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Identification of mammalian cell signaling in response to plasma membrane perforation:

Endocytosis of Listeria monocytogenes and The Repair Machinery

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

By

Jonathan Gat-Tze Lam

Graduate Program in Microbiology

The Ohio State University

2018

Dissertation Committee

Stephanie Seveau, Advisor

Dan Wozniak

John Gunn

Li Wu

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Copyrighted by

Jonathan Gar-Tze Lam

2018

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Abstract

The animal plasma membrane is a semi-fluid structural platform that maintains cellular homeostasis by regulating the passage of ions and small molecules in and out of the cell and modulating cell signaling activities. Disruption of its barrier function via mechanical damage or perforation by a pore-forming toxin is quickly followed by a sudden influx of extracellular Ca2+, which triggers efficient plasma membrane repair processes, the mechanisms of which are, to date, not fully elucidated. Efforts to understand cellular responses to plasma membrane damage have resulted in several non- mutually exclusive models of repair, each realized by the use of various cell types damaged using approaches that attempt to replicate normal physiological damage

(mechanical, osmotic, and sheer stress) or damage that occurs under infectious conditions (bacterial pore-forming toxins).

In the context of infection, evolutionarily distinct pathogens including the parasite

Trypanosoma cruzi, the Gram-positive bacterium Listeria monocytogenes, and the non- enveloped Adenovirus have been shown to damage the plasma membrane of non- professional phagocytic cells in order to co-opt the subsequent cellular responses to facilitate their entry into target cells. It was concluded that T. cruzi and Adenovirus mechanically damage or perforate the host cell plasma membrane in order to co-opt a

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Ca2+ influx-dependent repair mechanism involving the exocytosis of lysosomes, release of acid sphingomyelinase, invagination of the host plasma membrane and endocytosis of the invading pathogen along with the damaged membrane. It was revealed that addition of the cholesterol-dependent cytolysin (CDC) pore-forming toxin Streptolysin O, which forms ~30 nm diameter proteinaceous pores in cholesterol-containing membranes, could further facilitate the efficient entry of these pathogens. Studies using the related CDC pore-forming toxin listeriolysin O (LLO) showed that L. monocytogenes entry into hepatocyte epithelial cells also requires Ca2+ influx subsequent to LLO-mediated perforation of the target cell suggesting that like T. cruzi and Adenovirus, L. monocytogenes could co-opt the repair machinery to gain entry. Using a combination of biochemical assays and live-cell fluorescence resonance energy transfer imaging, we found that LLO-mediated plasma membrane perforation and influx of extracellular Ca2+ activates a signal cascade involving the recruitment and activation of a conventional protein kinase C at the plasma membrane, activation of the central actin regulator, the

Rho GTPase Rac1, and induction of Arp2/3-dependent F-actin polymerization leading to bacterial internalization. Inhibition of the cPKC/Rac1/Arp2/3 pathway prevents L. monocytogenes entry, but does not prevent membrane resealing, revealing that, in contract to the mechanism of entry of T. cruzi and Adenovirus, LLO-dependent L. monocytogenes endocytosis is distinct from the resealing machinery.

An additional focus of this work has been on unifying the mutually non-exclusive models of plasma membrane repair, which include the Patch hypothesis, lysosomal

ii exocytosis and subsequent endocytosis, exosome release, and ectocytosis/microvesicle shedding. Much of the pioneering work that led to these models required the use of morphologically unique cells (Xenopus laevis oocytes, sea urchin eggs, squid giant axons) damaged via microneedle puncture, laser ablation, or transection techniques.

Further studies using various somatic mammalian cells including myocytes, neurons, epithelial and endothelial cells damaged via mechanical or sheer stress, or exposed to pore-forming toxins have also led to the identification of numerous proteins involved in the resealing process, many of which are cell type specific. The goal of this work was to provide a platform in which to study plasma membrane resealing in any cell type in a high-throughput manner in order to a) quantify the efficiency of resealing, b) identify new proteins involved in the resealing process, c) determine any overlap across different cell types, and d) unify the various models or resealing into a global system of repair mechanisms. Here we describe a high-throughput microplate-based assay to quantify the membrane resealing efficiency of cells exposed to the pore-forming toxin listeriolysin O using a spectrofluorometric plate reader. Additionally, image cytometry was incorporated to automatically enumerate target cells expressing nuclear localized histone 2B-GFP

(HeLa H2B-GFP) before and after damage to account for differences in cell counts due to damage-induced cell detachment. Lastly, as a proof of concept, a siRNA library covering

