP6 deficiency was found to be harmful in the context of infection with enteric RNA viruses, where NLRP6-deficient mice were found to have impaired gut clearance of the ECMV and murine noroviruses when infected orally (92).

Neither of these inflammasome-independent functions has been consistently reproduced in the literature, so it remains to be seen whether these findings apply broadly beyond the limitations of the studied models. As a result, the significance of these findings remains dubious.

In summary, the function of NLRP6 remains poorly understood. Most of the studies to date have been restricted to the gut and demonstrate variable results depending on the model and controls employed.

27 1.6 Non-Canonical Inflammasome and Pyroptotic Cell Death

1.6.1 Non-Canonical Inflammasome

Kayagaki et al. first used the term “non-canonical inflammasome” in 2011 to describe findings that caspase-11 (the mouse ortholog of human caspases-4 and 5) was activated in a manner independent from the canonical NLR-ASC-caspase-1 inflammasome paradigm (101). This was the first study to demonstrate that certain stimuli, specifically exposure to Cholera toxin subunit B (CTB) and infection with various Gram-negative bacteria, triggered caspase-1-independent cell death and caspase-

1-dependent IL-1β and IL-18 secretion downstream of caspase-11 activation.

Furthermore, mice deficient in caspase-11 were protected following injection with a lethal dose of lipopolysaccharide (LPS) (101). This same group later discovered that LPS was the common molecule triggering non-canonical inflammasome activation downstream of each of the aforementioned stimuli, in a manner independent of TLR4

(102). Specifically, in terms of their toxin studies, LPS was bound to the B subunit of the

Cholera toxin and co-transported into the cell, in response to which the non-canonical inflammasome became activated. Our own work revealed that Escherichia coli Shiga toxin, a related AB5 bacterial toxin, acted similarly to drive non-canonical inflammasome activation through the co-transport of LPS.

At the time, it was thought that an unidentified LPS sensor existed upstream of caspase-11 activation in manner analogous to NLR-mediated caspase-1 activation. It was later found that caspases-4, 5, and 11 directly bound LPS, triggering both oligomerization

28 and activation of these caspases (103). In other words, they serve as both the PRR and effector molecule.

Non-canonical inflammasome activation has two downstream consequences: 1) the induction of pyroptosis, an inflammatory form of programmed cell death and 2) the secretion of the pro-inflammatory cytokines IL-1β and IL-18 via the activation of the canonical NLRP3 inflammasome (101, 102, 104). Gasdermin D (GSDMD), a 50 kDa protein containing a 30 kDa N-terminal domain and a 20 kDa C-terminal domain, is the effector molecule that links these two downstream processes (105, 106).

1.6.2 Pyroptosis

Pyroptosis is a form of programmed cell death. Programmed cell death can be subdivided into two broad categories: 1) apoptosis and 2) programmed necrosis.

Apoptosis is a non-inflammatory form of cell death that occurs as a result of exposure to various cell intrinsic and extrinsic, death-inducing stimuli. Cells undergoing apoptosis display a readily identifiable phenotype and morphology: the nucleus and cytoplasm condense, the cell undergoes blebbing, the DNA undergoes fragmentation, mRNA is degraded, and phosphatidylserine is expressed on the outer leaflet of the cell membrane (to facilitate recognition by scavenger receptors and clearance by macrophages) (107).

Apoptotic signaling occurs via the autocatalytic activation of the initiator caspases-8 and 9 (for the extrinsic and intrinsic pathways of apoptosis respectively); the proteolytic activation of the executioner caspases-3, 6 and 7; and the cleavage of a plethora of target substrates to induce cell death (Figure 1.4) (107). The resulting

29 apoptotic bodies can then be cleared by macrophages in a non-inflammatory manner.

Failure to efficiently clear these apoptotic bodies can result in their lysis, in a process known as secondary necrosis (107).

In contrast, programmed necrosis is a type of lytic, pro-inflammatory death characterized by cell swelling, loss of membrane integrity, and the release of cellular contents into the extracellular space (107, 108). There are a number of different programmed necrosis pathways including necroptosis, ferroptosis, NETosis, and, most relevant for the purposes of this thesis, pyroptosis (107). The term “pyroptosis” describes a process of lytic cell death downstream of inflammatory caspase activation (i.e. caspases-1, 4, 5, and 11), resulting from canonical or non-canonical inflammasome activation (Figures 1.4, 1.5). Gasdermin D (GSDMD), the molecular effector of pyroptosis, was only recently characterized.

Activation of any of the inflammatory caspases triggers the cleavage of gasdermin

D at Asp275 (Asp276 in mouse), allowing the pore-forming, 30 kDa N-terminal domain to disassociate from the auto-inhibitory C-terminal domain (106, 109, 110). The N- terminal fragment of GSDMD oligomerizes following cleavage, forms pores in the plasma membrane (with an inner diameter of approximately 13 nm), and triggers cell lysis (Figure 1.5) (110-114). Over-expression of the N-terminal fragment is sufficient to trigger death in HEK293T cells (105). Interestingly, cleaved GSDMD could only insert in membranes with select lipid compositions. Specifically, N-terminal GSDMD possessed affinity only for membranes rich in phosphatidic acid, phosphatidyl serine, mono- and bisphosphorylated phosphoinositols, and cardiolipin (111-113). This suggested that N- terminal GSDMD could bind to and induce permeabilization of the inner leaflet of the

30 plasma membrane, inner leaflet-derived membrane-bound organelles, and potentially both bacterial and mitochondrial membranes (111, 112). Notably, cleaved GSDMD had no affinity for the outer leaflet of the cell membrane, limiting damage to bystander cells following pyroptotic death (111). This pattern of lipid affinity is similar to MLKL, the pore-forming protein responsible for executing necroptotic cell death, suggesting a partially conserved mechanism across the known pathways of programmed necrosis

(115).

Figure 1.4 Domain Architecture of the Caspase Family of Proteases.

A colour-coded schematic diagram depicting the domain architecture of the inflammatory, initiator, and executioner caspases. Relevant domain abbreviations are as follows: caspase-recruitment domain (CARD); death-effector domain (DED); and the p20 and p10 catalytic subunits (p20/p10).

Figure 1.5 Activation of the Canonical and Non-Canonical Inflammasome Mediates

Both Cytokine Maturation and Pyroptotic Cell Death.

Figure legend on next page.

33 A schematic diagram depicting the activation of the canonical and non-canonical inflammasomes. Activation leads to the cleavage of caspases-1 and 4, which in turn triggers the cleavage of gasdermin D and the induction of pyroptotic cell death. Caspase-

1 also results in the maturation of the IL-1β and IL-18 into their active and secreted forms to drive inflammation.

34 1.6.3 NLRP3 Activation by the Non-Canonical Caspase-4 Inflammasome

The mechanisms underlying canonical NLRP3 inflammasome activation downstream of the non-canonical inflammasome are only beginning to be understood.

Potassium (K+) efflux following gasdermin D permeabilization of the plasma membrane is important in this process, consistent with its known role in NLRP3 activation (Figure

1.6) (116, 117). These findings are consistent with an analogous pathway described in the field of necroptosis, where MLKL-dependent potassium efflux activates the NLRP3 inflammasome to promote cytokine maturation (118, 119).

Knowing that the common pathways of NLRP3 activation include K+ efflux, mitochondrial ROS generation, lysosomal rupture, and calcium mobilization, it is a reasonable possibility that GSDMD-mediated permeabilization of a variety of membrane- bound organelles act collectively upstream of the NLRP3 inflammasome to induce both inflammation and cell death (Figure 1.6).

Figure 1.6 Potential Signaling Cascades that Activate the Canonical NLRP3

Inflammasome Downstream of the Non-Canonical Inflammasome.

A schematic diagram depicting the known and theorized pathways of NLRP3 inflammasome activation downstream of the non-canonical inflammasome. Hashed arrows and question marks denote speculative theories.

36 1.7 Normal Kidney Physiology and Disease States

1.7.1 Kidney Physiology

The kidney performs a variety of critical functions in normal animal physiology including electrolyte and acid-base homeostasis, the regulation of blood pressure, and the production of erythropoietin to induce erythrocyte production. By far the most important function of the kidney is to filter the blood, reabsorb important metabolites, and remove waste products (e.g. urea, ammonia, uric acid) and excrete these into the urine (120).

In clinical medicine, the effectiveness of the kidney in performing this function is estimated with a metric known as the glomerular filtration rate (GFR). In the event of kidney disease, this filtering and excretion process is impaired and these waste products are allowed to accumulate in the bloodstream. This is registered by a rapid decline in

GFR. These events can be defined as either acute or chronic, depending on the timeframe of the illness. Chronic kidney disease (CKD) arises when this reduction in GFR fails to rebound over the long term or progressively falls to the point of kidney failure (120).

1.7.2 Pathogenesis of Kidney Disease

The etiologies of kidney disease can be divided into three categories: pre-renal, renal, and post-renal (120). Pre-renal causes of kidney disease (e.g. congestive heart failure) significantly impair the perfusion of the kidney. Renal causes directly damage the intrinsic components of the kidney, namely the tubules, the glomeruli, the interstitium, and the vasculature (120). For the purposes of this thesis, a relevant example of renal acute kidney injury is hemolytic uremic syndrome (HUS): the systemic manifestation of

37 enterohemorrhagic E. coli infection. In this disease, Shiga toxin enters the systemic circulation and causes acute kidney injury through the activation of glomerular endothelial cells, the generation of microthrombi in the glomerular compartment, and the subsequent impairment of both renal perfusion and GFR (121). Post-renal causes of kidney disease (e.g. nephrolithiasis, vesicoureteral reflux, etc.) result in urinary tract obstruction, which triggers a backlog of urine into the kidney leading to injury and scarring in a process known as hydronephrosis (120).

All of these etiologies of kidney disease can give rise to CKD. Regardless of cause, the pathophysiology of CKD is similar. Cellular damage triggers the recruitment of pro-inflammatory cells to the kidney including neutrophils, monocytes, lymphocytes, and macrophages (120). These cells propagate the inflammatory response through the production of chemical mediators such as cytokines and chemokines. The parenchymal cells of the kidney undergo cell death via both apoptotic and necrotic pathways as a result of both the initial injury and resulting inflammation. In response to these processes, macrophages, fibroblasts, and epithelial cells secrete pro-fibrotic factors such as

TGFβ and CTGF in an attempt to repair the tissue damage (120). Prolonged damage and inflammation can lead to progressive interstitial fibrosis of the kidney, further lowering the GFR, and ultimately driving kidney failure (120).

38 1.8 Summary, Rationale, and Hypotheses

NLRs and inflammasome complexes are critical sensors of a variety of cell stressors and play key roles in affecting the innate immune response to these phenomena.

In the case of NLRs, this can be through both inflammasome-dependent and inflammasome-independent signaling pathways depending on the both the nature of the

NLR and the context of activating stimulus. The same is true for other pattern recognition receptor complexes, such as the non-canonical caspase-4 inflammasome, which can also feed into canonical inflammasome signaling through poorly understood mechanisms.

The purpose of this PhD was to explore the knowledge gaps that exist in the field of inflammasome-independent NLR and non-canonical caspase-4 inflammasome signaling. To accomplish this, two projects, linked by the canonical inflammasome- independent nature of the pathways they regulate, were undertaken:

1) To explore the biology of NLRP6, a non-inflammasome-forming NLRP in the

kidney under both homeostatic and pathological states (“NLRP6 Project”)

2) To investigate the activation of the non-canonical caspase-4/11 inflammasome

in a Shiga toxin model of macrophage-mediated inflammation (“Non-

Canonical Caspase-4 Inflammasome Project”)

What follows is an overview of the central hypotheses and research questions associated with each of these projects and the organization of this thesis in terms of addressing them.

39 1.8.1 NLRP6 Project

Relevant Background and Rationale:

1. NLRP6 mRNA is highly expressed in kidney tissue.

2. NLRP6 is reported to have an anti-inflammatory and anti-mitogenic role in

murine models of inflammatory disease.

3. NLRP6 has a high degree of sequence similarity with NLRP3, another NLRP

expressed in the kidney, which is known to have immunomodulatory and pro-

fibrotic functions.

It is the central hypothesis of this project that NLRP6 plays a significant role

in the pathogenesis of kidney disease through the regulation of inflammatory and/or

fibrotic signaling cascades.

To address this hypothesis, the following research questions were devised:

1. Where is NLRP6 expressed in the kidney and how is this gene regulated?

(Chapter 3)

2. What signaling pathways are regulated by NLRP6 and how do these pathways

contribute to the pathogenesis of kidney disease? (Chapter 3)

3. What are the key structure-function relationships in NLRP6 and how do these

mediate the protein’s activation and downstream effector function? How does

this compare to NLRP3, a known inflammasome-forming NLRP sharing high

sequence similarity? (Chapter 4)

40 1.8.2 Non-Canonical Caspase-4 Inflammasome Project

Relevant Background and Rationale:

1. Shiga toxins activate the NLRP3 inflammasome.

2. Cholera toxin, a related AB5 holotoxin, allows for the import of LPS into the

cell to activate the non-canonical caspase-4 inflammasome.

3. Cleaved gasdermin D has a high affinity for cardiolipin, a mitochondrial lipid.

4. Mitochondrial ROS is a known activator of the NLRP3 inflammasome.

It is the central hypothesis of this thesis that enterohemorrhagic Escherichia coli

Shiga toxins activate the non-canonical caspase-4 inflammasome and mitochondrial

reactive oxygen species upstream of NLRP3.

To address this hypothesis, the following research questions were devised:

1. Does Shiga toxin activate the non-canonical caspase-4 inflammasome?

(Chapter 5)

2. Is Shiga toxin-bound LPS (Stx/LPS) responsible for the activation of the non-

canonical caspase-4 inflammasome? (Chapter 5)

3. Does Stx/LPS induce mitochondrial ROS and does this occur as a result of

gasdermin D insertion into the mitochondrial membrane? (Chapter 5)

4. Is the non-canonical caspase-4 inflammasome-mediated generation of

mitochondrial ROS responsible for the activation of the NLRP3

inflammasome? (Chapter 5)

41 1.8.3 Contextual Importance of this Work

Until recently, the deep-rooted assumption in the literature was that all NLRPs form caspase-1-activating inflammasomes and these were the key signaling platforms underlying inflammation. The work presented in this thesis disputes this assumption. By understanding the biology of non-inflammasome-forming NLRP proteins and caspase-1- independent inflammasome signaling, the fundamental pathways that modulate inflammation in disease can be better understood and novel therapeutic agents can potentially be developed. The projects presented in this thesis contribute to the fundamental knowledge surrounding the function of pattern recognition receptors and will contribute to the elucidation of the pathogenesis of inflammatory disease.

42 Chapter Two: Materials and Methods

43 2.1 Materials

2.1.1 Reagents, Kits, and Buffers

Table 2.1 Reagents

Reagent Manufacturer Catalogue Number Complete Protease Inhibitor Sigma 4693132001 Glibenclamide (Glyburide) Sigma G2539 ICH-2 Sigma SCP0087 LEVD-FMK Enzo Life Sciences ALX-260-142-R020 Lipofectamine 2000 Thermo Fischer Scientific 11668027 LPS, E. coli 0157:H7 Alpha Diagnostic International LPS15-1 LPS, Ultrapure E. coli 0111:B4 Invivogen tlrl-eblps MitoSOX Thermo Fisher Scientific M36008 N-Acetyl-L-Cysteine Sigma A9165 Nigericin Tocris 4312 Penicillin-Streptomycin Invitrogen 15140122 PMA Sigma P1585 PolyJet Reagent SignaGen Laboratories SL100688 Polymyxin B-Agarose Sigma P1411 Protein Assay Dye Reagent Bio-Rad 5000006 Puromycin Dihydrochloride Sigma P9620 Sepharose 6B Sigma GE17-0110-01 Shiga toxin, Subunit B, Yeast LifeSpan BioSciences Inc. LS-G23305 Recombinant YVAD-CMK Sigma SML0429 Z-VAD-FMK Tocris 2163

Table 2.2 Commercial Kits

Kit Manufacturer Catalogue Number BD OptEIA TMB Substrate Reagent Set BD Biosciences 555214 BD OptEIA Human IL-1β ELISA BD Biosciences 557953 BD OptEIA Mouse IL-1β ELISA BD Biosciences 559603 CytoTox96 Non-Radioactive Cytotoxicity Assay Promega G1782 FAM-FLICA Caspase-1 Assay Kit ImmunoChemistry 97 Technologies Mouse Albumin ELISA Bethyl Laboratories E90-134 Phusion High Fidelity DNA Polymerase NEB M0530S Q5 Site Directed Mutagenesis Kit NEB E0554 QIAGEN Midi Prep Kit Qiagen 12143 QIAquick Gel Extraction Kit Qiagen 28704

44 QIAquick PCR Purification Kit Qiagen 28104 QIAshredder Qiagen 79654 RNEasy Mini Kit Qiagen 74104

Table 2.3 Cell Culture Media/Buffer

Medium/Buffer Manufacturer Catalogue Number Dulbecco’s Modified Eagle Medium (DMEM) (1X) Life Technologies 11965-092 Dulbecco’s Modified Eagle Medium (DMEM)/F12 Life Technologies 11320-082 (1X) Dulbecco’s Phosphate Buffered Saline (DPBS) (1X) Life Technologies 14190-144 Hank’s Balanced Salt Solution (HBSS) (1X) Life Technologies 14170-112 Roswell Park Memorial Institute Medium 1640 Life Technologies 11875-093 (RPMI 1640) (1X) Roswell Park Memorial Institute Medium 1640 (1X) Life Technologies 11835-030 (RPMI 1640) (Without Phenol Red)

Table 2.4 Buffers, Solutions, and Supplemented Media

Buffer/Solution Composition Blocking Buffer for 5% (w/v) non-fat milk or 5% (w/v) bovine serum albumin Immunoblotting (BSA) in PBS-T or TBS-T respectively HEK293T Media DMEM media supplemented with 10% (v/v) FBS and 100 U/mL penicillin/streptomycin Lower Gel Buffer 1.5 M Tris base and 0.4% SDS, pH to 8.8 NP-40 Lysis Buffer (1%) 1% IGEPAL (v/v), 1 M Tris pH 7.5, 150 mM NaCl, 5 mM EDTA pH 8.0, 1 mM Na3VO4, and 1 x tablet Complete Protease Inhibitor Cocktail (Roche) Phosphate Buffered 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 1.8 Saline (PBS) (1X) mM KH2PO4, pH to 7.4 Phosphate Buffered 1X PBS with 0.5% (v/v) Tween-20 Saline with Tween-20 (PBS-T) (1X) RIPA Buffer 140 mM NaCl, 10 mM Tris-Cl (pH 8.0), 1 mM EDTA, 0.5 mM EGTA, 1% (v/v) Triton X-100, 0.1% (w/v) sodium deoxycholate, 0.1% (w/v) SDS, and 1 x tablet Complete Protease Inhibitor Cocktail (Roche) Running Buffer (1X) 25 mM Tris base, 192 mM glycine, 1% (w/v) SDS SDS Sample Buffer (3X) 37.5% (v/v) stacking gel buffer pH 6.8, 34.4% (v/v) glycerol, 6% (v/v) SDS, and 0.03% (v/v) phenol red Stacking Gel Buffer 0.5 M Tris base and 0.2% SDS, pH to 6.8 THP-1 Media RPMI 1640 media supplemented with 10% (v/v) FBS, 100

45 U/mL penicillin/streptomycin, 1mM sodium pyruvate, and 0.05mM β-mercaptoethanol Transfer Buffer (1X) 25 mM Tris base, 192 mM glycine, +/- 1% (w/v) SDS, in 1:4 ratio of 100% ethanol to water. Tris-borate-EDTA (TBE) 89 mM Tris base, 89 mM boric acid, and 2 mM EDTA pH (1X) 8.0 Tris Buffered Saline 50 mM Tris pH 7.4, 150 mM NaCl (TBS) (1X) Tris Buffered Saline with 1X TBS with 0.1% (v/v) Tween-20 Tween-20 (TBS-T) (1X) Urea Buffer 7M urea, 2M thiourea, 4% (w/v) CHAPS, 1% (w/v) DTT, 2% (v/v) carrier ampholytes (pH 3-10), and 1 x tablet Complete Protease Inhibitor Cocktail (Roche)

2.1.2 Antibodies

Table 2.5 Primary Antibodies for Immunoblotting and Flow Cytometry

Antibody Target Manufacturer Catalogue Number Dilution α-Smooth Muscle Actin Sigma A2547 1:1000 ASC Adipogen AG-25B-0006-C100 1:1000 ATF6 Abcam ab122897 1:500 β-Tubulin Sigma T0198 1:1000 Caspase-1 Santa Cruz sc-622 1:250 Caspase-4 MBL M029-3 1:500 Caspase-8, Cleaved Cell Signaling 9496 1:1000 CD11b Abcam ab75476 1:1000 E-Cadherin BD Biosciences BD610181 1:1000 FLAG Sigma F3165 1:1000 GAPDH Cell Signaling 2118 1:1000 Gasdermin D Sigma G7422 1:500 GFP Thermo Fisher A-11122 1:1000 Scientific IRE1α Cell Signaling 3294 1:1000 IRE1α, S724 Abcam ab124945 1:1000 Phosphorylated IL-1β, Full Length R&D AF-201 1:1000 IL-1β, Cleaved Cell Signaling 12242 1:1000 Escherichia coli LPS Abcam ab211144 1:1000 NLRP3 Adipogen AG-20B-0014-C100 1:1000 NLRP6, Human Adipogen AG-20B-0046-C100 1:1000 NLRP6, Human R&D MAB9145 1:1000 NLRP6, Mouse Santa Cruz sc-50635 1:500 PERK Cell Signaling 3192 1:1000

46 Shiga Toxin 2, Subunit B Abcam ab101838 1:1000 TLR4 Santa Cruz sc-293072 1:1000 VDAC Cell Signaling 4866 1:1000

Table 2.6 Primary Antibodies for Immunohistochemistry/Immunocytochemistry

Antibody Target Manufacturer Catalogue Number Dilution ASC Adipogen AG-25B-0006-C100 1:100 CD77 (Gb3) Thermo Fischer Scientific MA5-16505 1:100 E-Cadherin BD Biosciences BD610181 1:100 GM130 Abcam ab52649 1:100 NLRP3 Adipogen AG-20B-0014-C100 1:100