287 gene targets involved in membrane trafficking, autophagy, lysosome biogenesis and function, ectocytosis, and cytoskeletal dynamics was used to identify proteins involved in the resealing of LLO-perforated HeLa cells. 26/287 knockdown conditions caused a defect in repair, which correspond to clathrin-mediated endocytosis (adaptor related

iii proteins, dynamin), vesicular fusion and fission (Rab, SNARE, and exocyst proteins), plasma membrane stabilization (Annexins), and vesicle packaging (COP and ESCRT proteins). Surprisingly 19/287 knockdown conditions actually improved repair indicating that plasma membrane repair can potentially be negatively regulated. These preliminary findings confirmed the applicability of our high-throughput assay in identifying proteins involved in plasma membrane repair, but confirmation screens, pathway analyses, use of different cell types and damaging conditions are still required to meet the goals of this work.

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Dedication

Dedicated to my parents Dr. Juan Lam and Dr. Flora Yeung for their guidance and support that have made me the man and scientist I am today. To my sister Jennifer Lam-

Gerjarusak who has been a model of dedication and hard work. And lastly, to my beautiful wife Katy Elizabeth Lam, for whom this work lays the future for our family.

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Acknowledgments

I would like to acknowledge the following people who have been supportive in and out of the lab: Christopher Phelps, Dr. Jordan Angle, Garrett Smith, Dr. Joanna

Marshall, Lauren Johnson, Siavash Azari, Jasneet Singh, Madison McQuate, Megan Linz and Bella Cho. I’d like to thank all of the professors in the Department of Microbiology and Department of Microbial Infection and Immunity who have been influential in my academic and technical training. To the researchers who have provided the scientific breakthroughs that helped my projects come to fruition: Dr. Adam Hoppe and Dr. Joel

Swanson (University of Michigan Medical School) who developed the fluorescence resonance energy transfer (FRET) stoichiometry method; Dr. Xiaoli Zhang, Eric

McLaughlin, and Dr. Chi Song (The Ohio State University) for the statistical analyses,

Dr. Alexandra Newton (University of California San Diego) for her guidance on the study of protein kinase C activation and providing us with protein kinase C vectors; Dr.

Stephen Vadia (The Ohio State University), a former graduate student of the laboratory, for his contribution to the identification of the novel listeriolysin O-dependent internalization mechanism of L. monocytogenes, and Dr. Sarika Pathak-Sharma (The

Ohio State University), a former postdoctoral trainee of the laboratory, for the initial development of the high-throughput membrane resealing assay. I would also like to thank

Dr. Robert Tabita and Dr. Jesse Kwiek (The Ohio State University) for allowing me to

vi use their French Press and SpectraMax i3X + MiniMax300, respectively. To my committee members Dr. John Gunn, Dr. Dan Wozniak, and Dr. Li Wu (The Ohio State

University), your guidance and input throughout my graduate career have ensured the success of my projects. Most importantly, to Dr. Stephanie Seveau developed the present research program, which was funded by the National Institutes of Health

(RO1AI107250), and dedicated her time to train me as a research scientist.

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Vita

2009-2011 Laboratory Technician, Department of Biology, University of California San Diego

2011 B.S. Biochemistry/Chemistry, University of California San Diego Revelle College

2011-2012 Contracted Research Associate Synthetic Genomics Inc. La Jolla, Ca

2013-2015 Graduate Teaching Associate Department of Microbiology, The Ohio State University

2016-2017 Vice President of the Students for the Advancement of Microbiology, The Ohio State University

Publications

Lam, J., et al., High-Throughput Measurement of Plasma Membrane Resealing Efficiency. Journal of Visualized Experiments, in Press.

Lam, J., et al., Host cell perforation by listeriolysin O (LLO) activates a Ca(2+)- dependent cPKC/Rac1/Arp2/3 signaling pathway that promotes L. monocytogenes internalization independently of membrane resealing. Molecular Biology of the Cell, 2017. 29(3): p.270-284.

Pathak-Sharma, S., et al., High-Throughput Microplate-Based Assay to Monitor Plasma Membrane Wounding and Repair. Frontiers in Cellular and Infection Microbiology, 2017. 7: p. 305.

Fields of Study

Major Field: Microbiology

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Table of Contents

Abstract ...... i Dedication ...... v Acknowledgments...... vi Vita ...... viii List of Tables ...... x List of Figures ...... xi Chapter 1: Introduction ...... 1 1.1 Mammalian Plasma Membrane Biology ...... 2 1.2 Plasma membrane damage under physiological and pathological conditions ..... 7 1.3 Models to Study Plasma Membrane Resealing ...... 10 1.4 Plasma Membrane Repair Models ...... 17 1.5 Signaling and repair mechanisms in response to plasma membrane damage in the context of host-pathogen interactions ...... 26 1.6 Intracellular Infectious cycle of Listeria monocytogenes ...... 28 1.7 Implications and Perspective ...... 31 2 Chapter 2. Plasma Membrane Damage, A Novel Mechanism of Pathogen Entry ... 33 2.1 Introduction ...... 33 2.2 Results ...... 36 2.3 Discussion ...... 55 2.4 Materials and Methods ...... 62 3 Chapter 3: A High-Throughput Assay to Quantify Membrane Repair Efficiency ... 77 3.1 Introduction ...... 77 3.2 Protocol ...... 82 3.3 Results ...... 95 3.4 Discussion ...... 119 Chapter 4: Future Directions and Final Comments ...... 128 References ...... 133 Appendix A. Supplemental Data Chapter 2 ...... 148

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List of Tables

Table 2.1 LLO induces Rac1 activation in a Ca2+- and pore-dependent manner ...... 41 Table 2.2: Rac1 is activated downstream of a conventional PKC ...... 52 Table 3.1 Fluorophore excitation/emission and imaging cytometer bandpasses...... 101 Table 3.2 Preliminary siRNA library hits ...... 115 Table A.1 Densitometry analysis for Figure 3.1 B ...... 152 Table A.2 Statistics for Table 3.1...... 152 Table A.3 Statistics for Figure 3.3D...... 153 Table A.4. Statistics for Figure 3.8...... 153 Table A.5 Statistics for Table 3.2...... 154 Table A.6 Primers used in this study...... 154

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List of Figures

Figure 1.1 The plasma membrane ...... 5 Figure 1.2 Cholesterol dependent cytolysin crystal structures ...... 14 Figure 1.3 Listeriolysin O crystal structure ...... 16 Figure 1.4 Plasma membrane repair models ...... 25 Figure 1.5 The Listeria monocytogenes intracellular lifecycle 122 ...... 29 Figure 2.1: Rac1, RhoA, and Cdc42 knockdown efficiencies ...... 37 Figure 2.2: Role of Rac1 in LLO-dependent entry of L. monocytogenes...... 38 Figure 2.3: FRET stoichiometry to monitor Rac1 activation by LLO...... 42 Figure 2.4: LLO induces F-actin rearrangement...... 45 Figure 2.5: Role of Arp2/3 in LLO-dependent entry of L. monocytogenes...... 47 Figure 2.6: Role of a conventional PKC in LLO-dependent entry of L. monocytogenes. 48 Figure 2.7: LLO induces PKCα translocation to the plasma membrane...... 49 Figure 2.8: A PKC is activated by LLO...... 51 Figure 2.9: The LLO-dependent entry pathway is dispensable for membrane resealing. 54 Figure 2.10: Model for LLO-mediated L. monocytogenes entry...... 55 Figure 3.1 Experimental Design ...... 84 Figure 3.2: Cell Counting accuracy ...... 97 Figure 3.3: Propidium iodide fluorescence measurement is not affected by Histone 2B- GFP expression ...... 98 Figure 3.4: Cell enumeration is unaffected by PI fluorescence...... 100 Figure 3.5: Measuring plasma membrane resealing efficiency...... 102 Figure 3.6: TO-PRO-3 as an alternative dye to assess membrane resealing...... 104 Figure 3.7: Effect of LLO concentration on the resealing efficiency, Z-factor, and SSMD...... 106 Figure 3.8: Cell exposure to desipramine causes a defect in resealing...... 108 Figure 3.9 siRNA knockdown efficiency ...... 110 Figure 3.10 Cell proliferation...... 113 Figure 3.11 Cell count ratios...... 114 Figure A.1 Rac1 FRET controls...... 148 Figure A.2 Hemolysis Assays...... 149 Figure A.3 Ca2+ influx triggers PKC activation at the plasma membrane...... 150 Figure A.4 LLO activates the Rho GTPase Cdc42...... 151