Table 2.7 Secondary Antibodies

Antibody Target Manufacturer Catalogue Dilution Number Chicken IgY, HRP Conjugated Calbiochem 401520 1:5000 Goat IgG, HRP Conjugated Jackson 705-036-147 1:5000 ImmunoResearch Mouse IgG, AlexaFluor 488 Thermo Fischer A-11029 1:600 Conjugated Scientific Mouse IgG, AlexaFluor 568 Thermo Fischer A-11004 1:600 Conjugated Scientific Mouse IgG, HRP Conjugated Jackson 115-035-003 1:5000 ImmunoResearch Rabbit IgG, AlexaFluor 568 Thermo Fischer A-11011 1:600 Conjugated Scientific Rabbit IgG, HRP Conjugated Jackson 111-035-003 1:5000 ImmunoResearch Rat IgG, AlexaFluor 488 Thermo Fischer A-11006 1:600 Conjugated Scientific Rat IgG, HRP Conjugated Jackson 112-035-003 1:5000 ImmunoResearch

47 2.1.3 Oligonucleotides

Table 2.8 Oligonucleotides used for Cloning and Site-Directed Mutagenesis

Construct Direction Sequence (5’ to 3’) IRE1α CRISPR Forward CACCGACACCAAAACCCGAGAGCTC gRNA Reverse AAACGAGCTCTCGGGTTTTGGTGTC GSDMD Forward CACCGACCAGCCTGCAGAGCTCCAC CRISPR gRNA Reverse AAACGTGGAGCTCTGCAGGCTGGTC NLRP3 CRISPR Forward CACCGCTGCAAGCTGGCCAGGTACC gRNA Reverse AAACGGTACCTGGCCAGCTTGCAGC NLRP3 C409A Forward CTTCACCATGGCCTTCATCCCCCTG Reverse AGGACCTCGTTCTCCTGAATC NLRP3 C415A Forward CCCCCTGGTCGCCTGGATCGTGTG Reverse ATGAAGCACATGGTGAAGAG NLRP3 C419A Forward CTGGATCGTGGCCACTGGACTGAAAC Reverse CAGACCAGGGGGATGAAG NLRP3 P412A Forward GTGCTTCATCGCCCTGGTCTGCT Reverse ATGGTGAAGAGGACCTCGTTC NLRP3 V414A Forward CATCCCCCTGGCCTGCTGGATCG Reverse AAGCACATGGTGAAGAGGACC NLRP3 W416A Forward CCTGGTCTGCGCCATCGTGTGCAC Reverse GGGATGAAGCACATGGTG NLRP3 WA Forward CGCCATCCTGGCCAGGAAGATGATGTTG Reverse GCGGCAATCCCTGCCGCCCCCTG NLRP3 WB Forward CGCCGCCCTGCAAGGTGCCTTTGAC Reverse AAGCCGGCCATGAGGAAGAGGATTCTGG NLRP6 Forward GACCTGTCGCGCACGTCC Δ379-406 Reverse GTTCTCCTTCACGAAGCGGTAG NLRP6 C385A Forward GTTCGCGCTGGCCTTCGTGCCC Reverse AGCGTCTCGTTCTCCTTC NLRP6 C391A Forward GCCCTTCGTGGCCTGGATCGTGTG Reverse ACGAAGCACAGCGCGAAC NLRP6 C395A Forward CTGGATCGTGGCCACCGTGCTG Reverse CACACGAAGGGCACGAAG NLRP6 P388A Forward GTGCTTCGTGGCCTTCGTGTG Reverse AGCGCGAACAGCGTCTCG NLRP6 V390A Forward CGTGCCCTTCGCCTGCTGGATCGTGTG Reverse AAGCACAGCGCGAACAGC NLRP6 W392A Forward CTTCGTGTGCGCCATCGTGTGCACCGTG Reverse GGCACGAAGCACAGCGCG

48 NLRP6 WA Forward GGCCATGGCGGCCAAAAAGATCCTGTACGAC Reverse GCGGCGATGCCCGCCGGGCCCTG NLRP6 WB Forward GGCCGCCCTGCCGGCGCTGGGGGGC Reverse GCGCCGGCCAGGATGAAGAGCAGCCGCTGCGG NLRP6 PYD Forward CGCAAGCTTATGGACCAGCCAGAGGCCCCC Reverse CGCGGATCCGCCTGCAGCCGCCGCTCCTGGAG NLRP6 NACHT Forward CGCAAGCTTATGCTGACCGTGGTGCTGCAGGGC Reverse CGCGGATCCGCCAGCAGGTAGGACAGTGCCGC NLRP6 Forward CGCAAGCTTATGGAGAAGGAACTGGAGCAACT LRR 1-5 G Reverse CGCGGATCCGCCTTTGCTCTCTTCACAGCCTG NLRP6 Forward CGCAAGCTTATGCTGTGCCATCTGAGCAGCCTC LRR 2-5 Reverse CGCGGATCCGCCTTTGCTCTCTTCACAGCCTG NLRP6 Forward CGCAAGCTTATGGACCAGCCAGAGGCCCCC PYD-NACHT Reverse CGCGGATCCGCCAGCAGGTAGGACAGTGCCGC NLRP6 Forward CGCAAGCTTATGCTGACCGTGGTGCTGCAGGGC NACHT-LRR Reverse CGCGGATCCGCCTTTGCTCTCTTCACAGCCTG NLRP6 Forward GAGGACGGCGGGGTGCCC ΔNACHT Reverse CGGCCGCCGGCCCTCCTC

Table 2.9 Oligonucleotides used for PCR, RT-PCR, and qRT-PCR

Target Direction/Probe Sequence (5’ to 3’) mCol1A1 Forward GGAAGAGCGGAGAGTACTGG Reverse TTGCAGTAGACCTTGATGGC mGAPDH Forward CCATCACCATCTTCCAGGAGC Reverse GCCTGCTTCACCACCTTCTTG mKIM-1 Forward CGTGGCTATCACCAGGTACATACT Reverse GCGTTCTGCAAAGCTTCAATC Probe TCTCTAAGCGTGGTTGC mNGAL Forward ACTGGATCAGAACATTTGTTCCAA Reverse CCACTTGCACATTGTAGCTCTGTA Probe TATGCACAGGTATCCTC mNLRP3 Forward AGAGCCTACAGTTGGGTGAAATG Reverse CCACGCCTACCAGGAAATCTC Probe CGTGCCTTAGAAGCG hNLRP6 Forward GACCCTCAGTCTGGCCTCTGT Reverse TCCGGCTTTGCTCTCTTCAC Probe TGAGCGAGCAGTCAC mNLRP6 Forward GTGAGACAATGACTACCCCGAAAT

49 Reverse GTCTCGGCAAACTGCATCAG Probe TCTGATCTTGTCACACTGC mNPHS2 Forward TCCGTCTCCAGACCTTGGAA Reverse TGCAGACAGCGTCTATCTCCATTA Probe TCCATGAGGTGGTAACCA hXBP-1 Forward GAAACTGAAAAACAGAGTAGCAGC Reverse GCTTCCAGCTTGGCTGATG

2.2 Methods

2.2.1 Animal Studies

2.2.1.1 Anti-Glomerular Basement Membrane Nephrotoxic Serum Nephritis (NTS)

All animal studies were conducted with the approval of, and in accordance with, the Animal Care Committee guidelines at the University of Calgary, Canada. All mice were bred under double barrier conditions and housed in a pathogen-free facility.

Polyclonal sheep anti-mouse glomerular basement membrane serum was obtained from

Laurent Mesnard, heat inactivated, and mice were treated as previously described (122).

In brief, eight- to ten-week old female Nlrp6 -/- mice (on a C57Bl/6 background, as generated in Chen et al. 2011 (79)) or Nlrp6 +/+ littermate control mice were weighed and injected intravenously (via tail vein injection) with 18 µL/g of the nephrotoxic serum, with the dose divided between days one and two. Weights were measured and urine collected on days zero, four, seven, and ten of the treatment course using metabolic cages. On day ten, the mice were sacrificed and the kidneys were harvested.

2.2.1.2 Unilateral Ureteral Obstruction (UUO)

Healthy male mice (8-10 weeks old), of the Nlrp6 +/+ and Nlrp6 -/- genotypes, underwent left unilateral ureteral obstruction or sham surgery. Mice were anesthetized with intraperitoneal injections of 115 mg/kg ketamine hydrochloride (Ketaset, Wyeth

Canada) and 12 mg/kg xylazine (Rompun, Bayer) along with inhalation anesthetic

50 isoflurane (Fresenius Kabi). A 1.5 cm incision through the disinfected skin and peritoneal membrane was made in the lower left quadrant of the abdominal region. The left ureter was isolated using blunt dissection and a 3-0 silk suture (Ethicon) was used to ligate the ureter. The peritoneal membrane was closed with silk sutures and the skin stapled shut with stainless steel surgical wound clips (Reflex 9, Harvard Apparatus). Mice were allowed to recover under a warming lamp and received food, water and analgesia. Sham surgeries were conducted in the exact same manner but the isolated left ureter was not ligated.

2.2.1.3 Tissue Preparation

Following the time course for either the NTS or UUO models, the mice were sacrificed and both kidneys were isolated. These are identical in the NTS model, whereas only one kidney is affected by ligation in the UUO model. The contralateral kidney thus serves as an additional negative control, alongside sham ligation. The kidneys were then divided into thirds. One third was fixed in 10% buffered formalin, before being sent to

Calgary Lab Services (CLS) for paraffin embedding, sectioning, and staining. Another third was flash frozen in OCT compound (Tissue-Tek, Sakura), and stored at -80°C for cryosectioning. The final third was sliced into small 1 mm2 pieces, flash-frozen with dry ice/ethanol, and stored at -80°C for protein extraction and mRNA isolation.

2.2.1.4 Mechanical Glomerular Isolation

For murine glomeruli, six to ten kidneys were harvested freshly from six to eight- week old, wild type C57Bl/6 mice. For human glomeruli, a one square inch piece of nephrectomy or partial nephrectomy tissue was obtained from Rockyview General

Hospital with appropriate consent. In either case, the renal capsule was removed, the

51 cortical regions were isolated, and the cortical kidney tissue was finely chopped in a 10 cm cell culture dish containing Hank’s Balanced Salt Solution (HBSS). The finely chopped pieces of kidney cortex were then transferred to a 100 µm sieve overlying a 50 mL conical, where they were crushed through the sieve using the plunger from a 10 mL syringe. The contents of the sieve were then washed several times with HBSS, collecting the flow through containing the glomeruli onto a 70 µm sieve. The collected glomeruli were then washed three times with HBSS, before being transferred to a new conical by washing them off the sieve with 8 mL HBSS. The glomeruli were then decapsulated by passing them through a 21.5-G needle twice. The decapsulated glomeruli were then spun down at 250 x g, and a sample was viewed by light microscopy to confirm the glomerular isolation. The supernatant was removed, and Buffer RLT was added to begin the process of RNA extraction.

2.2.1.5 Laser-Capture Microdissection

Snap frozen tissue in OCT was prepared for laser-capture microdissection (LCM).

8-µm thick sections were fixed with 75% ethanol and stained with the HistoGene

(Applied Biosystems, Foster City, CA, USA) under RNase-free conditions. After complete dehydration, the PixCell II LCM System (Arcturus Engineering, Mountain

View, CA, USA) was used to obtain laser captures. Samples were then stored for mRNA isolation.

2.2.2 Assays

2.2.2.1 Bradford Assay

The assay was completed in 96-well plates. Known BSA (bovine serum albumin) standards along with the diluted (1:10) experimental protein samples (10% of the final

52 volume) were mixed with the Bradford dye solution (20% (v/v) Bradford dye (Bio-Rad),

70% (v/v) ddH2O) and read at 595 nm. Unknown protein concentrations were calculated using a standard curve produced from the standard BSA protein samples.

2.2.2.2 Enzyme-Linked Immunosorbent Assay (ELISA)

Human and mouse IL-1β ELISA kits (BD Biosciences) were used to detect IL-1β secreted into the supernatant by THP-1 cells and BMDMs treated with various compounds, as per the manufacturer’s instructions. Briefly, 96-well plates were coated with 100 µL of the IL-1β capture antibody (1:250) overnight at 4°C in coating buffer (0.1

M sodium carbonate, pH 9.5). Subsequent steps were completed at room temperature.

Wells were washed 3X with 0.05% Tween-20 in 1X PBS. Wells were then blocked for 1 hour with 200 µL of blocking buffer (10% FBS in 1X PBS, pH 7.0), wells were washed

3X, then incubated with 100 µL of IL-1β standard (made at 1000 pg/mL, 500 pg/mL, 250 pg/mL, 125 pg/mL, 62.5 pg/mL, 31.3 pg/mL and 15.6 pg/mL concentrations in blocking buffer) or experimental supernatant samples (diluted 1:10) for 2 hours. Plates were washed 5X then incubated with 100 µL of biotin-conjugated secondary antibody (1:1000) for 1 hour, followed by 5 washes and then incubation for 30 minutes with 100 µL of

HRP- conjugated to streptavidin (1:250). Well were washed 7X then developed with 100

µL of TMB substrate (BD Biosciences) for 30 minutes in the dark. The reaction was stopped with 50 µL of 1 M H3PO4. Plates were then read at 450 nm in the Bio-Rad

Benchmark Microplate Reader.

Mouse albumin ELISA kits (Bethyl Laboratories) were used to quantify the albuminuria generated in the NTS model. Briefly, 96-well plates were coated with 100

µL of the mouse albumin capture antibody (1:100) for 1 hour in coating buffer (0.05 M

53 sodium carbonate, pH 9.6) at room temperature. Wells were washed 5X with 0.05%

Tween-20 in 1X TBS pH 8.0. Wells were then blocked for 30 minutes with 200 µL of blocking buffer (1% BSA in 1X TBS, pH 8.0), wells were washed 5X, then incubated with 100 µL of mouse albumin standard (made at 500 ng/mL, 250 ng/mL, 125 ng/mL,

62.5 ng/mL, 31.3 ng/mL, 15.6 pg/mL, and 7.8 ng/mL concentrations in blocking buffer with 0.05% Tween-20) or experimental urine samples (diluted 1:4000) for 1 hour. Plates were washed 5X then incubated with 100 µL of HRP-conjugated secondary antibody

(1:100,000) for 1 hour, followed by 5 washes. Well then developed with 100 µL of TMB substrate (BD Biosciences) for 15 minutes in the dark. The reaction was stopped with 50

µL of 1 M H3PO4. Plates were then read at 450 nm in the Bio-Rad Benchmark Microplate

Reader.

2.2.2.3 Immunoblotting

Protein samples were separated on 8%, 10%, 12%, or 15% SDS-PAGE gels (depending on protein size) under reducing conditions at 180V. The proteins were then transferred onto 0.2 µm pore nitrocellulose membranes (Hybond ECL, Amersham GE Healthcare

Life Sciences) at 100V for either 45 minutes (in SDS-free transfer buffer) or 1 hour (in

SDS-containing transfer buffer), again depending on the size of the proteins in question.

The membranes were stained with Ponceau red dye to confirm successful transfer and then blocked for 1 hour using either 5% (w/v) dry, non-fat skim milk or 5% (w/v) BSA diluted in 1X PBST (containing 0.5% (v/v) Tween-20) or 1X TBST (containing 0.1%

(v/v) Tween-20) respectively. Following the blocking step, the membranes were incubated at 4°C overnight with the primary antibody in appropriate buffer. Following washes with either PBST or TBST, membranes were incubated for 1 hour at room

54 temperature with the secondary antibody diluted 1:5000 in appropriate blocking buffer.

After the final wash, proteins were visualized using enhanced chemiluminescent (ECL) western blotting detection reagents (Amersham, GE Healthcare, Mississauga, ON) and images were captured using a ChemiDoc MP imaging device (Bio-Rad). Densitometry analysis was accomplished using Image Lab software version 5.1 build 8 (Bio-Rad).

2.2.2.4 Lactate Dehydrogenase Assay

Lactate dehydrogenase (LDH) release into the supernatant by dying cells was measured using the CytoTox 96 Non-Radioactive Cytotoxicity Assay (Promega). This was done to calculate percent cell death relative to a complete cell lysis control well.

Briefly, THP-1 cells were plated in 24 well plates and subjected to various treatments for a variety of time points in 500 µL of THP-1 media in a 37 degree Celsius incubator. 45 minutes prior to the end of the time point, 55 µL of 10X Lysis Buffer (included in the kit) was added to the complete cell lysis control wells and the plate was returned to the incubator. Following the completion of the time point, 50 µl of supernatant was collected from each well and transferred in triplicate to a 96 well plate. 50 µL of Assay

Diluent/Substrate Mix (both provided in the kit) was added to each well and the 96 well plate was allowed to incubate at room temperature in the dark for 30 minutes, before the addition of 50 µL of Stop Solution to end the reaction. The 96 well plate was then analyzed by spectrophotometry at 490 nm and percent cell death was calculated by the following formula: % Cell Death = 100 x (Experimental LDH Release

(OD490)/(Maximum LDH Release (OD490)).

2.2.2.5 Reconstituted Inflammasome Expression

55 HEK293T cells were grown to 70% confluence in 24-well plates and were provided with fresh media the day of experimentation. Using PolyJet reagent, cells were transfected with 15 ng/well of wild type or mutant NLRP3-GFP, 5 ng/well of FLAG-

ASC, 5 ng/well of FLAG-pro-caspase-1 and 150 ng/mL FLAG-pro-IL-1β. Transfection was allowed to proceed for 6 hours or overnight, with immunoblotting/ELISA or LDH assays used as a readouts of inflammasome activity respectively. Mutant NLRP3 results were compared to the wild type control wells. Transfections omitting key inflammasome components (e.g. ASC, NLRP3, pro-caspase-1, etc.) served as negative controls.

2.2.2.6 NLRP Oligomerization Assay

HEK293T cells were cultured to a confluency of 70% in 60 mm cell culture plates. Cells were then co-transfected with 3 µg mutant GFP-tagged NLRP3/6 and 3 µg wild type FLAG-tagged NLRP3/6 via calcium phosphate precipitation. The media was changed after 6 hours, and the transfected cells were allowed to incubate at 37°C for 48 hours.

Following this incubation, cells were washed with phosphate-buffered saline and lysed with 200 µL of 1% NP-40 lysis buffer (50 mM Tris pH 7.5, 150 mM NaCl, 5 mM

EDTA pH 8, Roche Complete Protease Inhibitor Tablet). Lysates were spun at 13,000

RPM in a microcentrifuge and the supernatants were transferred to Eppendorf tubes containing 10 µL of Sepharose 6B beads (Sigma Aldrich). Lysates were rotated for 1 hour at 4°C after which the samples were spun at 8000 RPM for 5 minutes (to remove proteins that bind non-specifically to the Sepharose beads). Supernatants were then divided into two fractions: 30 µL was collected for immunoblotting of the input lysate, with the remaining 170 µL transferred to new Eppendorf tubes containing 10 µL of

56 Protein G beads (GE Healthcare), 10uL lysis buffer, and 0.5 µg FLAG antibody (Sigma

Aldrich) each. Bead-containing samples were rotated at 4°C overnight, after which they were spun at 2000 RPM in a microcentrifuge. The supernatant was aspirated, and the beads were washed five times with 500 µL of lysis buffer.

Following the wash steps, 50 µL of 3XSDS sample buffer was added to each bead-containing tube (15 µL to the aforementioned input lysates), and the samples were incubated for 10 minutes at 95°C. The samples, both pre and post-IP, were subjected to

SDS-PAGE and immunoblotted with anti-NLRP3 or anti-NLRP6 antibodies (Adipogen).

Densitometry was performed and the quotient of GFP-tagged NLRP3/6 pulled down by

FLAG-tagged NLRP3/6 was calculated. Percent oligomerization was calculated by dividing this quotient by that of the positive control – wild type GFP-tagged NLRP3/6 pulled down by wild-type FLAG-tagged NLRP3/6. Co-immunoprecipitation experiments using cell lysates from untransfected, GFP-transfected, NLRP3/6-GFP alone or FLAG-

NLRP3/6 alone served as negative controls.

2.2.3 Cell Culture

2.2.3.1 Cell Line Culture

The human myelogenous leukemia THP-1 cell line was purchased from the

American Type Culture Collection (ATCC, Manassas, Virginia, United States). THP-1 cells were cultured in RPMI 1640 media (Gibco/Thermo Scientific) supplemented with

10% (v/v) heat-inactivated fetal bovine serum (FBS), 0.05 mM β-mercaptoethanol, and 1 mM sodium pyruvate. Cells were maintained in cell suspension in a 37 degree Celsius,

5% CO2, humidified incubator. Cells were differentiated with 100 nM phorbol-12- myristate-13-acetate (PMA) (Sigma-Aldrich) for 16-18 hours prior to experiments.

57 The human embryonic kidney (HEK293T) cell line was purchased from the

American Type Culture Collection (ATCC, Manassas, Virginia, United States).

HEK293T cells were cultured in DMEM media (Gibco/Thermo Scientific) supplemented with 10% (v/v) heat-inactivated FBS. Cells were maintained on uncoated 10 cm plates in a 37 degree Celsius, 5% CO2, humidified incubator. When confluent, cells were passaged

1:10 onto new cell culture plates.

2.2.3.2 Primary Cell Culture

Wild type C57Bl/6 mice were housed under standard conditions. Primary bone marrow-derived macrophages (BMDM) were isolated from mice between the ages of 8-

12 weeks of age. Bone marrow was collected from the femurs and tibia of the mice and was stored on ice in Hanks' Balanced Salt Solution (HBSS) (Gibco). Cells were centrifuged at 230xg for 5 minutes and subjected to two washes with HBSS.

Macrophages were resuspended in DMEM (Gibco) media containing 10% L929-cell medium, 1% penicillin streptomycin, 1% L-glutamine, and 10% FBS. BMDM were maintained for 4 days replacing the media every two days. On day 4, confluent macrophages were primed with 100 ng/mL of ultrapure LPS (Sigma-Aldrich) for 4 hours prior to initiating the experiments.

Human peripheral blood monocytes (PBMCs) and polymorphonuclear cells were isolated from healthy volunteers (20 mL) or from the pooled blood of 5-10 mice.

Heparinized blood was layered in 5 mL aliquots on 2.5 mL Histopaque 1119 (Sigma-

Aldrich) and 2.5 mL Histopaque 1077 gradient in 15 ml tubes and centrifuged for 30 minutes at 700 x g. The mononuclear and PMN layers were collected, pooled, and washed x3 in sterile PBS. After the final wash, the mononuclear cells were resuspended

58 in complete RPMI, plated at a density of 6.0 x 105 cells/well in a 12 well tissue culture plate. Cells were primed with 100 ng/mL ultrapure LPS for 4 hours before initiating experiments. All procedures on mice and humans were approved and performed under the guidelines set forth by the Conjoint Health Research Ethics Board and the Animal

Care Committee at the University of Calgary.

Primary renal tubular epithelial cells were isolated from mice between the ages of

6-12 weeks. Mice were sacrificed and the kidneys removed. Under sterile conditions the capsules surrounding the kidneys were detached and the renal cortex removed with a sterile scalpel. Renal cortex tissues were placed in 1.5 mg/ml collagenase/HBSS (Sigma) solution and incubated at 37°C with 5% CO2 for 60 minutes. Tissue was homogenized using sterile microscope slides and then passed through 70 µm nylon filters (BD Falcon).

The cell suspension was spun down for 5 minutes at 230 x g to remove excess collagenase and washed twice more in 1X HBSS. The cell pellet was resuspended in K1 media (10% (v/v) FBS, 1% (v/v) penicillin-streptomycin, 1% (v/v) hormone mix, 25 ng/ml mouse EGF, 25 mM HEPES (pH 7.4 in DMEM/F12) and plated onto tissue culture plates for 1 hour at 37°C, 5% CO2. The cell suspension was then re-plated onto collagen

IV-coated plates and allowed to grow overnight at 37°C, 5% CO2). Cells were washed and media changed the next day. Cells were maintained in K1 media and allowed to grow to confluency before passage onto experimental plates.

Human tubular epithelial cells were isolated from non-diseased nephrectomy samples from patients with renal cell carcinoma. Nephrectomy samples were placed in clean HBSS on ice for transport before isolation. Subsequent steps were performed in a fume hood under sterile conditions. The capsule and medulla were dissected away from

59 the cortex. Minced renal cortex tissues were then placed in a 1.5 mg/ml collagenase/HBSS (Sigma) solution and incubated at 37°C with 5% CO2 for 90 minutes.

Tissue was homogenized using two sterile microscope slides and then passed successively through a series of autoclaved metal sieves (250 µm, 106 µm and 75 µm).

The filtered cell suspension was then spun down for 10 minutes at 230 g to remove excess collagenase and washed 2 twice more in 1X HBSS. The cell pellet was resuspended in HPTC media (10% FBS (v/v), 1% (v/v) penicillin-streptomycin, 1% (v/v) hormone mix, 25 ng/ml human EGF, 25 mM HEPES (pH 7.4 in DMEM/F12) and plated onto uncoated tissue culture plates for 1 hour at 37°C, 5% CO2. After 1 hour, the cell suspension was re-plated onto collagen IV coated plates and allowed to grow overnight at

37°C, 5% CO2. Cells were washed and media changed the next day. Cells were maintained in HPTC media and allowed to grow to confluency before passage onto collagen coated experimental plates.

Renal fibroblasts were obtained from both human and murine kidney by allowing them to grow out of the aforementioned tubular epithelial cell cultures under identical culture conditions. If allowed to, fibroblasts will eventually overtake the plates.

2.2.3.2 Generation of CRISPR THP-1 Cell Lines

NLRP3 -/-, GSDMD -/-, and IRE1α -/- THP-1 cells were produced using the lentiCRISPRv2 shuttle plasmid (Addgene plasmid # 52961, a gift from Feng Zhang)

(123). The lentiCRISPRv2 included three expression cassettes: the Cas9 nuclease, puromycin resistance, and the guide RNA scaffold, into which a pair of annealed oligonucleotides complementary to bases 767-787 in the first exon of the NLRP3 gene, bases 225-245 in the second exon of the GSDMD gene, and bases 613-632 in the seventh

60 exon of the IRE1α, and were cloned using BsmI digestion sites. The following targeting oligonucleotides were synthesized: NLRP3: 5’-

CACCGCTGCAAGCTGGCCAGGTACC-3’, 5’-

AAACGGTACCTGGCCAGCTTGCAGC-3’; GSDMD: 5’-

CACCGACCAGCCTGCAGAGCTCCAC-3’, 5’-

AAACGTGGAGCTCTGCAGGCTGGTC-3’; IRE1α: 5’-

CACCGACACCAAAACCCGAGAGCTC-3’, 5’-

AAACGAGCTCTCGGGTTTTGGTGTC-3’. Oligonucleotides were annealed by the addition of 0.1 nmol of each to 0.5 µL of polynucleotide kinase in 10x T4 Ligation Buffer

(NEB) at 37°C for 30 minutes, 95°C for 5 minutes, followed by ramping down the temperature at 5°C/min to 25°C. The lentiCRISPRv2 plasmid was then digested with

BsmBI, and dephosphorylated with alkaline phosphatase (NEB) for 30 minutes at 37°C.

The annealed oligonucleotides were then diluted 1:200 into sterile water and ligated into

50 ng of the dephosphorylated and BsmBI-digested lentiCRISPRv2 with T4 DNA ligase in T4 DNA ligase buffer for 10 minutes at room temperature.

HEK293T cells (8 x 10 cm plates) were transfected using calcium phosphate with the NLRP3, GSDMD, or IRE1α targeting lentiCRISPRv2 construct (15 µg), viral envelope plasmid pCMV-VSVG (5.25 µg), and viral packing vector psPAX2 (9.75 µg).

The virus-containing medium was collected at 48 and 72 hours post-transfection and concentrated by ultra-centrifugation at 50,000 x g for 120 minutes. THP-1 cells were transduced with 5 µL concentrated viral particles and maintained at 37°C, 5% CO2 in complete RPMI containing 1 µg/mL puromycin for a total of 8 passages.

61 The effectiveness of the NLRP3 CRISPR gene editing was confirmed at the DNA and protein level using PCR and immunoblotting. For GSDMD and IRE1α CRISPRs, confirmation at the protein level was performed by immunoblotting.

2.2.4 Cloning and Polymerase Chain Reaction (PCR)

2.2.4.1 Cloning

Human NLRP6 cDNA was purchased from Origene and cloned into a pAcGFP-

N1 vector (Addgene). For the conventional cloning, Phusion High Fidelity Polymerase

(NEB) was used to clone the desired sections of NLRP6, flanked by restriction sites for

HindIII and BamHI respectively, as per the manufacturer’s instructions. Following PCR with appropriate primers, the products were run through an agarose gel and extracted using a QiaQuick Gel Extraction Kit (Qiagen). The products and the pAcGFP-N1 vector were then subjected to restriction digestion with HindIII and BamHI (NEB), as per the manufacturer’s instructions. The digested products and vector were then run through a second agarose gel, extracted, and ligated using T4 DNA ligase (NEB) per the manufacturer at a 3:1 insert-to-vector ratio. The ligated vectors were then transformed into NEB5α competent E. coli cells (NEB) and grown on kanamycin-containing LB-agar plates. Kanamycin-resistant colonies were then expanded in LB broth and Midipreps

(Qiagen) performed to obtain purified plasmid DNA encoding the various constructs. All generated constructs were sequenced prior to use to ensure sequence accuracy.

2.2.4.2 Quantitative Real Time PCR (qRT-PCR)

mRNA was isolated from 105-106 cells or 20-30 mg of frozen tissue with the

RNeasy Mini kit (Qiagen) as per the manufacturer's instructions. Reverse transcription was carried out using 500 ng of mRNA, random primers (3 µg/µL) (Invitrogen), and M-

62 MLV reverse transcriptase (200 U/µL) (Invitrogen) as per the manufacturer's protocol.

For probe chemistry, target gene reactions were formed using 10 µL 2X SsoAdvanced

Universal Probes Supermix (BioRad), 900 nM of each primer, 200 nM probe and 1 µL of cDNA template in a 20 µL reaction. For SYBR Green chemistry, target gene reactions were formed using 10 µL 2X SsoAdvanced Universal SYBR Green Supermix (BioRad),

500 nM of each primer, and 1 µL cDNA template in a 20 µL reaction. Amplification was performed in 96-well reaction plates using the CFX96 Touch™ Real-Time PCR

Detection System (BioRad) and results were analyzed using the CFX Manager software.

- Ct The results are expressed as fold change (2 ΔΔ ) relative to control samples and were replicated at least three times. A comprehensive list of the oligonucleotides and probes used for the qRT-PCR reactions can be found in Table 2.9. All of the probes used are 6-

FAM/MGB (Applied Biosystems). The 20X 18S rRNA FAM/MGB Probe (Applied

Biosystems) was used as the endogenous control for probe chemistry, and GAPDH was used for SYBR Green chemistry.

2.2.4.3 Site-Directed Mutagenesis

All site-directed mutagenesis was conducted using the Q5 Site-Directed

Mutagenesis Kit from NEB, as per the manufacturer’s instructions. All primers are listed in Table 2.8 and were designed using the NEBaseChanger website. Following induction of mutations, the mutated vectors were then transformed into NEB5α competent E. coli cells (NEB) and grown on kanamycin-containing LB-agar plates. Kanamycin-resistant colonies were then expanded in LB broth and Midipreps (Qiagen) performed to obtain purified plasmid DNA encoding the various constructs. All generated constructs were sequenced prior to use to ensure sequence accuracy.

63 2.2.4.4 XBP-1 Splicing Assay

The splice state of XBP-1 was analyzed as previously described (124). Briefly, mRNA was isolated from PMA-differentiated THP-1 macrophages, treated with nigericin, Stx2, or tunicamycin for 1, 3 or 6 hours, using a RNeasy kit (Qiagen), as per the manufacturer’s instructions. mRNA was reverse-transcribed to cDNA using M-MLV reverse transcriptase (Invitrogen), as per the manufacturer’s instructions. This cDNA was then used as a template for a PCR reaction, amplifying the XBP-1 transcript from nucleotide 544 to nucleotide 820. The following primers were used: 5’-

GAAACTGAAAAACAGAGTAGCAGC -3’ and 5’- GCTTCCAGCTTGGCTGATG -

3’. The amplified PCR product was then purified, quantified by NanoDrop (to ensure equal loading) and subjected to restriction digestion with PstI (NEB) for 1 hour at 37°C.

The digested PCR products were then separated on a 2% agarose gel containing SYBR

Safe dye and visualized using a ChemiDoc imaging device (BioRad).

2.2.5 Flow Cytometry

2.2.5.1 Flow Cytometric Analysis of CD77 Expression

Adhered, differentiated human THP-1 cells and murine BMDMs were washed in

PBS and treated with 0.05% (v/v) ethylenediaminetetraacetic acid (EDTA) for 10 minutes. Cells were gently removed and washed x3 in PBS supplemented with 0.2% (v/v)

Tween-20. A total of 5 x 105 murine cells were obtained and incubated with rat anti- human CD77 antibody (Thermo Scientific) or rat IgM isotype control (Abcam,

Cambridge, UK) at a final concentration of 10 µg/mL diluted in 3% (w/v) BSA/PBS. The cells were incubated for 45 min at 4°C, washed with PBS 0.2% (v/v) Tween and incubated with FITC-conjugated secondary mouse anti-rat antibody (Molecular Probes,

64 Eugene, OR) at a final concentration of 5 µg/mL for 45 min at 4°C in the dark. Following secondary antibody incubation the cells were washed and fixed in 4% (v/v) paraformaldehyde. Cells were washed, suspended in PBS and analyzed by flow cytometry (Becton Dickinson LSR II, Franklin Lakes, NJ) using BD FACSDivaTM version 6.1 software.

2.2.6 Imaging of Cells and Tissues

2.2.6.1 Histology

Paraffin-embedded kidney tissues were sectioned and stained with hematoxylin and eosin (H&E), Masson’s trichrome, Picrosirius red, and periodic acid-Schiff (PAS) stains using standard protocols by Calgary Lab Services (CLS), the lab of Dr. Margaret

Kelly, and by Takanori Komada respectively.

2.2.6.2 Immunocytochemistry

Differentiated THP-1 and BMDMs were grown to 70% confluency on sterile coverslips. The cells were then stimulated with 200 ng/mL of Stx2 in serum-free RPMI for 3 or 6 hours. Prior to fixation, the cells were incubated with 300 µL 1 x PBS or

FAM-YVAD-FMK (FLICA-caspase-1, ImmunoChemistry Technologies, Bloomington,

MN, USA) for 1 hour. Next, the FLICA-treated cells were washed and fixed in 4% (v/v)

PFA. Cells were incubated with NH4Cl (50 mM) for 10 min and permeabilized with 0.1%

(v/v) Triton X-100 for 5 min. The permeabilized cells were blocked using 3% (w/v) BSA for 30 min at room temperature and then incubated with primary antibodies (diluted

1:100) in blocking solution for 1 hour at room temperature. Antibodies used in these experiments were mouse anti-NLRP3 (Cryo-2, Adipogen), rabbit anti-ASC (Adipogen), rat anti-CD77 (Thermo Scientific), rabbit anti-IRE1α (Cell Signaling), mouse anti-

65 calreticulin (Abcam), rabbit anti-calreticulin (Abcam), and rabbit anti-GM130 (Abcam).

Following three sequential washes with PBS for 5 min each, the cells were incubated with secondary fluorescent antibodies (1:600, Alexa Fluor, Life Technologies, Carlsbad,

CA, USA) in 3% (w/v) BSA blocking solution for 1 hour at room temperature.

Coverslips were mounted onto slides in ProlongGold anti-fade reagent containing 4',6- diamidino-2-phenylindole (DAPI). Confocal microscopy was performed using an

Olympus IX-70 microscope equipped with Fluoview1000 system software.

To visualize Shiga toxin membrane binding, THP-1 and BMDM cells were grown as before and treated with 1 µg/mL Alexa Fluor 594-labeled Shiga toxin (Stx2-AF594) diluted in serum-free RPMI media for 20 minutes. The wells were then washed three times for five minutes each with PBS and fixed with 4% (v/v) PFA. Immunofluorescence was then conducted as previously described.

2.2.6.3 Immunohistochemistry

Immunofluorescence for CD77 was conducted in cryosections of murine kidney tissue as follows. The cryosections were fixed with 4% (v/v) PFA for 15 minutes, blocked with a 2% (v/v) normal goat serum (in PBS) blocking solution for 1 hour, and incubated overnight in rat anti-mouse CD77/A14GALT primary antibody (1:100 in blocking solution) at 4˚C. Species-matched immunoglobulin M (IgM) (1:100) was used instead of the primary antibody as a negative control. AlexaFluor 488-conjugated goat anti-rat secondary antibody (1:600 in blocking solution) was used to stain CD77. Mouse anti-mouse E-cadherin antibody (BD) (1:100 in blocking solution) and Alexa 568- conjugate goat anti-mouse secondary antibodies (1:600 in blocking solution) were used to stain tubular epithelial cells. 3X 5 minute washes with PBS were performed after primary

66 and secondary antibody incubations. Following washes, coverslips were mounted onto the slides using ProlongGold anti-fade reagent containing 4',6-diamidino-2-phenylindole

(DAPI). Confocal microscopy was performed using an Olympus IX-70 microscope equipped with Fluoview1000 system software.

2.2.6.4 Live Cell Imaging with MitoSOX

THP-1 cells were stained using MitoSOX Red mitochondrial superoxide indicator

(Molecular Probes). In brief, PMA-differentiated THP-1 macrophages adhered to a 24- well tissue culture plate were stained with 5 µM of MitoSOX reagent and incubated for

10 minutes at 37°C protected from light. Cells were gently washed with warm PBS and treated with 200 ng/mL of Stx. THP-1 cells were pretreated for 30 minutes with NAC,

LEVD, or ZVAD prior to MitoSOX staining and maintained in the cultures throughout the experimental period. Following Stx treatment, cells were immediately visualized for mitochondrial ROS production using an Incucyte Zoom automated fluorescent microscope (ESSEN Bioscience, Ann Arbor, MI, USA) over the course of 6 hours. The

ROS index value of the experimental groups is expressed as a fold increase of red- positive cells divided by total amount of cells over time, normalized to time zero of each treatment group.

2.2.7 RNASeq

2.2.7.1 RNASeq Analysis

RNASeq reads were pseudoaligned to the mouse NCBI RefSeq transcript database (125) dated January 2017, using Kallisto 0.42.4 (126). Sleuth (127) was used for differential gene expression using a linear model containing three terms: the source tissue, knockout status, and date of sample extraction (batch effect). Genes passing the

67 Likelihood Ratio Test with Benjamnini-Hochberg corrected p-values (a.k.a. false discovery rate, FDR) less than 0.05 and Wald test FDR < 0.05 were considered differentially expressed. Differentially expressed genes were annotated and analyzed for enrichment using Ingenuity Pathway Analysis (Qiagen N.V., Redwood City, CA).

RNASeq reads were also aligned to the Genome Reference Consortium’s mouse dataset

GRCm38 using bwa v0.7.15, with the resulting BAM files used for visual inspection of

UTR regions.

2.2.8 Shiga Toxin Preparation and Labeling

2.2.8.1 Shiga Toxin Preparation

Shiga toxin 1 (Stx1) and Shiga toxin 2 (Stx2) were isolated and affinity purified

K using an immobilized synthetic analog of the CD77 (GB3) receptor Synsorb-P (Alberta

Innovates Technology Futures, NuRX Services, CAN) as previously described (128, 129).

In brief, Stx1 was purified from E. coli JM101 (pJB28) that contained the Stx1 operon isolated from the genome of bacteriophage H-19B cloned into pTZ18-R in E. coli C600, which was a kind gift provided by Dr. J.L. Brunton (University of Toronto, Toronto,

ON). Stx2 was isolated from E. coli C600 containing the Stx2 bacteriophage 933W and was provided by Dr. David Acheson (Tufts University, Boston, MA, USA). Cultures were grown at 37°C and lysed using polymyxin B sulfate (0.1 mg/ml) (Sigma-Aldrich,

St. Louis, MO, USA) during the mid- to late log-phase, when the cells reached an absorbance value of 0.6 at wavelength of 540 nm. Lysates were then centrifuged at

10,000 x g for 20 minutes to sediment bacterial cell bodies and the cell free supernatant was collected. Synsorb-PK was subsequently added to the cell-free culture supernatant solution and incubated at 37°C on a shaker at 115 rpm. The mixture was then washed in

68 250 mM NaCl (pH 3.8) until the absorbance (λ=280 nm) of the wash solutions returned to baseline values. Stx2 was eluted from Synsorb-PK using three volumes of 50 mM Tris base (pH 10), 250 mM NaCl per gram of Synsorb-PK. Once eluted, Stx proteins were concentrated using an Amicon ultrafiltration (Amicon, Oakville, ON, CAN) unit with a

10 kilodalton (kDa) molecular weight cut off membrane and the final protein concentration was determined by Pierce™ BCA Protein Assay Kit (Thermo Scientific,

Waltham, MA, USA).

To generate the low endotoxin Stx2, excess lipopolysaccharide was removed using several incubations with Detoxi-Gel immobilized polymyxin B resin (Pierce). The subsequent Stx2 solution was found to contain less than 0.5 endotoxin units/mL as determined by the Limulus Amebocyte Lysate Chromogenic Endotoxin Quantitation Kit

(Thermo Scientific). Notwithstanding this purification step, the low-endotoxin Stx2 preparations contained residual LPS, as detected by immunoblotting, albeit at much lower amounts than the high endotoxin, standard preparation (see Figure 5.7). Stx preparations were diluted in PBS and stored at -80oC until ready for use. A mock Stx2 preparation was made using the same process on E. coli that did not contain the Stx2-encoding operon.

2.2.7.2 Shiga Toxin Labeling with AlexaFluor 594

Stx2 was labeled with the AlexaFluor 594 Protein Labeling Kit from Molecular

Probes, as per the manufacturer’s instructions. Briefly, 1 mg/mL Stx2 (in PBS) was brought to a pH of 7.5-8.5, using a 1:10 dilution of a 1 M sodium bicarbonate solution, and then incubated for 1 hour with the Alexa Fluor 594 dye reagent. Following incubation, the labeled Shiga toxin was purified by elution through a size exclusion

69 chromatography column, and quantified via Bradford assay. The labeled Shiga toxin was aliquoted and stored at -80°C.

2.2.9 Subcellular Fractionation

2.2.9.1 Subcellular Fractionation of Crude Membranes

THP-1 cells were collected in 1 mL of mannitol buffer (10 mM HEPES, 70 mM

Sucrose, 210 mM D-Mannitol, 0.1 mM EGTA, pH 7.4) containing protease inhibitor cocktail (Roche, Mississauga, ON, Canada). Fifteen strokes through a 27-G needle were performed to homogenize the cells, and the homogenate was centrifuged at 320 × g for

5 min to remove unbroken cells. The soluble fraction was centrifuged at 8000 × g for

15 min to collect crude membrane fraction. This was resuspended in 3X sample buffer containing 0.1% (v/v) Triton-X100 and boiled prior to immunoblotting.

2.2.9.2 Subcellular Fractionation of Mitochondria

THP-1 cells were collected in 1 mL of mannitol buffer (10 mM HEPES, 70 mM

5 min to remove unbroken cells. The soluble fraction was centrifuged at 8000 × g for

15 min to collect crude membrane fraction. The pellet, containing mitochondria, was washed in 1 mL of PBS twice and re-suspended in 0.5 M sucrose and gently layered on discontinuous sucrose gradient (30% w/w. 40% w/w, 50% w/w, 60% w/w) containing

1 mM EDTA, 0.1% (w/v) BSA, 10 mM Tris-HCl (pH 7.5). After ultracentrifugation at

51 000 × g for 3 h at 4 °C, the intact mitochondrial band was collected and sedimented by additional centrifugation at 26 000 × g for 30 min at 4 °C. Mitochondrial fractions were

70 subsequently resuspended in 3 × SDS sample buffer containing 0.1% (v/v) Triton-X100 in preparation for immunoblotting. Samples were heated to 95°C.for 5 min before loading into denaturing SDS gels for analysis. Fractions were characterized using immunoblotting and antibodies against GAPDH (cytosolic), VDAC (mitochondrial), and TLR4 (cell membrane).

2.2.10 Statistical Analysis

Data were analyzed using an unpaired Student’s t-test or ANOVA (multiple comparisons with Sidak’s or Dunnett’s tests where appropriate) where appropriate to determine statistical significance. Values are expressed as mean ± standard error of the mean (S.E.M). P values of < 0.05 were determined to be statistically significant.

GraphPad Prism software version 7.0 was used to perform all statistical analyses.

71 Chapter Three: Characterization of the Expression and Role of NLRP6 in the

Kidney Under Both Normal And Disease States

72 3.1 Rationale

Chronic kidney disease (CKD) is a major cause of morbidity and mortality in the general population, affecting over three million Canadians (130). Regardless of etiology, the pattern of disease progression is similar: the recruitment of pro-inflammatory immune cells, the propagation of tubulointerstitial inflammation, the development of renal fibrosis, and ultimately kidney failure (131-133). Given the significant expense and morbidity associated with renal replacement therapy, urgent research is needed to better understand the pathogenesis of kidney disease in order to explore novel therapeutic targets.