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Chapter 1: Introduction

The plasma membrane forms a biophysical barrier that separates the cell from its external environment. It is composed of a variety of lipids arranged in an asymmetrical bilayer and decorated with functionally and structurally diverse proteins. The multiple roles of the plasma membrane include preventing the free diffusion of ions and small molecules in and out of the cell, maintaining intracellular ionic and molecular contents distinct from the extracellular environment, and sensing and responding to environmental clues. Any perturbations that compromise the integrity of the plasma membrane can prove fatal for the cell in the absence of quick and efficient membrane repair processes that detect the breach in the plasma membrane and reseal it to re-establish its barrier function. It is thus understandable that plasma membrane integrity is one of the most critical aspects of cell viability and extensive research are dedicated to deciphering the molecular mechanisms that repair a damaged plasma membrane. Our understanding of membrane repair processes originated from observations that transected squid giant axons, microneedle injected sea urchin eggs and laser-ablated Xenopus laevis oocytes, could repair micrometer-sized lesions on the plasma membrane by the Ca2+-influx- dependent exocytosis of intraluminal vesicles, which facilitated resealing of the membrane and re-establishment of plasma membrane integrity 1-7. Since then, efforts to understand the molecular processes of plasma membrane repair have been motivated by 1 the devastating consequences associated with its dysfunctions, including Limb-Girdle muscular dystrophy, Duchene muscular dystrophy and Chediak-Higashi syndrome 8-11. In addition to understanding the underlying defaults in membrane repair associated with these genetic diseases, research in the field of infectious diseases have begun to illustrate how evolutionarily distinct pathogens damage target host plasma membranes in order to co-opt the subsequent cellular responses to promote their pathogenic lifestyles. In fact, much of our understanding of the molecular mechanisms involved in plasma membrane repair comes from studies using pathogen-derived pore-forming proteins or peptides, which create stable proteinaceous pores in membranes. Over the past 60 years of plasma membrane repair research, several mutually non-exclusive models of repair have emerged, however, none of which fully describes the intricate signaling network required for repair of a torn plasma membrane 12, 13. This introduction provides an overview of the history of plasma membrane repair research, the various experimental approaches that support our current understanding of cellular responses to plasma membrane damage, and the cellular responses to plasma membrane damage in the context of infectious diseases, with a special focus on listeriosis, a disease caused by the facultative intracellular pathogen Listeria monocytogenes.

1.1 Mammalian Plasma Membrane Biology

Since the first observation of a living cell was made in 1674 by Anton van

Leeuwekhoek, notably the pioneer of modern day microscopy, humanity’s intrigue of the cell has exploded into numerous topics including cell division, organelle biology, metabolism, and more. The cell as a singular functional entity is only made possible by 2 the barrier that separates it from the outside world, the plasma membrane. Structurally, the animal plasma membrane is a 2-dimensional semi-fluid bilayer composed of a variety of amphipathic glycerophospholipids (phosphatidylethanolamine, phosphatidylcholine, phosphatidylserine, phosphatidylinositol, phosphatidic acid, phosphatidylglycerol), sphingolipids (sphingomyelin and glycosphingolipids), along with other lipid species including sterols, triglycerides and glycolipids. Lipids are arranged in a bilayer with the large hydrophilic polar head groups facing the aqueous cytoplasmic and extracellular spaces, while the long hydrophobic fatty acyl chains face in towards the core of the bilayer. The hydrophilic head groups can be positively charged (ethanolamine), negatively charged (serine, phosphorylated inositol, phosphatidic acid), or neutral

(choline, glycerol, sphingomyelin, cardiolipin), and are asymmetrically distributed in the bilayer with the charged species mostly situated in the inner leaflet of the plasma membrane to regulate peripheral protein association, while the neutral phospholipids are located on the outer leaflet to limit the free flow of ions into the cell. For example, cytosolic peripheral proteins exhibit transient interactions with the plasma membrane through recognition of the charged phospholipids such as mono-, di-, or tri- phosphorylated phosphatidylinositol, within the inner leaflet of the membrane. The fluid nature of the plasma membrane is derived from the non-covalent interactions of the fatty acyl chains that can be completely saturated or exhibit varying degrees of unsaturation i.e. double bonds, in a cis- configuration. Long saturated fatty acyl chains of neighboring phospholipids can closely interact with each other via Van der Waals forces due to their linear structure thus resulting in lipid packing and decreased fluidity. Conversely, lipids

3 enriched in unsaturated acyl chains are more spaced out due to the kinked structure of the cis- configured fatty acyl chain resulting in loosely packed lipids and increased fluidity.

Under normal physiological conditions and temperature the fluid dynamics of the plasma membrane is characterized by a liquid ordered state in which phospholipids can freely move within the 2-dimensional plane. However, this lateral movement is heavily regulated by the presence of an essential component to all animal cells, cholesterol.