The NOD-like receptor (NLR) family of proteins is a group of pattern recognition receptors (PRRs) known to mediate the initial innate immune response to cellular injury and stress (2, 3). The NLRP proteins represent a fourteen-member subset of the NLR family that contains an N-terminal pyrin domain. Some NLRs are known to form multi- protein complexes known as inflammasomes. These consist of an NLR, the adaptor protein ASC, and the effector molecule pro-caspase-1. Once activated, these inflammasomes facilitate the cleavage and activation of caspase-1, which in turn mediates the cleavage of the pro-inflammatory cytokines IL-1β and IL-18 into their active and secreted forms (2, 3). To date, only 5 proteins are confirmed to assemble endogenous caspase-1 activating inflammasomes and include pyrin, NLRP3, NLRP1,

NLRC4 and the non-NLR protein AIM2.

A variety of chronic inflammatory diseases, including kidney disease, have been linked to the dysregulation of NLR proteins (2). Two NLRPs, NLRP3 and NLRP6, are highly expressed in the kidney with the former playing significant canonical and non-

73 canonical roles in chronic and acute kidney injury by propagating inflammation, cell death, and fibrosis (72, 73, 79, 86-88, 134). Prior to this thesis, the role of NLRP6 in kidney injury was unknown, as no in vivo or in vitro study of NLRP6 in in the context of the kidney had been conducted.

NLRP6 is a prototypical member of the NLRP family of proteins containing an N- terminal pyrin domain, central NACHT domain (containing a nucleotide-binding domain), and five C-terminal leucine-rich repeat domains. Though possible under overexpression conditions (80), its ability to form an inflammasome in vivo remains controversial and has not been demonstrated biochemically. Most existing studies of

NLRP6 function have been conducted in the gut, where NLRP6 was reported to have an anti-inflammatory and anti-mitogenic role in the context of DSS colitis models of inflammatory bowel disease (79, 87). NLRP6 function in this context is thought to arise through two mechanisms: 1) the regulation of the gut microbiota via control of mucous secretion and cytokine production and 2) the negative regulation of the NF-κB and

MAPK pathways in leukocytes (76, 88, 93-96, 99, 100). These ascribed functions remain under debate, with two recent studies concluding that NLRP6 played no role in the regulation of intestinal microbiota when the appropriate littermate and ex-germ-free control experiments are performed (97, 98).

Up to this point, little was known regarding the specific expression pattern and function of NLRP6 in the kidney.

74 3.2 Results

3.2.1 Nlrp6 mRNA, But Not Protein, is Expressed in Murine Glomerular Tissue

Previous studies have demonstrated that murine Nlrp6 RNA is most highly expressed in intestinal, hepatic, and renal tissue (79, 86-88). These results were replicated in the current study using quantitative real-time PCR (qRT-PCR), with the additional finding that the bulk of the renal murine Nlrp6 mRNA expression is restricted to the glomerular compartment (i.e. the functional unit of the kidney responsible for filtering the blood) (Figure 3.1A-C). Transcript levels were determined with both mechanical isolation of murine glomeruli and laser-capture microdissection, with the purity of the glomerular fractions confirmed by qRT-PCR for enrichment of Nphs2 (podocin, a marker of glomeruli) and lack of Kim-1 (a marker of damaged tubules) (Figure 3.1D). Virtually no murine Nlrp6 mRNA was detected in tubular epithelial cells (TECs), podocytes (a cell type resident to the glomerulus), renal fibroblasts, or leukocytes. In contrast to the findings at the mRNA level, the 97 kDa murine NLRP6 protein was only detectable in intestinal tissue (Figure 3.1E). Interestingly, endogenous NLRP6 protein was also not detected in liver, which also expresses high levels of Nlrp6 mRNA. These findings are consistent with previous work by Wang et al. that found no expression of NLRP6 in murine kidney or liver using a transgenic mouse expressing FLAG-tagged NLRP6, bypassing any restrictions of the commercially available murine NLRP6 antibodies (92).

Finally, since NLRP6 is reported to be upregulated in response to interferons (92), wild type mice were injected intravenously with poly(I:C) to induce a systemic type I

75 interferon response. Similar to the UUO and NTS disease models, NLRP6 protein could not be detected in the kidney or liver of these mice (Figure 3.1F).

Figure 3.1 Nlrp6 mRNA, But Not Protein, is Expressed in Murine Glomerular

Tissue.

Figure legend on next page.

77 A, B. Nlrp6 mRNA expression in a variety of murine (A.) organ systems and (B.) tissue/cell types. TECs = tubular epithelial cells. PBMCs = peripheral blood mononuclear cells. PMNs = polymorphonuclear cells. C. Nlrp6 mRNA expression in whole kidney, glomerular, and non-glomerular fractions, as isolated by laser-capture microdissection

- Ct (courtesy of Takanori Komada). Results are expressed as fold change (2 ΔΔ ) compared to spleen (A., B.) or whole kidney (C.). Bars indicate mean +/- SEM of three animal replicates. D. Nlrp6, Nphs2 (podocin), and Kim-1 mRNA expression by qRT-PCR in murine whole kidney and glomerular fractions from wild type mice. Results are

- Ct expressed as fold change (2 ΔΔ ) compared to the expression in whole kidney. E.

Immunoblotting of NLRP6 in a variety of murine tissue types derived from both wild type and Nlrp6 -/- mice on a C57Bl/6 background. F. Immunoblotting of NLRP6 in a variety of murine tissue types derived from both wild type and Nlrp6 -/- mice injected with poly(I:C). Saline injections served as a negative control. Wild type and Nlrp6- deficient ileum were used as positive and negative controls respectively.

78 3.2.2 NLRP6 mRNA, But Not Protein, is Expressed in Human Glomerular Tissue

Having demonstrated the expression pattern of Nlrp6 in murine kidney, we next sought to confirm this pattern in human tissue. As before, human NLRP6 mRNA was readily detectable in colon and kidney tissue, with the majority of the expression found in the glomerular compartment (Figure 3.2A). Human NLRP6 mRNA expression was also found to be robust in human polymorphonuclear cells (PMNs), which differed from the aforementioned murine findings. Little to no expression of human NLRP6 mRNA was found in tubular epithelial cells (TECs), glomerular endothelial cells (GECs), dermal microvascular endothelial cells (DMECs), or peripheral blood mononuclear cells

(PBMCs) (Figure 3.2A). Once again, the 98 kDa human NLRP6 protein was not detected in kidney tissue by immunoblotting, despite the readily detectable signal in fresh ileal samples (Figure 3.2B). Of note, a single freeze-thaw cycle resulted in a complete loss of

NLRP6 protein detection in the human (but not murine) ileal samples, suggesting that this protein (in human) is either very unstable or prone to precipitating out of solution (Figure

3.2B). As such, all immunoblots conducted on human tissue for NLRP6 were run using new protein extractions from freshly isolated tissue. Given the difficulties in obtaining fresh human ileal tissue for each experiment, the majority of the remaining experiments were conducted using murine tissue.

Figure 3.2 NLRP6 mRNA, But Not Protein, is Expressed in Human Glomerular

Tissue.

A. NLRP6 mRNA expression in a variety of human tissue and cell types. TECs = tubular epithelial cells. GECs = glomerular endothelial cells. DMECs = dermal microvascular endothelial cells. PBMCs = peripheral blood mononuclear cells. PMNs =

- Ct polymorphonuclear cells. Results are expressed as fold change (2 ΔΔ ) compared to tubular epithelial cells. Bars indicate mean +/- SEM of three replicates. B.

Immunoblotting of NLRP6 in a variety of human tissue types.

80 3.2.3 NLRP6 Protein Remains Undetectable Under Various Protein Extraction

Conditions

Hypothesizing that murine NLRP6 may simply be insoluble under the attempted lysis/immunoblotting protocol, protein extractions were performed under various conditions including standard RIPA lysis buffer, the more stringent urea-based lysis buffer, and boiling the tissue pellet in SDS-containing sample buffer. Immunoblotting was conducted on the prepared lysates in ileum derived from wild type and Nlrp6 -/- mice, both untreated and subject to nephrotoxic serum nephritis (Figure 3.3, Figure 3.7C).

NLRP6 was detected in all of the tested wild type ileal samples, but not the Nlrp6 -/- ileal or wild type kidney samples (Figure 3.3, Figure 3.7C). These findings suggest that murine NLRP6 protein should be soluble under the tested conditions, and that the lack of detectable expression may be due to other causes such as different tissue-specific isoforms or post-transcriptional regulation of protein expression.

Figure 3.3 NLRP6 Protein Remains Undetectable Under Various Protein Extraction

Conditions.

Immunoblotting of NLRP6 in murine kidney and ileum samples prepared via an assortment of protein extraction methods (RIPA buffer extraction, urea buffer extraction, boiling of insoluble tissue pellet with 3X SDS sample buffer) from both wild type and

Nlrp6 -/- mice on a C57Bl/6 background.

82 3.2.4 The Commercially Available Antibodies Against Human NLRP6 Bind Different

Domains of NLRP6

To test the possibility that different isoforms of NLRP6 may exist, thereby hindering antibody binding, epitope mapping of the commercially available human anti-

NLRP6 antibodies was conducted. Constructs were generated encoding the full-length human NLRP6 protein, as well as the individual domains and combinations of domains.

These were then expressed in HEK293T cells and blotted using anti-human NLRP6 antibodies from Adipogen (Clint-1) and R&D Systems. The former was found to bind to an epitope internal to the NACHT domain of NLRP6, whereas the latter was found to bind to an epitope covering both part of the pyrin domain and part of the downstream interdomain (Figure 3.4A-C). Since the antibodies bind different conserved domains spanning the N-terminal and central regions of the protein, it appears unlikely that the presence of a novel isoform explains the inability to detect the protein by immunoblotting and that the protein is simply not expressed in the kidney. Considering the high levels of mRNA produced and the lack of NLRP6 protein in the kidney, the NLRP6 gene may be subject to post-transcriptional regulation that inhibits its translation.

Figure 3.4 The Commercially Available Antibodies Against Human NLRP6 Bind

Different Domains of NLRP6.

Figure legend on next page.

84 A, B. Immunoblotting, using (A.) the Adipogen (Clint-1) anti-human NLRP6 antibody or

(B.) the R&D anti-human NLRP6 antibody, of GFP-tagged NLRP6 gene constructs transiently transfected into HEK293T cells encoding full-length NLRP6, single NLRP6 domains, or NLRP6 domain combinations generated through both cloning and site- directed mutagenesis of the human NLRP6 gene. Immunoblotting for the common GFP tag was performed as a control for protein expression. C. Model demonstrating the epitopes bound by each antibody.

85 3.2.5 Nlrp6 +/+ and Nlrp6 -/- Mice Have Indistinguishable Phenotypes in Murine

Models of Kidney Disease

Despite the lack of detectable NLRP6 protein by immunoblotting in kidney at baseline, experiments were performed in vivo to investigate the possibility that NLRP6 protein translation may be triggered during kidney injury and inflammation. The rationale for this being that if translation of the NLRP6 protein is induced following the introduction of renal injury/inflammation, it may play a role in the pathogenesis of inflammatory kidney disease.

First, unilateral ureteral obstruction (UUO) was performed to induce chronic kidney inflammation and fibrosis. In the UUO model, the left ureter of the mouse is surgically ligated in order to induce ipsilateral hydronephrosis. Over a 7 or 14-day time course, the backlog of urine results in mechanical and chemical stresses that induce both inflammatory and fibrotic responses in the affected kidney, characterized by leukocyte infiltration (e.g. macrophages, neutrophils) and the deposition of matrix proteins (e.g. type I collagen). The contralateral kidney is left unligated both to prevent the systemic manifestations of kidney failure (one kidney is sufficient to maintain an otherwise normal phenotype) and to serve as an internal negative control (134).

In a manner consistent with earlier studies in other organ systems, murine Nlrp6 mRNA levels were found to decrease markedly in murine kidneys subject to UUO for seven or fourteen days, likely reflecting a loss of Nlrp6-expressing glomerular cells observed in this model (Figure 3.5A) (87, 89, 90). This is in contrast to murine Nlrp3 mRNA levels, which increased as the disease model progressed, in agreement with

86 previous publications (Figure 3.5B) (134). Furthermore, no NLRP6 protein was expressed in the kidney following induction of the disease state (Figure 3.5C).

Histologically, no significant differences were observed between wild type and

Nlrp6-deficient mice following seven and fourteen day UUO time points in terms of inflammatory cell infiltration or fibrosis as determined by H&E staining, Masson’s

Trichrome staining, and Picrosirius red staining (Figure 3.6A). These findings were supported biochemically, where no significant differences in CD11b, α-SMA protein and

Col1a1 mRNA expression were noted by immunoblotting and qRT-PCR respectively

(Figure 3.6B-D).

Figure 3.5 NLRP6 Expression in the Unilateral Ureteral Obstruction (UUO) Model of Murine Kidney Disease.

Figure legend on next page.

88 A, B. (A.) Nlrp6 and (B.) Nlrp3 mRNA expression by qRT-PCR in murine kidney samples from both wild type and Nlrp6 -/- mice following unilateral ureteral obstruction

(UUO) for 7 and 14 days. Sham-ligation procedures and contralateral kidneys served as

- Ct negative controls. Results are expressed as fold change (2 ΔΔ ) compared to wild type mice that underwent sham surgery. Bars indicate mean +/- SEM of at least three animal replicates. Double asterisks indicate statistical significance (P < 0.01). C. Immunoblotting of NLRP6 in murine kidney from both wild type and Nlrp6 -/- mice on a C57Bl/6 background, untreated or subject to 7 or 14-day time courses of UUO. Wild type and

Nlrp6-deficient ileum were used as positive and negative controls respectively.

Figure 3.6 Nlrp6 +/+ and Nlrp6 -/- Mice Have Undistinguishable Histological and

Biochemical Phenotypes in the Unilateral Ureteral Obstruction (UUO) Model of

Murine Kidney Disease.

Figure legend on next page.

90 A. Representative H&E (hematoxylin and eosin), Masson’s Trichrome, and Picrosirius red staining of murine kidney sections from both wild type and Nlrp6 -/- mice following

14 days of UUO, observed under 20X magnification. Polarized light microscopy was employed to visualize the Picrosirius red staining. Histology of contralateral kidneys served as a negative control. B, C. Densitometry of western blots for (B.) CD11b and

(C.) αSMA (normalized to β-tubulin) in murine kidney samples from both wild type and

Nlrp6 -/- mice, following unilateral ureteral obstruction (UUO) for 7 and 14 days. Sham- ligated and contralateral kidneys served as negative controls. D. Type I collagen (Col1a1) expression by qRT-PCR in murine kidney samples from both wild type and Nlrp6 -/- mice following unilateral ureteral obstruction (UUO) for 7 and 14 days. Contralateral

- Ct kidneys served as negative controls. Results are expressed as fold change (2 ΔΔ ) compared to the expression in contralateral kidneys. Bars indicate mean +/- SEM of at least three replicates.

91 Given that Nlrp6 mRNA is restricted to the glomerular compartment of the kidney, we next sought to examine the impact of this gene on a murine model of glomerulonephritis induced by nephrotoxic serum targeting the glomerular basement membrane (135). Disease pathogenesis in this model consists of an acute inflammatory reaction at the glomeruli, resulting in the progressive development of proteinuria, hematuria, and ascites (122). Specifically, as a classic example of type II hypersensitivity, anti-glomerular basement membrane antibodies (anti-GBM antibodies) bind to the basement membrane of the glomeruli. Antibody binding results in the activation of complement (through the canonical complement cascade) resulting in both the generation of chemotactic anaphylotoxins (i.e. C3a, C5a) and the complement-mediated destruction of nearby tissue. Additionally, antibody binding and anaphytoxin generation results in the recruitment, activation, and degranulation of circulating leukocytes, further driving glomerular tissue damage and the death of bystander cells (in the glomerulus, largely podocytes and endothelial cells). The loss of endothelial- and podocyte-mediated barrier function in this disease model results in hematuria and proteinuria, the inability to maintain osmotic pressure in the vasculature, and ultimately the development of ascites.

This disease model also results in the formation of “crescents” in the glomeruli, as observed by PAS staining. Crescents are indicative of cellular proliferation in Bowman’s space, arising in response to the induced inflammation as part of a repair response (136,

137).

Wild type and Nlrp6 -/- mice were administered sheep serum containing anti- glomerular basement membrane antibodies over two days, with weight measurements and

92 urine collected at days 0, 4, 7, and 10 (Figure 3.7A). On day 10, the mice were sacrificed and the kidneys harvested for analysis.

Similar to the UUO findings, Nlrp6 mRNA expression decreased following the

10-day course of the NTS model and no NLRP6 protein was expressed as a result of the induced glomerulonephritis (Figure 3.7B, C). Also consistent with the UUO data, no phenotypic differences were observed between wild type and Nlrp6-deficient mice following induction of experimental glomerulonephritis. Specifically, there were no significant differences in terms of proteinuria, albuminuria, or glomerular crescent formation (Figure 3.8A-D). Although there appeared to be a significant improvement in the proteinuria of Nlrp6-deficient mice on day 7 of the model, this is finding was primarily due to sample bias arising from differences in urine output between Nlrp6 -/- and wild type controls (Figure 3.8E). Over the entire course of this model however, there were no observable differences between the genotypes in terms of proteinuria.

Finally, since NLRP6 is known to be upregulated in response to interferons (92), wild type mice were injected intravenously with poly I-C to induce a systemic type I interferon response. Similar to the UUO and NTS disease models, NLRP6 protein could not be detected in the kidney or liver of these mice (Figure 3.1F).

Altogether, these results seem to indicate that NLRP6 protein is not expressed in kidney tissue and it does not play a role in the pathogenesis of the studied models of inflammatory kidney disease.

Figure 3.7 NLRP6 Expression in the Nephrotoxic Serum Nephritis (NTS) Model of

Murine Kidney Disease.

Figure legend on next page.

94 A. Model of nephrotoxic serum nephritis treatment course (tail vein injection and urine collection performed by Bjorn Petri and the Mouse Phenomics Core at the University of

Calgary). B. Nlrp6 mRNA expression by qRT-PCR in murine kidney samples from both wild type and Nlrp6 -/- mice following 10 days of NTS. Results are expressed as fold

- Ct change (2 ΔΔ ) compared to the expression in untreated kidneys. Bars indicate mean +/-

SEM of at least three animal replicates. Double asterisks indicate statistical significance

(P < 0.01). C. Immunoblotting of NLRP6 in murine kidney prepared via an assortment of protein extraction methods (RIPA buffer extraction, urea buffer extraction, boiling of insoluble tissue pellet with 3X SDS sample buffer) from both wild type and Nlrp6 -/- mice on a C57Bl/6 background. The kidney samples were taken from both untreated mice

(-) or mice subjected to a ten-day course of nephrotoxic serum nephritis (NTS). Murine ileum samples served as a positive control for NLRP6 protein expression.

Figure 3.8 Nlrp6 +/+ and Nlrp6 -/- Mice Have Undistinguishable Histological and

Biochemical Phenotypes in the Nephrotoxic Serum Nephritis (NTS) Model of

Murine Kidney Disease.

Figure legend on next page.

96 A. Representative Periodic Acid-Schiff (PAS) staining of murine kidney sections from both wild type and Nlrp6 -/- mice following 10 days of NTS, observed under 40X magnification. B, C. Quantification of (B.) urine albumin by ELISA and (C.) urine protein by Bradford assay at zero, four, seven, and ten days of NTS treatment in both wild type and Nlrp6 -/- mice. D. Quantification of crescentic glomeruli in kidney sections from both wild type and Nlrp6 -/- mice following 10 days of NTS, represented as a percentage of total glomeruli counted in each genotype. E. Quantification of wild type and Nlrp6 -/- mice demonstrating anuria following 7 days of NTS, represented as a percentage of total mice in each genotype. Bars indicate mean +/- SEM of at least three animal replicates. Asterisk indicates statistical significance (P < 0.05).

97 3.2.6 RNASeq Identifies a Novel Fusion Transcript and Reveals Select Differences in

Gene Expression Between Nlrp6 -/- and Wild Type Mice at Baseline

The lack of protein expression observed in the mouse and human kidney suggested that the Nlrp6 gene might be regulated though some form of post- transcriptional modification, which would be informative in order to determine the role of this gene in the kidney. To understand this mechanism of regulation and to examine at a global level whether or not Nlrp6 was a relevant gene in the kidney, we performed

RNASeq analysis in the kidney and gastrointestinal tract (colon) of Nlrp6 -/- and wild type mice at baseline.

Interestingly, RNAseq revealed that there was a kidney-specific fusion of the long, non-coding RNA (lncRNA) BC024386 to the 5’ end of the Nlrp6 gene (Figure

3.9C). BC024386, which encodes a lncRNA thousands of base pairs upstream of Nlrp6, is known to possess three exons (Figure 3.9A). The sequencing performed for these experiments revealed that, in a kidney-specific manner, the first exon of the BC024386 locus is alternatively spliced to two novel downstream exons, spanning between

140917371-140917464 and 140919679-140919816 respectively (Figure 3.9B). These three exons from BC024386 are then spliced directly to the 5’ end of the Nlrp6 mRNA

(Figure 3.9C). This fusion transcript does not appear to contain a Kozak consensus sequence and as such it is highly unlikely that NLRP6 translation is possible. This phenomenon was not observed in colon tissue, revealing a potential mechanism for the lack of NLRP6 protein expression in murine kidney: gene silencing through the generation of a novel fusion, lncRNA transcript (Figure 3.9C). Additionally, an NCBI

Gene search revealed that the tissues expressing the highest levels of BC024386 (based

98 on transcriptomics data) are kidney and liver (138). These findings are consistent with work by Halpern et al. that concluded that Nlrp6 mRNA is sequestered in the nucleus of select tissue types, specifically liver (139). Seeing as many lncRNAs demonstrate this compartmental restriction, it is possible that the described fusion event occurs in non- renal tissue as well and is the mechanism underlying the observed sequestration event in liver tissue.

Analysis of gene expression between wild type and Nlrp6-deficient mice revealed

42 genes that are differentially expressed in kidney and colon, 4 that are differentially expressed only in the colon, and 7 that are differentially expressed only in the kidney

(Figure 3.10A, B). The 42 genes common to both kidney and colon may be due to genetic drift in our mouse lineage, which seems likely given the entirely different mechanisms of gene regulation at play (i.e. protein expression versus fusion transcript generation).

Though more likely, pathway analysis of the differentially expressed genes found that 13 of these genes (including Mecp2, a gene differentially expressed only in gut) are part of causal network downstream of Apoc2, suggesting that these findings may be significant

(Figure 3.10D). Additional work will need to be done to validate these findings and explore their potential implication with regard to Nlrp6.

The finding that only 4 transcripts are differentially expressed in the colon also supports recent data that demonstrate no microbiota changes between Nlrp6 -/- and wild type littermates (Figure 3.10A) (97, 98).

The fact that 7 genes are differentially expressed in the kidney notwithstanding the existence of a lncRNA fusion transcript suggests that the fusion transcript may yet have a functional role in the kidney at a post-transcriptional level (Figure 3.10B).

99 Currently, lncRNAs are thought to regulate gene expression via both the regulation of microRNAs and the direct interaction with transcriptional and epigenetic machinery both in cis and in trans (140). The mechanism through which this lncRNA-Nlrp6 mRNA fusion transcript mediates gene expression remains to be studied.

Of the seven genes differentially expressed in the kidney, one in particular is worth noting: the retinoblastoma protein gene Rb1. Mice deficient in Nlrp6 expressed 210 fold more Rb1 mRNA than wild type controls and preliminary immunoblotting suggests that RB1 is indeed upregulated in Nlrp6-deficient mice, suggesting a possible role for this gene in renal tumorigenesis (Figure 3.10B, C). Specifically, given its role as a tumor suppressor, the upregulation of RB1 in Nlrp6-deficient mice may render these animals resistant to renal cell carcinoma. Further experimentation is required to confirm the preliminary immunoblotting and to explore the effect on downstream signaling through both in vitro and in vivo models.

100

Figure 3.9 Unique Fusion Splicing of BC024386 to Nlrp6 As Determined by RNAseq

Analysis.

Figure legend on next page.

101 A. The gene organization of the mRNA-encoding murine Nlrp6 gene and the lncRNA locus BC024386, as denoted by their exonic organization and nucleotide position according to NCBI Gene. Solid boxes (black or white) denote exons, with shorter boxes denoting known untranslated regions (UTRs). B. The unique splice variant of BC024386 discovered in murine kidney tissue, where the canonical exon 1 is spliced to two novel exons downstream. C. A diagram illustrating the kidney-specific fusion of three exons

(one known, two novel) of the lncRNA BC024386 locus to the 5’ end of the Nlrp6 mRNA, as determined by RNAseq. The RNAseq analysis was performed by Paul Gordon and the DNA Sequencing Lab at the University of Calgary. Experiments were performed in triplicate, with each replicate composed of three biological replicates.

102 A.

Target ID P Value Q Value b Gene Symbol NM_001081979.2 3.48E-27 8.46E-23 -6.585392771 MECP2 NM_133946.2 3.97E-19 6.44E-15 -8.60407242 NLRP6 XM_006508512.3 7.01E-19 8.52E-15 -5.548409274 SLC22A18 NM_001013753.2 3.60E-10 3.50E-06 -5.879538573 PCDH17 NM_146155.3 7.07E-10 5.28E-06 -5.001930985 AHDC1 NR_037992.1 7.60E-10 5.28E-06 -5.606593897 JMJD4 NM_001271727.1 1.44E-09 8.73E-06 -6.594470698 TRIM2 NM_001305671.1 1.66E-09 8.99E-06 -6.199365684 GLIS3 NM_001146084.1 2.81E-08 0.00013671 -5.457255708 FASTKD5 NM_001159964.1 3.86E-08 0.000170685 -5.145082016 EPS15 XM_017315983.1 8.73E-08 0.000354146 -6.47061329 PCDH17 NM_001037298.1 9.63E-08 0.000360595 -5.738581879 PIEZO1 XM_011243460.2 1.44E-07 0.000501265 -5.891499207 SYN3 XM_006532399.2 2.10E-07 0.000682253 -5.685721668 MGAT1 XM_006526086.3 3.46E-07 0.001052835 -5.093539454 LDLRAD4 XM_011240821.2 1.01E-06 0.002903167 -4.885214064 NCOR2 NM_001080774.1 1.23E-06 0.003318109 -5.19505094 MYO1C XM_006519338.3 1.53E-06 0.003909671 -5.022107393 SLC22A17 NM_001037955.4 2.14E-06 0.005206733 -4.255643332 DUSP22 XM_006497911.3 2.76E-06 0.006400582 -4.836717641 SEC16A XM_006495441.3 3.92E-06 0.008680082 -5.521505231 PREX2 NM_015767.4 5.02E-06 0.009769039 -4.995079408 TTPA XM_006510334.3 4.65E-06 0.009769039 -5.245287459 ZC3H12C XM_006530114.3 4.82E-06 0.009769039 -4.96554645 ACACB NM_001013786.2 6.41E-06 0.011550165 -5.192801489 ZSCAN26 NM_025914.2 6.18E-06 0.011550165 -4.786931618 ACTR6 XM_006500973.2 9.73E-06 0.016906424 -5.632311246 CRYZ NM_027871.2 1.20E-05 0.019562189 -4.833266285 ARHGEF3 XM_017320098.1 1.21E-05 0.019562189 -5.08173146 ARHGEF19 XM_006517337.3 1.59E-05 0.0227772 -4.810636725 AGTPBP1 XM_006521703.3 1.46E-05 0.0227772 -5.511436757 MYLK XM_017315041.1 1.54E-05 0.0227772 -5.614949315 ELK4 XR_383160.3 1.56E-05 0.0227772 -5.814766678 CHDH NM_177378.4 1.67E-05 0.023047254 -4.734386215 RNF150 XM_006501721.2 1.70E-05 0.023047254 -4.950215324 GBP4 XM_006516563.3 1.91E-05 0.025105607 -5.530032872 LYST XM_006503000.2 2.08E-05 0.026636369 -5.813034501 SLC5A9 NM_028181.5 2.37E-05 0.029536773 -4.854919139 CCPG1 XM_017313235.1 2.44E-05 0.029673113 -4.796668976 SMARCA4 XM_006506004.3 2.80E-05 0.033250384 -5.677631589 FRMD4B XM_011248440.2 3.19E-05 0.036048262 -6.375686142 ATXN1L XR_001783713.1 3.15E-05 0.036048262 -5.559625064 FUBP1

103 XM_006526177.3 3.55E-05 0.038443181 -6.022331184 CXXC5 XM_006528763.3 3.55E-05 0.038443181 -4.797895183 REPS2 XM_006529814.2 4.03E-05 0.042677985 -5.512742895 ILKAP NR_003364.1 0.0037969 0.9776717 2.537491 BC018473

Target ID P Value Q Value b Gene Symbol XM_006523654.3 1.73E-07 0.001684242 10.91285017 FOXN2 XR_383150.3 2.33E-06 0.016196822 10.52618648 RB1 NM_001201414.1 3.57E-06 0.019318876 11.11541016 APBB2 XM_006523391.3 3.21E-06 0.019318876 5.490205401 FAM234A NM_146155.3 6.54E-06 0.028912722 5.783916972 ZNF598 XR_001783772.1 7.45E-06 0.030219969 6.309848444 ATP11B NM_172937.3 1.28E-05 0.047849757 5.740361554 SHPRH NM_027922.2 9.69E-07 0.007862009 -5.94118799 ANKLE2

104

Figure 3.10 Differential Expression of Genes Between Nlrp6 +/+ and Nlrp6 -/- Mice in Colon and Kidney Tissue as Determined by RNASeq.

Figure legend on next page.

105 A. Genes differentially expressed between wild type and Nlrp6-deficient mice in both kidney and colon, with the exception of the bolded genes that were only differentially expressed in colon. B. Genes differentially expressed between wild type and Nlrp6- deficient mice only in the kidney. The DNA Sequencing Lab at the University of Calgary performed the RNAseq analysis. Experiments were performed in triplicate, with each replicate composed of three biological replicates. All differentially expressed genes that are listed passed the Wald test with regard to statistical significance. C. Immunoblotting of RB1 in murine kidney and liver from both wild type and Nlrp6 -/- mice on a C57Bl/6 background. Actin was used as a loading control. D. Pathway analysis (courtesy of Paul

Gordon of the DNA Sequencing Lab of the University of Calgary) of 13 of the genes differentially expressed between wild type and Nlrp6 -/- mice on a C57Bl/6 background, all of which are downstream of Apoc2. Genes highlighted in green are significantly differentially expressed, with the b values listed below each gene. The network has a network bias corrected p-value of 0.02 and a directionality Z-score of -3.2.

106 3.3 Summary

The results in this chapter (Chapter 3) demonstrate that NLRP6 protein is not expressed in the kidney under either normal homeostatic conditions or pathological states, in spite of the high levels of detected mRNA. The inability to detect NLRP6 protein via immunoblotting cannot be ascribed to limitations of the available antibodies or the existence of non-detectable isoforms. It appears that Nlrp6 is subject to a mechanism of post-transcriptional regulation involving the generation of a fusion lncRNA-Nlrp6 transcript and perhaps nuclear mRNA sequestration that remains incompletely understood (139). Its role in regulating RB1 expression, likely at a post- transcriptional level, requires further study given the relevance of said protein in tumorigenesis and renal cell carcinoma. Further research is required to explore the regulation of Nlrp6 gene expression in order to elucidate the mechanisms underlying this function and subsequently its biological properties in the kidney and other organs.

To do this, the following experiments will be undertaken. First, the sequence of the fusion lncRNA-Nlrp6 mRNA transcript will be cloned and submitted for Sanger sequencing. Additional work will also examine whether this phenomenon is occurring in liver tissue via PCR. Second, the observed kidney-specific differential expression of genes will need to be validated at the protein level, with specific interest in the expression of RB1: a known tumor suppressor important in the development of renal cell carcinoma.

If RB1 is indeed upregulated in Nlrp6-deficient animals (as appears to be the case in preliminary immunoblotting), an in vivo model of renal cell carcinoma will be investigated to ascertain the role of this protein in the pathogenesis of renal cancer after which the mechanism of fusion transcript-mediated regulation of RB1 expression will

107 need to be elucidated. These findings will then have to be examined in human tissue to see if this form of gene regulation is applicable across species.

108 Chapter 4: Characterization of the Fundamental Molecular Biology Underlying

NOD-like Receptor Activation

109 4.1 Rationale

At the outset of this thesis, it was understood that the NOD-like receptors (NLRs) are a family of intracellular pattern recognition receptors (PRRs) known to respond to a wide range of pathogen- and danger-associated molecular patterns (PAMPs and DAMPs respectively) (2). Upon recognition of these stimuli, NLRs are known to oligomerize and modulate a number of downstream functions, including the maturation of pro- inflammatory cytokines via inflammasome formation and the regulation of immune signaling pathways (i.e. NF-κB and MAPK pathways) (2, 75-77). As can be seen in the previous chapter, these facts are not necessarily applicable to every member of the NLR family and, in fact, the functions of these genes may be diverse and varied depending on the conditions and tissue types in which they are expressed.

When expressed as proteins, the NLRs share highly conserved domain architecture: they all possess C-terminal leucine-rich repeat (LRR) domains and a central, nucleotide-binding NACHT domain. They vary structurally at their N-termini, with most

NLRs falling into two broad categories: 1) NLRs containing pyrin domains (PYDs) and

2) NLRs containing caspase-recruitment domains (CARDs). These differences at the N- terminus allow for the formation of different effector complexes, depending on the NLR family member nucleating the reaction (2, 3).

A number of the PYD-containing NLRs (or NLRPs) (i.e. NLRP1, NLRP3) are known to form multi-protein complexes known as inflammasomes, consisting of an

NLRP, the bipartite adaptor protein ASC, and the effector protein caspase-1. When activated, NLRPs nucleate the oligomerization of ASC and the autocatalytic cleavage of

110 caspase-1, whose active form cleaves the cytokines pro-IL-1β and pro-IL-18 into their active and secreted forms (2, 3). Active caspase-1 also cleaves gasdermin D: a protein whose pore-forming, N-terminal domain is responsible for executing the programmed, inflammatory form of cell death known as pyroptosis (105, 106).

Apart from their well-defined roles in inflammasome formation, certain NLRPs are reported to regulate immune signaling cascades, such as the NF-κB and MAPK pathways. These include NOD2, NLRP6, and NLRP12 (70, 76, 77). The precise mechanisms underpinning this regulation are unknown; however, it is known that NLR oligomerization is a critical step in this process. Other functions of NLRs, such as the regulation of gene expression by NLRP6 described in Chapter 3, have only recently begun to be characterized.

In addition to the aforementioned conserved domain architecture, NLRs also share considerable homology with both plant disease resistance proteins (R proteins) and the apoptotic mediator APAF-1 (9, 141-143). These proteins, members of the AAA+ family of ATPases, all possess a central nucleotide-binding domain (NBD) that is thought to be critical in regulating protein function (8, 144). In fact, several auto-inflammatory diseases

(e.g. Cryopyrin-Associated Periodic Syndromes (CAPS), Blau syndrome, early onset sarcoidosis, etc.) are known to arise due to gain of function mutations in the NBD of select NLR proteins (9, 82, 84, 85, 145).

Despite the considerable body of research conducted on NLR proteins, the precise sequence of events governing their activation, in terms of ATP-binding and hydrolysis, remains poorly defined. The current assumption in the literature is that ATP hydrolysis is required for NLR function, which has not been demonstrated; though ATP-binding is

111 known to be essential for downstream function, the roles of ATP-binding and hydrolysis in this regard have not been parsed.

Given the structural homology of NLR proteins and the known link between NBD mutations and auto-inflammatory disease, we hypothesized that NLRs share a common mechanism of activation that is dependent on the cycling of ATP. In this chapter, we demonstrate that NLRs require ATP-binding to oligomerize and regulate downstream pathways, in a manner analogous to small GTPases and plant R proteins. This is true of both canonical (i.e. caspase-1-activating, inflammasome-forming NLRs) and non- inflammasome-forming NLRs (with NLRP3 and NLRP6 serving as prototypical examples respectively). The rationale for comparing these particular NLRs was three- fold: 1) it enabled the comparison of an inflammasome-forming NLR with a non- inflammasome-forming NLR in order to examine commonalities and differences in activation, 2) it enabled the extrapolation of findings in NLRP6 (an NLR with no widely accepted functional readout) to an NLR with well-defined effector functions and 3) it was hoped that in understanding its fundamental biology, screening could be performed to investigate the function of the NLRP6 protein, given the controversial and contradictory findings present in the literature to date. Given the findings presented in Chapter 3, these findings would be very applicable in gut tissue, where the NLRP6 protein is highly expressed and readily detectable.

Furthermore, we show that mutations that abrogate ATP hydrolysis lock the

NLRs in an active, ATP-bound state that leads to hyperactivity and auto-inflammatory disease. These findings support a model wherein ATP-binding licenses NLR activation in

112 response to PAMPs and DAMPs, and hydrolysis maintains the NLR protein in an ADP- bound, inactive state.

113 4.2 Results

4.2.1 ATP-Binding in NLRs is Mediated by Residues in the Walker A, Walker B, and

Lid Regions of the ATP-Binding Pocket.

In silico analysis has revealed significant sequence similarity among NLR family members, particularly at the site of the ATP-binding pocket (Figure 1.3) (82). Apart from the well-characterized Walker A and Walker B motifs, in silico analysis has identified a region approximately 200 residues downstream of the Walker A motif that is highly similar across most NLR family members (Figure 4.1A). This area of conservation consists of 12 amino acids with the consensus sequence ChhPhhCWhhCS/T, where h represents a hydrophobic amino acid (Figure 4.1A). In addition to the conserved cysteine and hydrophobic residues, this region contains a central proline residue positionally similar to the GxP motif found in APAF-1 (Figure 4.2) (143). Conserved across most

NLRs, mutagenesis of this proline residue is known to abrogate downstream signaling in

NOD2 (145). Given its known interaction with the adenine moiety of ATP in the case of

APAF-1 and the general hydrophobicity of this region, it was postulated that this region might serve as the lid to the ATP-binding pocket to facilitate the binding of ATP.

To investigate this possibility, this region was examined by in silico modeling using Phyre2 in two model NLRs: NLRP6, a non-inflammasome-forming NLRP of poorly characterized function; and NLRP3, a canonical, inflammasome-forming NLRP

(Figure 4.1B).

Phyre2 modeling, based on the crystal structure of APAF-1, revealed that this potential lid region directly overlays the ATP-binding Walker A site in both NLRP6 and

114 NLRP3 (Figure 4.1B). The hydrophobic residues that insert towards the Walker A site of

NLRP3 and NLRP6 appear to be coordinated by the conserved proline residue, whose conformational rigidity results in the formation of a turn at this site (Figure 4.1B). Given this positioning, it is highly likely that this conserved proline residue (henceforth referred to as the GxP motif) directly interacts with the adenine moiety in a manner consistent with the GxP motif of APAF-1 (Figure 4.2) (143).

115

116

117

Figure 4.1 Structural Similarity and Phyre2 Modeling of NLRP3 and NLRP6.

Figure legend on next page.

118 A. Schematic diagrams depicting the relative locations of the Walker A sites and conserved hydrophobic regions in NLRP3 and NLRP6, alongside sequence alignments of the latter region across the entire NLR family of proteins. B. Phyre2 modeling of NLRP6 and NLRP3, using APAF-1 as a related protein with a defined crystal structure, zooming in on the Walker A sites and conserved hydrophobic regions.

119

Figure 4.2 Model of the ATP-Binding Pocket of NLRP3.

A schematic diagram depicting the ATP-binding pocket of NLRP3. The arginine finger motif (R262) is highlighted in purple, the residues of the Walker B motif (D305, E306) are highlighted in orange, and the relevant serine/threonine residue of the Walker A motif

(T233) is highlighted in cyan. The residues of the putative lid region are represented by the yellow, with the key proline residue highlighted (P412).

120 4.2.2 Mutation of Either the Walker A Motif or GxP Motif-Containing Lid Region

Attenuates NLR Oligomerization.

NLR proteins are mechanistically linked by their need to oligomerize in order to transduce signals. Nucleotide-binding is known to be essential for NLR oligomerization, the abrogation of which abolishes downstream function, even in hyperactive mutants (5).

The residues of the Walker A site are known to be involved in ATP-binding via electrostatic interactions with the β and γ phosphates of the bound ATP molecule. Given the modeled position of the NLRP GxP motif and its known role in APAF-1 and NOD2, it was hypothesized that mutation of this residue or the region as a whole would impair

NLR oligomerization by impairing ATP-binding.

To test this hypothesis, an in vitro oligomerization assay was developed under over-expression conditions in HEK293T cells. When FLAG and GFP-tagged wild type

NLR proteins are co-expressed under these conditions, they spontaneously oligomerize in a complex that can be pulled down via co-immunoprecipitation against the FLAG tag

(Figure 4.3A, C). To examine the effects of mutagenesis on the lid region (specifically the GxP motif and several other conserved amino acids), wild type FLAG-NLRP6 (or

NLRP3) was co-transfected with mutant NLRP6 (or NLRP3)-GFP and co- immunoprecipitation against the FLAG tag was performed as before. Following immunoblotting, densitometry was conducted to determine the ratio of mutant NLRP-

GFP-to-wild type FLAG-NLR pulled down, with the final result expressed as a percentage of this ratio compared to that calculated for the wild type control pull-down.

Significant defects in oligomerization were observed in the lid region deletion

(Δ403-430 in NLRP3, Δ379-406 in NLRP6) and proline mutants (P412A in NLRP3,

121 P388A in NLRP6) of both NLRP3 and NLRP6, with comparable phenotypes to the

Walker A mutants (WA: 226GAAGIGKT233 à 226GAAGIAAA233 in NLRP3,

202GPAGIGKT209 à 202GPAGIAAA209 in NLRP6) (Figure 4.3A-D). This would suggest that the putative lid region acts in concert with the Walker A site to promote ATP binding and efficient oligomerization of NLR proteins.

122

Figure 4.3 Mutation of the Putative Lid Region Impairs NLR Protein

Oligomerization.

Figure legend on next page.

123 A. Immunoblotting of input lysates or co-immunoprecipitation for FLAG in

HEK293T cells co-transfected with wild type FLAG-NLRP3 and either wild type or mutant NLRP3-GFP. B. Quantification of FLAG-NLRP3-to-NLRP3-GFP ratio by densitometry, represented as a percentage of the control wild type co- immunoprecipitation ratio. C. Immunoblotting of input lysates or co- immunoprecipitation for FLAG in HEK293T cells co-transfected with wild type FLAG-

NLRP6 and either wild type or mutant NLRP6-GFP. D. Quantification of FLAG-

NLRP6-to-NLRP6-GFP ratio by densitometry, represented as a percentage of the control wild type co-immunoprecipitation ratio. WT = wild type, WA = Walker A mutants

(226GAAGIGKT233 à 226GAAGIAAA233 in NLRP3, 202GPAGIGKT209 à

202GPAGIAAA209 in NLRP6), Δ403-430/Δ379-406 = lid region deletion mutants in

NLRP3 and NLRP6 respectively. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001,

****: p<0.0001).

124 4.2.3 Mutation of Either the Walker A Motif or GxP Motif-Containing Lid Region

Attenuates NLRP3 Inflammasome Activation.

Oligomerization subsequent to ATP-binding is required for downstream NLR signaling. Seeing as NLRP6 has no universally accepted functional readout, the effects of mutagenesis on NLRP3 function were examined. Specifically, the effects of mutagenesis on IL-1β maturation and secretion downstream of the NLRP3 inflammasome were investigated.

To examine if mutation of the GxP motif-containing lid region inhibits downstream NLR signaling as well as oligomerization, the NLRP3 inflammasome was reconstituted in HEK293T cells using both wild type and mutant constructs of NLRP3.

Activation was examined by the cleavage of caspase-1 and the cleavage and secretion of

IL-1β into the cell supernatant. Loss of either the entire lid region (Δ403-430) or the GxP motif alone (P412A) significantly reduced NLRP3 inflammasome activation, as denoted by reduced caspase-1 cleavage and reduced IL-1β secretion, to a level comparable to both the NLRP3-deficient and Walker A mutant (WA) controls (Figure 4.4). Interestingly, the

C409A mutant also had reduced NLRP3 inflammasome activation, despite no apparent oligomerization defect (Figures 4.3, 4.4). Given the sensitivity of thiol-containing residues to the redox state of the cell, it’s possible that the conserved cysteine residues play a role in inflammasome signaling when appropriate redox conditions are met.

These findings are consistent with previous studies of NOD2, which found that loss of the Walker A or GxP motifs result in decreased activation of the NF-κB signaling pathway compared to wild type controls (145). As such, it appears that both

125 inflammasome-dependent and independent functions of NLRs are dependent on ATP- binding mediated by the Walker A and GxP motifs of these proteins.

126

Figure 4.4 NLRP3 is Functionally Impaired Following Mutation of the Putative Lid

Region.

HEK293T cells, grown in a 24-well plate, were transfected with ASC (5 ng/well), pro- caspase-1 (5 ng/well), pro-IL-1β (300 ng/mL), and either wild type or mutant NLRP3 (15 ng/well). 6 hours post-transfection, cells were lysed, supernatants were collected for IL-

1β ELISA, and immunoblotting was performed to examine efficiency of inflammasome activation (experimental data courtesy of Kuo Chieh Liao). WT = wild type, WA =

Walker A mutant (226GAAGIGKT233 à 226GAAGIAAA233), and Δ403-430 = lid region deletion mutant. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001, ****: p<0.0001).

127 4.2.4 Abrogation of ATPase Activity in NLRs Results in Hyperactivity.

NLRs also have intrinsic ATPase activity, in keeping with their membership with the AAA+ family of ATPases. NLRP3, NLRP7, NLRP12, and NOD2 have confirmed hydrolytic activity (5-7, 9). The role of ATP hydrolysis in NLR function remains controversial, with studies linking ATP hydrolysis to both NLR activation and inactivation (61, 85, 145). No study has rigorously parsed the distinct roles of these processes in NLR activation.

Mutagenesis studies and modeling of NOD2 have found that known hyperactive

SNPs cluster around residues thought to be responsible for ATP hydrolysis, based on the conserved mechanism of ATP hydrolysis in AAA+ ATPases (85, 145). An example of this proposed mechanism of hydrolysis for NLRP3 is provided (Figure 4.5). Specifically, mutation of NOD2 residues theorized to be responsible for the activation of the water molecule and promoting the nucleophilic attack of the γ phosphate of the bound ATP molecule (e.g. R334, D382, E383) are known to result in hyperactivity (85). Homologous residues are present in NLRP3 (e.g. R262, D305, E306) and several of them are associated with CAPS (82). Given these facts, it was hypothesized that the inability to hydrolyze ATP results in NLR hyperactivity across both inflammasome-forming and non-inflammasome forming NLRs.

To examine this model, oligomerization and inflammasome reconstitution assays were conducted in HEK293T cells with mutant NLRP3. Specifically, the R262W construct, a known hyperactive mutant associated with CAPS, was employed (82).

Inflammasome activation (indirectly examined by LDH release following pyroptotic cell death) was enhanced compared to wild type NLRP3 (Figure 4.6). Similar findings were

128 noted for the Walker B mutant (WB: D302A/D305A/E306A), a combination of gain of function mutations linked to the CAPS phenotype (84). Consistent with Figure 4.4, mutation of residues responsible for ATP-binding result in decreased pyroptotic cell death (Figure 4.6).

129

Figure 4.5 Proposed Mechanism of NLRP3 ATPase Activity.

A schematic diagram depicting the proposed mechanism of NLRP3 ATPase activity, based on the AAA+ ATPase mechanism described in the literature. The arginine finger motif (R262) is highlighted in purple, the residues of the Walker B motif (D305, E306) are highlighted in orange, and the relevant threonine residue of the Walker A motif

(T233) is highlighted in cyan. The residues of the putative lid region are represented by the yellow area, with the conserved proline residue highlighted (P412). The curved arrows depict a nucleophilic attack on the γ-phosphate moiety of the ATP molecule, catalyzed by the aforementioned residues of the Walker A, Walker B, and arginine finger motifs in concert with a magnesium ion (Mg2+).

130

Figure 4.6 NLRP3 is Functionally Hyperactive Following Mutation of Amino Acids

Responsible for ATP Hydrolysis.

HEK293T cells, grown in a 24-well plate, were transfected with ASC (5 ng/well), pro- caspase-1 (5 ng/well), pro-IL-1β (300 ng/mL), and either wild type or mutant NLRP3 (15 ng/well). After 24 hours of transfection, cell supernatants were collected and LDH assays were performed. WT = wild type, WA = Walker A mutant (226GAAGIGKT233 à

226GAAGIAAA233), WB = Walker B mutant (D302A/D305A/E306A) and Δ403-430 = lid region deletion mutant. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001, ****: p<0.0001).

131 4.2.5 NLRs Require ATP-Binding for Activation, ATPase Activity for

Inactivation.

The loss of function associated with abrogated ATP-binding in tandem with the gain of function associated with abrogated ATP hydrolysis lends itself to a model of NLR activation where the ATP-bound protein is active and licensed to oligomerize upon

PAMP/DAMP recognition, and is deactivated via hydrolysis of the bound ATP molecule in a manner of nucleotide cycling analogous to small G proteins (Figure 4.7) (83).

Interestingly the plant R protein tomato I-2, which has similar domain architecture to

NLR proteins, is known to utilize this exact mechanism of activation suggesting a degree of conserved signaling across at least two kingdoms (141, 142).

132

Figure 4.7 Model for the Activation of NLR Proteins.

A schematic diagram depicting a proposed model for the activation of NLR proteins. At baseline, NLRs bind and hydrolyze ATP to ADP. Detection of PAMPs or DAMPs locks the NLR in an ATP-bound state (i.e. hydrolysis activity stops), licensing it to oligomerize, recruit downstream effector proteins, and mediate a variety of both inflammasome-dependent and independent signaling cascades.

133 4.3 Summary

The data presented in this chapter support the notion that NLR proteins share a common biology with other members of the AAA+ family of ATPases: they possess several conserved ATP binding and hydrolysis motifs (Walker A, Walker B, GxP, hydrophobic lid region) and are theorized to utilize a similar mechanism of ATPase activity. Functionally, the data support a model similar to that observed in plant R proteins wherein ATP-binding licenses the activation of NLR proteins in response to inflammatory stimuli and hydrolysis mediates the inactivation of these proteins. This is noted in that mutations thought to impair ATP-binding prevent the oligomerization and downstream functions of these proteins, whereas the hyperactive CAPS mutants are associated with a loss of ATPase activity. Further work is required to validate this paradigm of activation across the NLR family, but it may represent a common mechanism of action for both inflammasome-forming and non-inflammasome-forming

NLR proteins (including both NLRP3 and NLRP6). Such a finding would be invaluable in the study of these proteins, and would allow for a thorough screening of protein function across the NLR family.

To evaluate this, the following experiments should be performed. First, to parse the distinct roles of ATP-binding and hydrolysis in the activation step of these proteins, additional mutants should be generated that individually mutate the key residues of the

Walker A and B motifs (i.e. the G, K, and T/S residues of Walker A motif, and the D/E residues of Walker B motif). Given the known overlap of these motifs/residues in terms of binding and hydrolysis, this will help to clarify the roles of specific residues contained therein. Second, ATP-binding and hydrolysis assays will need to be performed on all of

134 the aforementioned mutants (using ATP-sepharose column elution assays for the former and hydrolysis assays on purified protein for the latter). This data, in combination with the functional and oligomerization data presented, will allow for the precise mechanisms of NLR activation to be uncovered, especially when supplemented with data from known hyperactive mutations. Ultimately, crystal structures with and without bound ATP will be required to confirm the validity of these conclusions, but that well exceeds the scope of this PhD thesis.

135 Chapter 5: Caspase-4 and Gasdermin D Trigger the NLRP3 Inflammasome

Through the Regulation of Mitochondrial Reactive Oxygen Species

136 5.1 Rationale

Pyroptosis is an inflammatory form of programmed cell death mediated by the cleavage of gasdermin D (GSDMD) downstream of both canonical and non-canonical caspase-4 inflammasome activation (105, 106, 111). The inflammatory proteases caspase-1, caspase-11 (mouse) and 4 (human) are the main effectors of gasdermin D cleavage, that leads to insertion of the N-terminal domain into the cell membrane, pore formation and pyroptotic cell death (111, 112). The non-canonical caspase-4/11 inflammasome is activated by intracellular lipopolysaccharide (LPS) bound and co- transported into the cell by AB5 bacterial toxins such as Cholera toxin (101, 102).

Pyroptosis has also been implicated in a few invasive Gram-negative infections, including Salmonella and Shigella (24, 146).

The non-canonical caspase-4 inflammasome is inextricably linked to the NLRP3 inflammasome. NLRP3 is an intracellular pattern recognition receptor that senses multiple pathogen and host-derived cellular insults. Upon activation by pathogen and damage-associated molecular patterns, canonical NLRP3 signaling triggers oligomerization of the adaptor protein ASC and the effector protein caspase-1 via homotypic interactions, resulting in the formation of the inflammasome that regulates cytokine maturation and pyroptosis. The NLRP3 inflammasome is downstream of a myriad of stimuli that likely converge upon a single common mechanism of activation related to reactive oxygen species (ROS) or potassium efflux (3). Non-canonical and inflammasome-independent functions for NLRP3 have also been identified, including in the regulation of mitochondrial ROS (73). The precise mechanisms governing the

137 crosstalk between NLRP3 and caspase-4/11 inflammasomes in response to intracellular

LPS remains poorly defined.

Shiga toxins (Stx) are cytotoxic exotoxins expressed by the enteric pathogens

Shigella dysenteriae serotype 1 and several E. coli serotypes, including EHEC (also known as Shiga toxin-producing Escherichia coli). A member of a larger family of toxins including Cholera toxin and Pertussis toxin, Shiga toxins consist of a catalytic A subunit and a pentameric, receptor-binding B subunit (147). Several genetic variants of Stx exist, however the broader categories are Stx from S. dysenteriae, and Stx type 1 (Stx1) and Stx type 2 (Stx2) produced by EHEC. Stx is the primary virulence factor in EHEC infection that induces significant inflammation in the development of colitis as well as hemolytic uremic syndrome (HUS), a systemic complication of the disease. In addition to the direct cytotoxic effects of Stx on epithelial and endothelial cells of the gut and kidney, patients with EHEC infections display significant local and systemic inflammation, in the gastrointestinal tract and during HUS respectively, (148) with increased serum concentrations of the pro-inflammatory cytokines IL-1β and TNFα that correlate with disease severity (149, 150).

Stx have previously been shown to activate the NLRP3 inflammasome (52), but the precise mechanism underlying this observation is not defined. Given that Stx production during a clinical EHEC infection would occur in an LPS-rich environment, we hypothesized that Stx-induced inflammation and cell death also involved the non- canonical caspase-4 inflammasome. In this study, we show that the activation of the

NLRP3 inflammasome by the Stx2/LPS complex occurs downstream of mitochondrial

ROS triggered by the non-canonical caspase-4 inflammasome and cleaved GSDMD. The

138 N-terminal domain of GSDMD was enriched at the mitochondria, clarifying the relationship between the caspase-4 and NLRP3 inflammasomes and increasing the understanding of GSDMD-mediated pyroptosis.

139 5.2 Results

5.2.1 Shiga Toxins Activate the Canonical and Non-Canonical Caspase-4

Inflammasome in Human Macrophages.

Previous in vitro studies demonstrated that human macrophages secrete IL-1β following Stx treatment in an NLRP3-dependent manner (52, 151). Consistent with these findings, Shiga toxin 2 (Stx2) treatment of the PMA-differentiated THP-1 human monocytic cells triggered caspase-1 activation and IL-1β maturation in both a concentration- and time-dependent manner (Figure 5.1A-C). To validate these findings in primary human cells, peripheral blood mononuclear cells (PBMC) were treated with Stx2 and also secreted significant amounts of IL-1β (Figure 5.1D). The activation of the inflammasome by Stx2 was caspase-dependent and effectively inhibited by the pan- caspase inhibitor zVAD and the NLRP3 inhibitor glibenclamide (152) (Figure 5.1D).

Immunofluorescence microscopy verified inflammasome assembly and ASC speck formation in THP-1 cells following Stx2 treatment (Figure 5.2A). ASC specks were also found released into the surrounding environment, indicative of pyroptosis (Figure 5.2B).

Indeed, Stx2 induced significant pyroptosis in a concentration- and time- dependent fashion, as determined by the cleavage of caspase-4 and GSDMD, as well as

GAPDH and LDH release into the cell supernatant (Figure 5.3A-D). Stx2 also induced caspase-8 activation as previously reported (Figure 5.3A, B) (153, 154). Pre-treating macrophages with the inhibitors zVAD and glibenclamide significantly reduced Stx2- induced cell death (Figure 5.3E). Collectively, these data show that Stx2 activates both

140 the canonical and non-canonical inflammasomes to trigger IL-1β maturation and pyroptosis.

141

Figure 5.1 Stx2 Activates the Canonical Inflammasome in Human Macrophages.

Figure legend on next page.

142 A. Immunoblot analysis of THP-1 cells treated with increasing concentrations of Stx2 (2,

20, 200 ng/mL) for 6 hours, probing for caspase-1 (antibody directed against N-terminal, detects p11 CARD fragment following caspase-1 activation) in supernatant (SUP) and cell lysates (LYS). B. ELISA for IL-1β in supernatants of THP-1 cells treated with Stx2.

C. Time course of THP-1 cells treated with Stx2 (200 ng/mL). Immunoblot analysis probing for caspase-1 in supernatant (SUP) and cell lysates (LYS). D. ELISA for IL-1β in human peripheral blood mononuclear cells pre-treated with the pan-caspase inhibitor zVAD (50 µM) or glibenclamide (100 µM) followed by challenge with Stx2 for 6 hours

(200 ng/mL) (data courtesy of Adom Bondzi-Simpson). NT = no treatment. Nigericin treatment (NGC, 50 µM) for 1 hour was used as a positive control. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001, ****: p<0.0001).

143

Figure 5.2 Shiga Toxins Induce NLRP3, ASC, and Caspase-1 Containing

Inflammasomes.

Stx2-induced (200 ng/ml) intracellular (A.) and secreted (B.) NLRP3, ASC

(immunolabeling) and caspase-1 (FAM-YVAD-FMK or FLICA) containing specks in

THP-1 cells at 6 hours as determined by confocal microscopy (data courtesy of Hyunjae

Chung). NT = no treatment. Images are representative of n=3 independent experiments.

144

Figure 5.3 Stx2 Triggers Non-Canonical Caspase-4 Inflammasome Activation and

Pyroptosis in Human Macrophages.

Figure legend on next page.

145 A, B. Immunoblot analysis of THP-1 cells treated with Stx2 over both a range of concentrations (2, 20, 200 ng/mL) (A.) and time points (1, 3, 6, 12, 24 hours with 200 ng/mL Stx2) (B.) probing for caspase-4, caspase-8, and gasdermin D cleavage (GSDMD) in the lysate (LYS) and GAPDH release into the supernatant (SUP). C, D. Lactate dehydrogenase (LDH) assay in THP-1 cells treated with increasing concentrations of

Stx2 for 6 hours (C.) and over a 24-hour time course (D.) (data courtesy of Adom

Bondzi-Simpson). E. LDH assay in Stx-treated THP-1 cells pre-treated with pan-caspase inhibitor zVAD (50 µM) or NLRP3 inhibitor glibenclamide (100 µM) at 6 hours (data courtesy of Adom Bondzi-Simpson). NT = no treatment. Positive control is nigericin

(NGC) treatment (50 µM) for 1 hour. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001,

****: p<0.0001).

146 5.2.2 Canonical and Non-Canonical Inflammasome Activation by Stx Requires

NLRP3.

Short-interfering RNAs were utilized in a prior study by Lee et al. to knock down

NLRP3 and to identify a role for the NLRP3 inflammasome in Stx-mediated macrophage activation (52). To prove at a genetic level that Stx-induced inflammasome activation required NLRP3, experiments were first conducted using mouse bone marrow-derived macrophages (BMDM). Unlike human cells, mouse BMDM were unresponsive to Stx2 demonstrating no caspase-1 activation, IL-1β secretion, or cell death (Figure 5.4A-C).

Mouse BMDM were also unresponsive to Stx type 1 (Stx1). Although these data are consistent with previous reports concluding that Shiga toxins were dispensable for

NLRP3 inflammasome activation in murine macrophages (155), surface expression of the

Shiga toxin receptor globotriaosylceramide (GB3 or CD77) (156) was found to be absent in mouse BMDM (Figure 5.4D), but expressed in human THP-1 cells and other mouse tissues such as kidney epithelium (156) (Figure 5.4D, E). In support of these findings,

AlexaFluor 594-conjugated Stx2 readily adhered to the cell membrane and induced CD77 internalization in THP-1 macrophages but not murine BMDMs, as observed by confocal microscopy (Figure 5.4F, G). These results therefore prevented the use of genetic mouse strains to study the biology of Stx-induced inflammasome activation.

To bypass this issue, NLRP3-deficient THP-1 cells were generated using

CRISPR-Cas9-mediated gene editing. NLRP3 expression was undetectable in PMA- differentiated NLRP3 -/- THP-1 cells, as indicated by immunoblotting, verifying the effectiveness of the gene-editing procedure (Figure 5.5A). NLRP3 -/- THP-1 cells expressed normal levels of caspase-1, caspase-4 and ASC (Figure 5.5A). Furthermore,

147 although NLRP3 -/- macrophages were resistant to nigericin, IL-1β production in response to double-stranded DNA, the agonist for AIM2 (34), was intact (Figure 5.6) ruling out both a general defect in inflammasome formation and/or off-target effects induced by the CRISPR construct on other inflammasome genes. When stimulated with

Stx2, caspase-1 cleavage and IL-1β secretion were abolished in NLRP3 -/- cells compared to wild type cells, confirming that Stx2-mediated canonical inflammasome activation is dependent on NLRP3 (Figure 5.5A, B). NLRP3-deficient THP-1 cells also demonstrated a significant reduction in pyroptosis. In response to Stx2 treatment, NLRP3

-/- cells exhibited reduced caspase-4 activation, GSDMD cleavage, and LDH release

(Figure 5.5A, C, D), identifying a role for NLRP3 also in the regulation of the non- canonical inflammasome.

148

149

Figure 5.4 Murine Macrophages are Resistant to Shiga Toxins.

Figure legend on next page.

150 A. IL-1β ELISA and LDH assay (C.) in supernatants of LPS-primed mouse bone marrow derived macrophages (BMDM) treated with Stx1 and Stx2 (data courtesy of Adom

Bondzi-Simpson). B. Caspase-1 immunoblotting in lysates and supernatants of BMDM treated with Stx1 and Stx2 (data courtesy of Adom Bondzi-Simpson). D. Flow cytometric analysis for the Shiga toxin receptor CD77 on the surface of BMDM in comparison to

THP-1 cells (data courtesy of Takanori Komada). E. Immunohistochemistry for CD77 in frozen, murine kidney sections. E-cadherin is used as a positive control (data courtesy of

Takanori Komada). F. Confocal microscopy for AlexaFluor 594 fluorophore-labeled Stx2 bound to THP-1 cells and murine BMDMs. G. Confocal microscopy for CD77 in THP-1 cells and BMDM treated with Stx2 (data courtesy of Hyunjae Chung). GM130 is used as a staining control. Data are representative of three independent experiments. NT = no treatment. Positive control is nigericin (NGC) treatment (50 µM) for 1 hour. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001, ****: p<0.0001).

151

Figure 5.5 Canonical and Non-Canonical Inflammasome Activation by Stx Requires

NLRP3.

Figure legend on next page.

152 A. Immunoblotting for caspase-1, caspase-4, GSDMD, and IL-1β cleavage; and GAPDH release in cell lysates (LYS) and supernatants (SUP) derived from wild type or NLRP3 -/-

THP-1 macrophages treated with Stx2 (200 ng/mL) for 6 hours. B. IL-1β ELISA in supernatants from wild type and NLRP3 -/- THP-1 cells treated with Stx2 for 6 hours. C.

Quantification of caspase-4 cleavage (p30/p43) by densitometry, normalized to wild type caspase-4 cleavage in response to Stx2 treatment (200 ng/mL) for 6 hours. D. LDH assay in supernatants from wild type and NLRP3 -/- THP-1 cells treated with Stx2 for 6 hours

(data courtesy of Adom Bondzi-Simpson). NT = no treatment. Nigericin (50 µM) at 1 hour was used as a positive control. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001,

****: p<0.0001).

153

Figure 5.6 Validation of the NLRP3 -/- (CRISPR) THP-1 Cell Line.

A. IL-1β secretion, measured by ELISA, in response to nigericin (50 µM) and double- stranded DNA (2 µg/mL) in wild type and NLRP3 -/- THP-1 macrophages (data courtesy of Adom Bondzi-Simpson). NT = no treatment. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01,

***: p<0.001, ****: p<0.0001).

154 5.2.3 Shiga Toxin-Mediated Inflammasome Activation is Dependent on Co-

Transported LPS.

Shiga toxins are part of a larger family of bacterial proteins known as AB5 toxins, so designated due to their conserved structure: a single catalytic A subunit complexed with a pentameric, receptor-binding B subunit. In addition to Shiga toxin, this family of protein complexes includes Cholera toxin, Pertussis toxin, and Subtilase cytotoxin. The B subunits of these toxins bind glycan receptors on the cell membranes of host cells to facilitate toxin import (147). The B subunit of the Cholera toxin is known to bind certain serotypes of LPS with a very high affinity, allowing for entry into the cytoplasm and activation of the non-canonical inflammasome (101, 102).

Given the functional conservation of the B subunit across AB5 family members, we reasoned that the Shiga toxin-mediated activation of caspase-4 may be dependent on the co-transport of toxin-associated or bound LPS (Stx/LPS). Not surprisingly, Stx2 preparations contained significant amounts of LPS (Figure 5.7A). To further explore the potential contribution of LPS to Stx-mediated effects, a “low-endotoxin” preparation of

Stx2 was generated and employed in the subsequent experiments. Regardless of the method of Stx production and purification, significant amounts of residual LPS remained bound to Stx by immunoblotting (Figure 5.7B). However, “low endotoxin” Stx2 was much less potent at inducing caspase-1 activation and IL-1β maturation compared to Stx2 isolated directly from E. coli (Figure 5.7C-E). Furthermore, 200 ng/mL of “low- endotoxin” Stx2 pre-incubated with either polymyxin B-coated beads or 100 ng/mL of

LPS (derived from E. coli 0157:H7) for 30 minutes prior to being added to PMA-

155 differentiated THP-1 cells reduced or amplified caspase-4 and GSDMD cleavage respectively (Figure 5.7C).

Similarly, Stx2-induced caspase-1 activation and IL-1β secretion were enhanced by LPS and attenuated by polymyxin B (Figure 5.7C-E), suggesting that the NLRP3 inflammasome is also dependent on activation of the non-canonical inflammasome. LPS alone (as demonstrated with the mock Stx2 preparation) or heat-denatured Stx2 did not activate IL-1β production or cell death consistent with the premise that toxin-mediated internalization of the LPS was required (Figure 5.7F). Together, these data show that while Stx2 alone may also contribute to the activation of the canonical and non-canonical inflammasomes, cellular effects are enhanced by and likely attributable to co-transported

LPS.

156

Figure 5.7 Stx-Mediated Inflammasome Activation is Dependent on Co-Transported

LPS.

Figure legend on next page.

157 A. Immunoblotting of 10 µg of the Shiga toxin preparation used in this paper alongside a dose curve of E. coli LPS (5 µg, 1 µg, 100 ng, 10 ng, 1 ng). Recombinant Stx2 subunit B made in yeast (LSBIO) was used as a positive control for LPS-free Stx2b blotting. B.

Immunoblotting for LPS in a dose curve of the low and high endotoxin preparations of

Stx2. C. Immunoblot analysis of THP-1 cells treated with Stx2 that has been pre- incubated for 30 minutes with either polymyxin B-coated beads or 0157:H7 LPS (100 ng/mL) prior to treatment for 6 hours, probing for caspase and GSDMD cleavage in the cell lysate (LYS) and supernatants (SUP). D, E. ELISA for IL-1β in supernatants of

THP-1 cells treated with Stx2 that has been pre-incubated for 30 minutes with either

0157:H7 LPS (100 ng/mL) (D.) or polymyxin B-coated beads (E.) prior to treatment for 6 hours. F. ELISA for IL-1β in THP-1 cells treated with either heat-inactivated (HI) Stx2

(200 ng/mL) or a bacterial extract (200 ng/mL) derived from a mock Stx2 preparation

(mock) (data courtesy of Adom Bondzi-Simpson). NT = No treatment. Positive control is nigericin (NGC) treatment (50 µM) for 1 hour. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01,

***: p<0.001, ****: p<0.0001).

158 5.2.4 Shiga Toxin-LPS-Mediated Non-Canonical Inflammasome Activation Induces

Mitochondrial ROS Upstream of NLRP3.

The precise sequence of events leading to NLRP3 inflammasome activation in response to the Stx2/LPS complex is unknown. In this context, K+ efflux and mitochondrial ROS (mitoROS) may represent common pathways that converge on

NLRP3 (3). In addition, previous studies have linked endoplasmic reticulum (ER) stress and IRE1α signaling to the generation of mitoROS and subsequent NLRP3 inflammasome activation (157).

Based on prior reports of Stx inducing ER stress responses (154, 158, 159), we first investigated the role of these pathways in Stx2/LPS-treated THP-1 cells. Stx2/LPS stimulation of THP-1 cells for 6 hours did not result in IRE1α phosphorylation, nor did it trigger the splicing of XBP-1 (Figure 5.8A-C). Furthermore, an IRE1α-deficient THP-1 cell line (CRISPR) responded normally to Stx2/LPS in terms of inflammasome activation, as compared to wild-type cells (Figure 5.8D, E). Stx2/LPS also did not activate any other arm of the ER stress response over 6 hours, as indicated by the lack of both PERK phosphorylation and ATF6 cleavage (Figure 5.8A). Thus, Stx2/LPS does not activate the inflammasome as a result of the ER stress response.

159

Figure 5.8 Shiga Toxin/LPS-Induced Inflammasome Activation is Independent of the ER Stress Response.

Figure legend on next page.

160 A. Immunoblotting for IRE1α, PERK and ATF6 pathways in THP-1 cells treated with

Stx2 (200 ng/mL) over a number of time points. B. PhosTag SDS-PAGE of THP-1 cells treated with Stx2 (200 ng/mL) over a number of time points, analyzed by immunoblotting

(data courtesy of Hyunjae Chung). C. PCR reactions characterizing the splicing of XBP-1 in response to Stx2 treatment (200 ng/mL) over various time points in wild type and

IRE1α -/- (CRISPR) cells. D, E. IL-1β secretion, measured by ELISA (D.), and inflammasome activation, demonstrated by immunoblotting of cell lysates (LYS) and supernatants (SUP) for caspase, GSDMD, and IL-1β cleavage and GAPDH release (E.), in response to Stx2 treatment (200 ng/mL) over 6 hours in wild type and IRE1α -/-

(CRISPR) cells. NT = no treatment. Positive controls are nigericin (NGC) (50 µM) for 1 hour and tunicamycin (TC) (10 µg/mL) treatment for 1 and 3 hours for NLRP3 inflammasome activation and ER stress respectively. Data are representative of n=2 independent experiments

161 The cleavage of GSDMD following non-canonical inflammasome activation is known to activate the NLRP3 inflammasome, but the mechanisms through which this occurs remain poorly characterized (105). Upon cleavage, the 30 kDa GSDMD N- terminal fragment is a pore-forming protein with an affinity for the inner leaflet of the cell membrane, as well as microbial and other cardiolipin-containing membranes (111,

112). In agreement with these findings, cleaved GSDMD was found to localize to both crude membrane and mitochondrial subcellular fractions following Stx2 treatment, suggesting that it may play a role in mitoROS production (Figure 5.9A, B). Live cell imaging with MitoSOX Red showed that Stx2/LPS treatment potently upregulated mitoROS over time, a response that was limited by pre-treating macrophages with either caspase-4 or pan-caspase inhibitors (Figures 5.10A and 5.11). Furthermore, Stx2/LPS induction of mitoROS was substantially reduced in NLRP3 -/- and GSDMD -/- THP-1 cells generated using CRISPR-Cas9 gene editing (Figures 5.10B, C and 5.12). Stx2/LPS- induced caspase-4 cleavage was not affected by GSDMD-deficiency, establishing its position upstream of GSDMD and mitoROS production (Figure 5.10D). GSDMD -/- cells also demonstrated a complete loss of caspase-1 cleavage and IL-1β secretion, as well as reduced pyroptotic cell death (Figure 5.10D-F). Together, these data suggested that mitoROS production in response to Stx2/LPS was regulated by the inflammasome and

GSDMD.

To further probe the impact of ROS on canonical and non-canonical inflammasome activation, N-acetylcysteine (NAC), a glutathione precursor and general cellular antioxidant, was employed. In Stx2/LPS-treated cells, NAC reduced caspase-1 cleavage, IL-1β secretion, GSDMD cleavage, and pyroptosis in a concentration-

162 dependent fashion at 6 hours but did not affect caspase-4 cleavage, verifying that ROS played a major role in Stx2/LPS activation of the canonical NLRP3 inflammasome

(Figure 5.13A). NAC was as effective as the caspase-4 inhibitor LEVD-FMK and pan- caspase inhibitor zVAD-FMK at preventing GSDMD cleavage, caspase-1 cleavage and

IL-1β maturation (Figure 5.13B, C). The observed reduction in GSDMD and caspase-1 cleavage, but not caspase-4 activation, following NAC treatment was consistent with a feed-forward mechanism whereby caspase-4 initially cleaved a small pool of GSDMD to trigger mitoROS, which in turn activated the NLRP3 inflammasome and caspase-1 to amplify GSDMD processing and mitoROS generation (Figure 5.13E).

To explore this possibility, caspase-4 and GSDMD cleavage was evaluated more closely using fractionation studies in NLRP3 -/- THP-1 cells. Although caspase-4 activation was reduced and GSDMD cleavage was barely detectable in whole cell lysates of Stx2/LPS-treated NLRP3 -/- macrophages (Figure 5.5A, Figure 5.10D), cleavage products for both were clearly seen in the crude membrane fraction containing mitochondria (Figure 5.13D). Furthermore, caspase-4 inhibition using LEVD-FMK reduced caspase-4 and GSDMD processing, confirming that the non-canonical inflammasome is activated by Stx2/LPS in the absence of NLRP3 (Figure 5.13D).

163

Figure 5.9 GSDMD p30 Localizes to Membrane and Mitochondrial Fractions of

Human Macrophages Following Stx/LPS Treatment.

A, B. Immunoblotting for caspase-4, GSDMD, and organelle/membrane markers following either crude subcellular fractionation (A) or subcellular fractionation by sucrose gradient/ultracentrifugation (B) of wild type THP-1 cells treated with Stx2/LPS for 6 hours (200 ng/mL). Wild type THP-1 membrane fraction lysate (from both untreated and Stx2/LPS-treated samples) was used as a positive control for GSDMD and caspase-4 cleavage. HOMO = homogenate, CYTO = cytoplasmic fraction, MEMB = crude membrane fraction, and MITO = mitochondrial fraction. Fractionations performed with the assistance of Hyunjae Chung.

164

Figure 5.10 Stx/LPS-Mediated Non-Canonical Inflammasome Activation Induces

Mitochondrial ROS Production To Drive Activation of the Canonical

Inflammasome Upstream of NLRP3.

Figure legend on next page.

165 A. Quantification of MitoSOX Red fluorescence (using Incucyte Zoom automated fluorescent microscopy) in THP-1 cells pre-treated with the caspase-4 inhibitor LEVD

(100 µM), the pan-caspase inhibitor zVAD (100 µM), or the antioxidant N-acetylcysteine

(NAC) (25 mM) followed by challenge with Stx2/LPS for 6 hours (200 ng/mL).

Stx2/LPS treatment was compared to Stx2/LPS + inhibitor. B. Quantification of

MitoSOX Red fluorescence (using Incucyte Zoom automated fluorescent microscopy) in wild type and NLRP3 -/- THP-1 cells challenged with Stx2/LPS for 6 hours (200 ng/mL).

WT cells compared to NLRP3 -/- cells. C. Quantification of MitoSOX Red fluorescence

(using Incucyte Zoom automated fluorescent microscopy) in wild type and GSDMD -/-

THP-1 cells challenged with Stx2/LPS for 6 hours (200 ng/mL). WT cells compared to

GSDMD -/- cells. D. Immunoblotting for caspase-1, caspase-4, GSDMD, and IL-1β cleavage; and GAPDH release in cell lysates (LYS) and supernatants (SUP) derived from wild type, NLRP3 -/-, or GSDMD -/- THP-1 macrophages treated with Stx2/LPS (200 ng/mL) for 6 hours. IL-1β ELISA (E) and LDH assay (F) in supernatants from wild type and GSDMD -/- THP-1 cells treated with Stx2/LPS (200 ng/mL) for 6 hours. NT = No treatment. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001, ****: p<0.0001).

166

Figure 5.11 Inhibitor Effects on Stx2/LPS Mediated ROS Production.

A. MitoSOX Red staining (fluorescence and DIC microscopy superimposed), using

Incucyte Zoom automated fluorescent microscopy, in THP-1 cells pre-treated with the caspase-4 inhibitor LEVD (100 µM), the pan-caspase inhibitor zVAD (100 µM), or the antioxidant N-acetylcysteine (NAC, 25 mM) followed by challenge with Stx2 for 6 hours

(200 ng/mL).

167

Figure 5.12 Gene Knockout Effects on Stx2/LPS Mediated ROS Production.

A. MitoSOX Red staining, using Incucyte Zoom automated fluorescent microscopy, in wild type and NLRP3 -/- THP-1 cells challenged with Stx2 for 6 hours (200 ng/mL). B.

MitoSOX Red staining, using Incucyte Zoom automated fluorescent microscopy, in wild type and GSDMD -/- THP-1 cells challenged with Stx2 for 6 hours (200 ng/mL). Images are representative of n=3 independent experiments.

168

Figure 5.13 NLRP3 and GSDMD Are Required For Shiga Toxin/LPS-Induced ROS

Generation.

Figure legend on next page.

169 A. Immunoblot analysis of THP-1 cells pre-treated with increasing concentrations of

NAC (1, 5, 25 mM) prior to challenge with Stx2/LPS (200 ng/mL) for 6 hours probing for caspase and GSDMD cleavage and GAPDH release in the cell lysates (LYS) and supernatants (SUP) respectively. B. Immunoblot analysis of THP-1 cells pre-treated with the caspase-4 inhibitor LEVD (100 µM), the pan-caspase inhibitor zVAD (100 µM), or the antioxidant NAC (25 mM) followed by challenge with Stx2/LPS for 6 hours (200 ng/mL), probing for caspase and GSDMD cleavage and GAPDH release in the cell lysates (LYS) and supernatants (SUP) respectively. C. IL-1β ELISA in supernatants from

Stx2/LPS stimulated THP-1 cells pre-treated with the caspase-4 inhibitor LEVD (100

µM), the pan-caspase inhibitor zVAD (100 µM), or the antioxidant N-acetylcysteine

(NAC) (25 mM) at 6 hours (200 ng/mL). NT = no treatment. Positive control is nigericin

(NGC) treatment (50 µM) for 1 hour. Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01, ***: p<0.001,

****: p<0.0001). D. Immunoblotting for caspase-4, GSDMD, and mitochondrial membrane marker VDAC following crude subcellular fractionation of NLRP3 -/-

(CRISPR) THP-1 cells treated with Stx2/LPS alone or Stx2/LPS + LEVD-FMK (5 µM) for 6 hours (200 ng/mL). HOMO = homogenate, CYTO = cytoplasmic fraction, MEMB

= crude membrane fraction. Fractionations performed with the assistance of Hyunjae

Chung. E. Model for Stx2-induced canonical NLRP3 inflammasome activation downstream of non-canonical inflammasome activation.

170 5.3 Summary

In this study, we shed light on the virulence properties of enterohemorrhagic E. coli Shiga toxin and the mechanism by which it activates the inflammasome. First, and not surprisingly, the effects of Stx2 on inflammasome activation are highly dependent on the co-transport and cellular internalization of LPS bound to the toxin. We show that

Stx2/LPS activates the non-canonical caspase-4 inflammasome and promotes GSDMD cleavage, a key driver of mitochondrial ROS production upstream of the canonical

NLRP3 inflammasome (Figure 5.13E). The NLRP3 inflammasome and caspase-1 in turn potentiate GSDMD cleavage, mitoROS generation, IL-1β maturation and pyroptosis.

Hemorrhagic colitis is the primary and presenting symptom in patients infected with EHEC. It has been proposed that Stx produced by EHEC cross the gastrointestinal epithelium in order to reach the vasculature to cause damage at peripheral sites (i.e. the kidney) (160). Through its cytotoxic actions, Stx directly cause apoptosis and cell death of the gastrointestinal epithelium (160). Inflammation induced at the intestinal interface may be linked to the development of systemic complications, such as HUS, due to compromised barrier function, increased intestinal permeability, and the severity of colitis (161, 162). A number of clinical and experimental studies place IL-1β as a central mediator in EHEC-induced inflammation and disease (149, 150, 163). Alongside its local pro-inflammatory role in the gut, systemic IL-1β upregulates the genes involved in CD77 synthesis in renal cells, thereby increasing the sensitivity and susceptibility of the kidney to Stx-induced injury (164). Thus, the activation of the non-canonical caspase-4 and canonical NLRP3 inflammasomes by Stx likely plays direct and indirect roles in the pathogenesis of EHEC-induced colitis and HUS.

171 Lee et al. demonstrated a role for NLRP3 in Stx-induced inflammasome activation using RNA interference in THP-1 macrophages (52). Our data using CRISPR- mediated gene editing confirm these results. However, our study further probes the mechanism by which Stx activates the NLRP3 inflammasome. We show that LPS co- transported with Stx during cellular internalization activates the non-canonical caspase-4 inflammasome first to trigger mitochondrial ROS production. The generation of mitochondrial ROS then results in NLRP3 inflammasome activation, which in turn regulates caspase-1 activation and further amplifies GSDMD cleavage. Although our results do not rule out an intrinsic ability for Stx to activate the inflammasome, the presence of LPS-bound to Stx is relevant since there would be no circumstance during a clinical EHEC infection where Stx would be produced in an LPS-free environment. Thus,

LPS is a key co-factor in Stx virulence and the pathogenesis of EHEC infection.

These data are consistent with past studies in two respects: 1) they corroborate the link between inflammasome activation and mitochondrial damage/mitoROS production and 2) they support the paradigm of NLRP3 inflammasome activation downstream of the non-canonical inflammasome. In this regard, mitoROS triggered by GSDMD is one of the signals leading to NLRP3 inflammasome activation. Yu et al. demonstrated that canonical inflammasome activation triggers mitochondrial damage and ROS production in a caspase-1-dependent manner (165). Our findings support this hypothesis and further show that GSDMD cleavage and insertion into the mitochondrial membrane likely drives

ROS production. The potential for cleaved GSDMD to insert into the mitochondrial membrane is supported by existing literature given its affinity for mitochondrial lipids

(111, 112). The exact mechanisms underlying ROS induction by GSDMD in this context

172 remain unclear. ROS generation may occur as a result of disruption of the mitochondrial electrochemical gradient or the entry and activity of proteolytic enzymes, such as caspases, on mitochondrial substrates (e.g. components of the electron transport chain)

(166). This latter pathway would be analogous to the granzyme/granulysin-mediated killing of intracellular bacteria (167).

Additionally, our data support the work by Cunha et al. in noting that the activation of the non-canonical inflammasome can serve to activate the NLRP3 inflammasome to result in a feed-forward amplification cascade (117). It is worth noting however that their study limited its scope to K+ efflux as the effector of NLRP3 inflammasome activation, whereas we submit that this process is at least co-dependent on the generation of mitochondrial ROS. In this regard, inhibition of K+ efflux also inhibited

IL-1β production in response to Stx2/LPS (Figure 5.14). Interestingly, caspase-4 activation was found to be partially dependent on NLRP3, suggesting that it may have a role in promoting or stabilizing the formation of the non-canonical caspase-4 inflammasome. This is not an unreasonable supposition, given NLRPs known role in regulating the activity of other caspase-activating complexes (71, 168).

173

Figure 5.14 Potassium Efflux is Required for Stx-Mediated NLRP3 Inflammasome

Activation.

A. IL-1β ELISA in supernatants from THP-1 cells pre-treated with 100 mM extracellular potassium chloride followed by challenge with Stx2 for 6 hours (200 ng/mL). NT = No treatment (data courtesy of Adom Bondzi-Simpson). Data are representative of n=3 independent experiments, with bars representing mean +/- SEM (*: p<0.05, **: p<0.01,

***: p<0.001, ****: p<0.0001).

174 Our results also address the ongoing discrepancy between the effects of Stx in human disease and experimental models in mice. Kailasan et al. examined the role of several EHEC virulence factors in activating the NLRP3 inflammasome (155). Co-culture experiments of wild type and Stx mutant EHEC strains with murine BMDM showed similar IL-1β responses, allowing the authors to conclude that Stx were dispensable for inflammasome activation. Our data suggests that these observations may be due to differential expression of CD77 in murine versus human macrophages and underscore the limitations of extrapolating EHEC studies in mice to human disease. Similarly, our results contradict a prior study that demonstrated a role for ER stress and IRE1α in Stx mediated-apoptosis in macrophages (154). While ER stress has been linked to the NLRP3 inflammasome, our studies using CRISPR-generated IRE1α-deficient THP-1 cells demonstrated no activation of ER stress pathways in response to Stx2/LPS in these cells.

The explanation for this discrepancy is unclear, but likely due to differences in methodology and the use of chemical inhibitors that may have off-target effects.

Collectively, our findings place caspase-4 and NLRP3 as critical pathways activated by Stx, the primary virulence factors associated with EHEC infection.

Furthermore, we describe a central role for LPS as a virulence co-factor in the inflammatory response to Stx. Finally, we clarify further the relationship between the caspase-4 non-canonical inflammasome and the NLRP3 inflammasome via GSDMD and mitochondrial ROS production in human macrophages. This finding provides further insight into the pathogenesis of EHEC infection and development of Stx-induced inflammation and systemic disease.

175 Chapter 6: Discussion and Future Directions

176 6.1 Discussion and Future Directions

6.1.1 Overview

NOD-like receptors are a family of pattern recognition receptors expressed in a variety of tissue types and known to regulate a variety of signaling cascades. Some are responsible for the maturation and secretion of pro-inflammatory cytokines through the formation of caspase-1 activating inflammasomes. Others, such as NLRP6 and NLRP12, have been reported to regulate cell pathways that govern both the growth and the immune response to stimuli, including the MAPK and NF-κB pathways. The inflammasome- independent functions of NLRs remain poorly defined, and recent studies in the NLR field suggest that the functions of these genes may be much more diverse than initially thought.

Furthermore, not all inflammasomes are NLR-dependent: the non-canonical caspase-4 inflammasome, a recently characterized death-inducing complex, is known to be a key driver of the inflammatory response. Though it is known to cause NLRP3 inflammasome activation, the mechanisms driving this activation event have been hitherto understudied.

The ultimate goal of this thesis was to explore the biology of these caspase-1 inflammasome-independent signaling pathways in the pathogenesis of inflammatory disease. The discussion will proceeds as follows with the first section discussing the biology of NLRP6, a non-inflammasome-forming NLR, in terms of expression and biochemistry that were extensively studied in Chapters 3 and 4 respectively. A separate section will discuss the biology of the non-canonical caspase-4 inflammasome and the

177 mechanisms through which it activates the NLRP3 inflammasome, as was presented in

Chapter 5.

6.1.2 NLRP6 Project: Expression and Gene Regulation

Chapter 3 explored the biology of NLRP6: a non-inflammasome-forming NLR whose function remains both poorly characterized and controversial. When this thesis project was undertaken, only a handful of papers had been published on the subject and these had exclusively been conducted in murine intestinal tissue. Since this time, the postulated role of NLRP6 in the gut has been hotly contested with some groups suggesting that the protein has direct roles in negatively regulating both inflammatory and tumorigenic signaling pathways, and others suggesting that the observed phenotypic differences are exclusively as a result of changes to the intestinal microbiome (76, 79, 87,

88, 97). Both of these conclusions remain in doubt due to conflicting data, a lack of demonstrable protein expression in some of the studied cell types, and improperly controlled experimentation.

When the studies conducted in Chapter 3 began, only two key facts were known for certain: 1) Nlrp6 mRNA expression was found to be highest in murine intestine, liver, and kidney and 2) NLRP6 had not been demonstrated to form a canonical inflammasome by biochemical means in any of the tested models (79, 86, 88). As such NLRP6 was classified as a non-canonical NLRP (in that it does not form a caspase-1 activating inflammasome) and, given the high level of expression and lack of study in the kidney as a model system, we sought to characterize the role of NLRP6 in the pathogenesis of kidney disease.

178 Our own expression studies at the transcript level confirmed the published results:

Nlrp6 mRNA expression is predominantly restricted to the intestine, liver, and kidney tissues in mouse. Specifically, the renal Nlrp6 mRNA expression was restricted to the glomerular compartment of the kidney. These findings are similar in human, with expression also found in polymorphonuclear cells (PMNs). Surprisingly, when we examined protein level expression of NLRP6, it was only detectable in intestinal samples from both mouse and human. No NLRP6 protein was detected in the kidney of either species.

To determine if this was due to either an inability to solubilize the protein or an inability of the commercially available antibodies to bind a kidney-specific isoform of the protein, we explored both the lysis conditions and epitope maps of the utilized antibodies.

NLRP6 protein was undetectable in kidney under a variety of lysis/protein solubilization methods, whereas it was readily detected in gut under these same conditions.

Furthermore, epitope mapping of the available antibodies revealed that they bind separate and distinct domains of NLRP6, rendering it unlikely that a truncated isoform was responsible for the inability to detect NLRP6 in the kidney by immunoblotting. In support of this conclusion, Wang et al. was unable to detect FLAG-tagged NLRP6 in kidney (or liver) in a transgenic mouse expressing FLAG-NLRP6 under its endogenous promoter

(92).

To further explore the function, if any, of this gene in the kidney, we compared the phenotypes of wild type and Nlrp6-deficient mice in two different models of murine kidney disease: unilateral ureteral obstruction (UUO) and nephrotoxic serum nephritis

(NTS). The former is a fulminant model of fibrotic kidney disease and the latter is a

179 model of autoimmune glomerulonephritis. No significant differences were observed between the two genotypes in either of these models in terms of inflammatory cell recruitment, fibrosis, histological findings, or albuminuria (in the case of the NTS model).

The paradox between the levels of Nlrp6 mRNA expression when compared to protein led us to postulate that some form of post-transcriptional regulation of the gene was taking place. This theory was supported in 2015 in a paper by Halpern et al., which found that Nlrp6 mRNA is sequestered in the nucleus of cells from select tissue types

(139). Specifically, cytoplasmic Nlrp6 mRNA is readily observed in intestinal epithelial cells, whereas the majority of Nlrp6 mRNA is nuclear in hepatocytes. This suggests that there may be a tissue-specific form of gene regulation at play allowing or preventing the translation of NLRP6 protein in select tissue types under certain conditions.

Hypothesizing that Nlrp6 mRNA may be similarly regulated post- transcriptionally in the cells of the kidney, we conducted RNASeq analysis in colon and kidney tissue derived from wild type and Nlrp6 -/- mice. Two specific analyses were performed: 1) a comparison of Nlrp6 transcript structure between colon and kidney tissue and 2) a comparison of the transcriptomes between wild type and knockout mice in both tissues types. The results of both analyses were both significant and novel.

First, comparison of the Nlrp6 transcripts between colon and kidney tissue revealed that there is a kidney-specific splicing event that results in the fusion of three exons from the upstream long, non-coding RNA locus BC024386 to the 5’ end of the

Nlrp6 transcript. This fusion product is not observed in murine colon tissue. Interestingly, an NCBI Gene search revealed that BC024386 is most highly expressed in murine kidney

180 and liver, suggesting that there may be a similar fusion transcript expressed in liver.

Bioinformatic analysis failed to find a Kozak consensus sequence in the lncRNA-Nlrp6 fusion transcript suggesting that this transcript will not be translated in the kidney, consistent with the lack of detectable protein. Furthermore, if the same fusion transcript were expressed in liver, this would lend credence to nuclear sequestration of Nlrp6 transcript in hepatocytes; several of the predominantly nuclear transcripts they characterized (e.g. Malat1) are in fact lncRNAs (139). These findings suggest that a rather unique form of post-transcriptional control regulates Nlrp6 expression in select tissue types, where fusion to an upstream lncRNA prevents translation of the protein. The mechanism(s) underlying and biological significance pertaining to the expression of

Nlrp6 as an lncRNA fusion transcript specifically in kidney and liver remains to be seen, but may relate to the important metabolic roles these organs occupy.

Notwithstanding the generation of a lncRNA-Nlrp6 fusion transcript in the kidney, the gene is not silenced in terms of function. Transcriptomic analysis of wild type and Nlrp6-deficient kidney (but not colon) samples revealed a 210 fold upregulation of

Rb1 in Nlrp6 -/- kidney. RB1 protein was also found to be upregulated in Nlrp6-deficient animals. Given the known role of Rb1 as a tumor suppressor gene, this suggests that

Nlrp6-deficiency may confer resistance to the development of renal cell carcinoma.

Having only recently discovered this upregulation, further work is required to both validate the regulation of the Rb1 transcript and RB1 protein expression and to explore the potential role of Nlrp6 in driving renal tumorigenesis (140). Additionally, the mechanisms underlying this regulation would require elucidation. It is possible that the fusion transcript either serves as a miRNA sink to indirectly mediate miRNA-controlled

181 gene expression or directly regulates the expression of select genes via direct interaction with transcriptional and/or epigenetic machinery (140).

The discovery of a kidney-specific lncRNA-Nlrp6 fusion transcript is entirely novel: no such phenomenon has been described in the field of NLR biology and a literature search has failed to find any example of a translational silencing event mediated in such a fashion. The fact that the gene in gut (expressed as a protein) and in kidney

(expressed as an lncRNA-mRNA fusion transcript) may have radically different functions depending on the nature of their expression is a very significant finding meriting a very detailed study in terms of mechanism and effect. Furthermore, additional work is required to examine the regulation of the NLRP6 gene in different human tissue types, where similarly unique gene regulation events may be taking place.

RNAseq also revealed 4 genes differentially expressed in colon and 42 genes differentially regulated in both kidney and colon between wild type and Nlrp6-deficient mice. Whilst the 42 genes common to colon and kidney were initially ascribed to genetic drift in our mouse lines (owing to the radically different mechanisms of gene regulation across these tissues), pathway analysis revealed 13 genes (including Mecp2, a gene whose differential expression is restricted to colon) that are part of a causal network downstream of Apoc2. Given this finding, additional experiments (e.g. qRT-PCR and immunoblot validation, rederivation of the mouse colony and additional RNAseq, etc.) will need to be done to validate the downregulation of this pathway and to explore the link with Nlrp6 expression.

182 6.1.3 NLRP6 Project: Biochemistry

Having characterized some of the gene expression and regulation of NLRP6,

Chapter 4 sought to better characterize the biochemistry of this protein, and of the NLR family as a whole.

The NLRs are part of the larger AAA+ family of ATPases and several NLRs have been confirmed to bind and hydrolyze ATP (e.g. NLRP3, NLRP7, NLRP12, etc.) (5-7,

82). All studies to date agree that ATP-binding is essential to NLR activation and oligomerization; abrogation of which abolishes downstream NLR function, even in the context of hyperactive, gain-of-function mutations. The role of ATP hydrolysis remains hotly contested; with studies suggesting it has a role for both the activation and inactivation of these proteins (61, 85, 145). The original study by Duncan et al. in NLRP3 considered both possibilities and no subsequent publication has diligently parsed the differing roles of ATP-binding and hydrolysis of this protein (5).

With respect to the NOD proteins (e.g. NOD1 and NOD2), several papers have come to support a model whereby ATP-binding allows for protein activation, whereas

ATP hydrolysis is an inactivating event in a manner of nucleotide-cycling analogous to small G proteins (9, 85, 145). Mutational analysis found that several of the hyperactive

NOD2 mutations associated with the autoinflammatory Blau syndrome affect amino acid residues associated with ATP hydrolysis, as per the common ATPase mechanism shared by AAA+ family members (85). Specifically, the affected residues (contributed from both the Walker B and arginine finger motifs) are thought to drive the activation of a water molecule and facilitate the nucleophilic attack of the γ phosphate of the ATP moiety. The same mutagenesis assays found that mutation of key ATP binding motifs,

183 such as the Walker A site and GxP motif, abrogate NOD2 function (85, 145). Given the conservation of these motifs across NLRs, we postulated that the same roles for ATP- binding and hydrolysis apply to both inflammasome and non-inflammasome-forming

NLRP proteins, namely NLRP3 and NLRP6 respectively. Given the difficulties in studying the NLRP6 protein and the conflicting data in the established literature, we sought to use its structural similarity to a well-studied inflammasome-forming NLR (i.e.

NLRP3) to develop a model of activation whereby we could attempt to elucidate its function as a protein. This would be of most relevance in the gut, where NLRP6 protein is readily expressed and detectable.

To test this hypothesis, we modeled both NLRP3 and NLRP6 using Phyre2, with

APAF-1 as a model protein. The ATP-binding pocket of these NLRPs greatly resembled that of NOD2, with the Walker A and GxP motifs (and surrounding hydrophobic amino acids of the putative lid region) coordinating the ATP molecule and several conserved

ATP hydrolyzing residues contributed from both the Walker B site and, in the case of

NLRP3, a conserved arginine finger motif. These models support the theory that NLRs utilize the same conserved mechanism of ATP-binding and hydrolysis as other AAA+

ATPases (8, 82, 144).

Using a self-designed oligomerization assay, mutation of the Walker A site, the proline of the GxP motif, or the entire lid of the ATP-binding pocket significantly reduced the ability of NLRP3 and NLRP6 to oligomerize. The introduction of the hyperactive CAPS mutation R262W, analogous to a similar mutation in NOD2 (i.e.

R334W) responsible for the development of Blau syndrome, enhanced this oligomerization event. With specific regard to NLRP3, mutagenesis of residues

184 responsible for ATP-binding resulted in decreased activity of the NLRP3 inflammasome

(i.e. caspase-1 cleavage, IL-1β secretion) and decreased NLRP3-driven pyroptosis, when reconstituted in HEK293T cells. Consistent with the oligomerization findings, the introduction of NLRP3 containing the R262W mutation to the HEK293T inflammasome reconstitution system resulted in enhanced NLRP3-dependent pyroptosis.

Overall, these data support a model of NLR activation similar to that seen in the related plant R proteins, specifically tomato I-2: at baseline the proteins cycle between

ATP- and ADP-bound forms, disallowing oligomerization of these proteins (141, 142).

Upon detection of noxious stimuli (or in the presence of mutations that prevent nucleotide hydrolysis), the proteins stop nucleotide cycling and remain in an active, ATP-bound state. This allows for the oligomerization of the NLR proteins, the recruitment of adaptor and effector proteins and, ultimately, the execution of downstream functions. In the case of canonical inflammasome-forming NLRs, these are the maturation of pro-inflammatory cytokines and the induction of pyroptotic cell death. With non-inflammasome-forming

NLRs, these events allow for the direct regulation of inflammatory and growth-associated signaling cascades.

These findings are significant for a number of reasons. First, they support a unified model of NLR activation that encompasses both the inflammasome and non- inflammasome-forming NLR proteins. This is the first work to attempt to parse the differing roles of ATP binding and hydrolysis in NLRP protein activation and downstream function. Second, they provide a set of tools through which the functions of non-inflammasome-forming NLR proteins can be screened: by understanding the conditions that confer hypo or hyperactivity on NLR proteins as a family, novel effector

185 functions can be detected in vitro. Finally, and perhaps more as a matter of diligence, these data correct an improperly cited “fact” in the literature that ATP hydrolysis (as well as binding) is required for NLR activity. The fact of the matter is that in preventing ATP- binding, hydrolysis becomes moot, and researchers have been too quick to ascribe the loss of function to hydrolysis when in fact no appropriately controlled experimentation has been performed (5, 61).

In terms of future directions, there is much work to be done with this project.

First, additional mutants of NLRP3 and NLRP6 need to be generated; the existing Walker

A and Walker B mutants mutate too many key residues simultaneously, some of which are theorized to have different effects in terms of ATP-binding and hydrolysis. By mutating the key residues individually (G, K, T/S residues of the Walker A motif, D/E residues of the Walker B motif, rather than all at once) and subjecting them to the aforementioned oligomerization and reconstitution assays, these functions can be effectively parsed and their effects on downstream activation can be precisely determined. Second, ATP-binding and hydrolysis studies will need to be performed on all of the generated mutants, to confirm their theorized effects. The ability to conduct these assays to date has been hindered due to the difficulty in purifying NLR proteins. The current strategy, which appears to be promising, is to purify GFP-tagged mutant proteins using the proprietary GFP Trap technology: in essence, an affinity column designed for the purification of GFP-tagged proteins. Once purified, ATP-binding can be quantified using ATP-Sepharose columns and hydrolysis can be quantified through the detection of both generated ADP and inorganic phosphate during controlled ATPase reactions.

186 Finally, additional inflammasome reconstitution experiments will be required to validate the effects of the new mutants on downstream NLR function.

By properly understanding the biochemistry of these proteins, therapeutics can be designed to specific target these proteins and limit their pro-inflammatory effector functions. Furthermore, in understanding mutations that confer hypo and hyperfunctionality in NLRs, the function of novel and understudied NLRs can be screened by in vitro assays in cell lines.

6.1.4 Non-Canonical Caspase-4 Inflammasome Project

Chapter 5 explored the mechanism by which the non-canonical, caspase-4 inflammasome activates the canonical NLRP3 inflammasome. This project evolved from two observations: 1) that Shiga toxin 2 derived from enterohemorrhagic Escherichia coli

(EHEC) activates both the NLRP3 and caspase-4 inflammasomes and 2) that these activation events are dependent on co-transported lipopolysaccharide in a manner analogous to that previously described with Cholera toxin.

In terms of clinical relevance, this finding is highly significant: clinical EHEC infections arise in the LPS-rich intestinal tract and as such LPS-driven, toxin-dependent inflammasome activation is almost certainly a contributing factor to disease pathogenesis.

In fact, our model demonstrates that LPS, rather than any intrinsic activity of the toxin, is the main driver of the acute, leukocyte-derived inflammatory response, at least in terms of IL-1β secretion. Specifically, over the time points studied, there was no induction of the ER stress response that is characteristic of the catalytic activity of the A subunit of

Shiga toxins. Given the known correlation between serum IL-1β levels and disease

187 severity, therapeutic targeting of the non-canonical caspase-4 inflammasome may prove to be fruitful in the treatment of EHEC-mediated hemorrhagic colitis and the prevention of hemolytic uremic syndrome, a known systemic complication of this infection.

Notwithstanding the practical applications of the aforementioned findings, the far more exciting data is that which characterizes the cross-talk between the non-canonical caspase-4 inflammasome and the canonical NLRP3 inflammasome. Previous studies have found that activation of the former can drive the activation of the latter, but the mechanisms through which this occurs remain unclear (101, 102, 105, 106). Recent studies have linked NLRP3 inflammasome activation to K+ efflux downstream of the non-canonical inflammasome and gasdermin D (GSDMD) cleavage, but an in depth analysis of this pathway had not been conducted until this thesis work.

Having noted both canonical and non-canonical inflammasome activation in THP-

1 macrophages following Stx2-LPS treatment, we used inhibitor studies to determine that caspase-4 lies upstream of the NLRP3 inflammasome. This was consistent with the order of signaling identified in previous studies.

Next, we sought to understand the mechanisms through which caspase-4 activity triggers the activation of the NLRP3 inflammasome. When activated, caspase-4 is known to cleave gasdermin D: a 50 kDa protein whose N-terminal fragment is known to have pore-forming properties, with specific affinities for phosphoinositide and cardiolipin-rich lipid membranes (105, 106, 111, 112). Through our earlier experiments with the Stx2-

LPS complex, we noted that the addition of this complex to macrophages results in the generation of mitochondrial ROS (mitoROS) and that the inhibition of this ROS decreased the activity of the NLRP3 inflammasome. Given the affinity of cleaved

188 GSDMD for cardiolipin, we hypothesized that N-terminal GSDMD inserts into the mitochondrial membrane, disrupts proper mitochondrial function, and drives the generation of mitoROS to activate NLRP3. Interestingly, a paper published following the submission of this work found that induction of pyroptosis results in a GSDMD- dependent loss of mitochondrial membrane potential, confirming this hypothesis and supporting our data (166).

Chemical inhibition and gene disruption confirmed that the generation of mitochondrial ROS is dependent on both GSDMD and NLRP3, suggesting a feed- forward loop of activation where the NLRP3 inflammasome serves as a signal amplifying complex downstream of caspase-4 to drive both the production of inflammatory cytokines and end-stage pyroptotic death.

These findings are both novel and significant contributions to the field of inflammasome biology. No group has previously established a direct link between caspase-4-mediated GSDMD cleavage, mitoROS generation, and NLRP3 inflammasome activation. This axis, which likely acts in tandem with GSDMD-dependent K+ efflux, is a potent activator of the NLRP3 inflammasome and ultimately drives the majority of the inflammatory response in our PAMP-driven model of inflammation.

Furthermore, when one considers the known activators of the NLRP3 inflammasome (e.g. K+ efflux, mitoROS generation, lysosomal disruption, calcium mobilization from the endoplasmic reticulum), these findings support a unified model of

NLRP3 biology where this protein serves as an inflammatory signal amplifier downstream of biological membrane disruption (both in the context of the cell membrane and membrane-bound organelles). It seems likely that GSDMD is also able to

189 permeabilize these other membrane-bound organelles, allowing for NLRP3 activation downstream of multiple stimuli. This is sensible biologically for two reasons: 1) it introduces redundancy into the system by utilizing multiple activating pathways upstream of NLRP3, limiting the effect of inhibition of any one pathway and 2) it introduces an additional signal amplification effect to maximize the activation and effector functions of

NLRP3. Additional work is required to explore GSDMD-dependent permeabilization of these membrane-bound organelles and their relative contribution to NLRP3 inflammasome activation.

In the specific example presented in Chapter 5, caspase-4 is the upstream sensor and GSDMD is the upstream effector which triggers said membrane disruption. Other membrane disrupting, NLRP3-activating effector molecules have been characterized, including MLKL: the pore-forming protein that executes cell lysis in necroptosis (118,

119). This suggests that there is considerable cross-talk between the various pathways of programmed necrosis and inflammasome activation, a fact consistent with the inflammatory nature of these pathways. Additional effectors have also been postulated, including other members of the gasdermin family (i.e. GSDMA3). Future work should investigate the role of caspase-4 in this model via CRISPR-mediated gene editing, as well as seek out the existence of additional sensors of membrane disruption upstream of

NLRP3. Furthermore, the existence of other pyroptotic mediators should be examined, given the incomplete loss of cell death downstream of GSDMD-deficiency.

Interestingly, the work conducted in Chapter 5 also revealed a role for NLRP3 in the activation of caspase-4; cleavage of caspase-4 was significantly reduced in cells deficient in NLRP3. The mechanism driving this observation remains to be studied

190 however, knowing that NLRP3 serves as a scaffold for the formation of various caspase- activating complexes, it seems a reasonable hypothesis to suggest that NLRP3 may have a facilitating role in the formation or stabilization of the caspase-4 inflammasome complex, with or without the adaptor protein ASC (71). The existence and importance of such a complex is an important question to be answered in future studies.

6.1.5 Summary

Overall, this PhD thesis has challenged a number of the existing assumptions in the field of NLR biology, has contributed novel and exciting new data to the field, and has laid the groundwork for several fascinating new avenues of study. With the input of additional resources, it is hoped that 1) the diverse functions of NLR genes (including those at the post-transcription/epigenetic level) can be elucidated; 2) the fundamental and conserved biochemistry governing the activity of these proteins can be solved; and 3) the over-arching paradigm of NLRP3 as a signal amplifier downstream of disruptions in cellular homeostasis, including those mediated by the non-canonical caspase-4 inflammasome, can be better understood.

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208 Appendix A: List of Publications

1. Platnich, J.M., Sandall, C.F., Chung, H., Bondzi-Simpson, A., Komada, T., Lau, A., Brandelli, J.R., Chun, J., Trotman-Grant, A., Beck, P.L., Philpott, D.J., Girardin, S.E., Ho, M., MacDonald, J.A., Armstrong, G.D., and Muruve, D.A. 2018. Caspase-4 and Gasdermin D Trigger the NLRP3 Inflammasome Through the Regulation of Mitochondrial Reactive Oxygen Species. Cell Rep Under Review.

2. Komada, T., Chung, H., Lau, A., Platnich, J.M., Beck, P.L., Benediktsson, H., Duff, H.J., Jenne, C.N., and Muruve, D.A. 2018. Macrophage Uptake of Necrotic Cell DNA Activates the AIM2 Inflammasome to Regulate a Proinflammatory Phenotype in CKD. J Am Soc Nephrol. In press.

3. Lemire, P., Robertson, S.J., Maughan, H., Tattoli, I., Streutker, C.J., Platnich, J.M., Muruve, D.A., Philpott, D.J., and Girardin, S.E. 2017. The NLR Protein NLRP6 Does Not Impact Gut Microbiota Composition. Cell Rep 21 (13): 3653- 3661.

4. Lau, A., Chung, H., Komada, T., Platnich, J.M., Sandall, C.F., Choudhury, S.R., Chun, J., Naumenko, V., Surewaard, B.G.J., Nelson, M.C., Ulke-Lemée, A., Beck, P.L., Benediktsson, H., Jevnikar, A.M., Snelgrove, S.L., Hickey, M.J., Senger, D.L., James, M.T., Macdonald, J.A., Kubes, P., Jenne, C.N., and Muruve, D.A. 2017. Immune Surveillance by Renal Phagocytes and Tubular Dipeptidase-1 is Essential to the Pathogenesis of Contrast-Induced Acute Kidney Injury. J Clin Invest Under Review.

5. Muruve, N., Feng, Y., Platnich, J.M., Hassett, D., Irvin, R., Muruve, D., and Cheng, F. 2017. A peptide-based biological coating for enhanced corrosion resistance of titanium alloy biomaterials in chloride-containing fluids. J Biomater Appl 31 (8): 1225-1234.

6. Chung, H., Vilaysane, A., Lau, A., Stahl, M., Morampudi, V., Bondzi-Simpson, A., Platnich, J.M., Bracey, N.A., French, M.C., Beck, P.L., Chun, J., Vallance, B.A., and Muruve, D.A. 2016. NLRP3 regulates a non-canonical platform for caspase-8 activation during epithelial cell apoptosis. Cell Death Differ 23 (8): 1331-46.

7. Chun, J., Chung, H., Wang, X., Barry, R., Taheri, Z.M., Platnich, J.M., Ahmed, S.B., Trpkov, K., Hemmelgarn, B., Benediktsson, H., James, M.T., and Muruve, D.A. 2016. NLRP3 Localizes to the Tubular Epithelium in Human Kidney and Correlates With Outcome in IgA Nephropathy. Sci Rep 6: 24667.

8. Davis, E.M., Platnich, J.M., Irvin, R.T., and Muruve, D.A. 2015. Peptide- Mediated PEGylation of Polysulfone Reduces Protein Adsorption and Leukocyte Activation. ASAIO J 61 (6): 710-7.

209