Cholesterol is a large sterol structure incorporated into the plasma membrane

(typically at a 1:1 ratio with phospholipids) and is critical to the structural and biophysical quality of the plasma membrane. The amphipathic sterol moiety is oriented with its hydroxyl group facing the polar heads of the neighboring lipids while the large hydrophobic ring structure interacts with the hydrophobic fatty acyl chains of the phospholipids. The presence of cholesterol in the membrane limits the free diffusion of small molecules into the cell by filling in the gaps created by the kinked structure of unsaturated fatty acyl tails, and modulates the fluidity of the membrane by acting as a bidirectional regulator. At normal physiological temperature (37˚C), the phospholipids and cholesterol exist in a liquid ordered state in which the fatty acyl chains are closely packed and exhibit dynamic lateral movement. A decrease in temperature causes slower lateral movements that are compounded by a cis-to-trans isomerization of unsaturated double bonds thereby increasing fatty acyl chain interactions and lipid compaction.

However, the presence of cholesterol acts as a spacer to interrupt lipid packing thus maintaining fluidity at temperatures above the melting temperature (Tm) of the membrane. Conversely, an increase in temperature decreases acyl chain interactions due 4 to increased Gibbs-free energy leading to highly sporadic lateral movement of lipids, termed the liquid-disordered state. Cholesterol acts to limit lateral movements by maintaining Van der Waals interactions between it and the neighboring fatty acyl chains thus maintaining a liquid ordered state. This unique property of cholesterol acts to buffer the effects of major swings in temperature thereby maintaining membrane fluidity. Aside from its structural and biophysical attributes, cholesterol is an essential component to the plasma membrane that facilitates protein integration and cellular signaling.

Figure 1.1 The plasma membrane Depicted are the chemical structures of 1-12:0-2-14:1Δ9-phosphatidylethanolamine (black) and cholesterol (orange). The plasma membrane is a dynamic 2-dimensional structural platform composed of various lipid species interspersed with glycolipids, integral proteins such as single pass or beta-barrel proteins, and associated peripheral proteins that bind to the charged phospholipids on the inner leaflet of the plasma membrane 14.

The foremost accepted description of the plasma membrane comes from SJ Singer and GL Nicolson’s fluid mosaic model, which depicts the plasma membrane as a 2- dimensional homogenous fluid lipid bilayer decorated with integral and peripheral

5 proteins that are free to move laterally within the plane of the membrane. Integral and peripheral proteins can transmit environmental cues into the cell, and reciprocally, promote cell-to-cell signaling through direct contacts or release of proteins, lipids, or nucleic acids. Measurements of lateral diffusion of model lipids with a cholesterol mole ratio of >0.3 exhibit diffusion coefficients of 3-5x10-8 cm2 s-1 whereas integral proteins of different sizes exhibit diffusion coefficients ranging from 8-20 μm2 s-1, indicating that the fluid dynamics of lipids surrounding proteins differ from that of free lipids, thus indicating that the membrane is not homogenous 15-17. Furthermore, developments in membrane biology have identified regions of plasma membrane that could not be solubilized in non-ionic detergents indicating a biophysically heterogeneous membrane

18. The composition of these detergent-insoluble microdomains known as lipid rafts is quite variable, but in general has been found to be highly enriched in cholesterol and saturated sphingomyelin, which, together, mediate a higher order of lipid packing while maintaining fluidity. These densely packed regions maintain a liquid ordered state, but exhibit more rigid-like characteristics, which promotes liquid-liquid phase partitioning.

The propensity for certain types of lipids to aggregate together suggested that these domains may act as functional platforms. Indeed, lipid rafts were found to be enriched in a subset of proteins including, but not limited to, glycosylphosphatidylinositol (GPI)- anchored proteins on the outer leaflet, acylated low molecular weight G proteins, and palmitoylated scaffolding proteins such as the mitogen-activated protein kinases (MAPK) on the inner leaflet. These findings proposed an amendment to the Singer Nicolson model that instead of the free-flowing homogenous sea of lipids and proteins, the plasma

6 membrane is a heterogeneous field of loosely packed lipids with biophysically distinct islands of densely packed lipid microdomains that exhibit spatially limited lateral movement due to characteristically distinct liquid phases that partition them from the open sea. These lipid rafts exist in a higher state of liquid-ordered relative to the more liquid-disordered state of the surrounding lipids, and act as biophysical platforms that facilitate the association, integration, movement and function of proteins.

The animal plasma membrane is a dynamic structure suited to sense environmental cues, send signals, and importantly exists as a biophysical barrier to maintain cellular homeostasis. Unlike the membrane structures of plant, fungal, and bacterial cell walls, which are composed of more rigid components (cellulose, chitin, peptidoglycan) that can resist physical perturbations to their membranes, the animal plasma membrane is fundamentally malleable. However, this pliability increases its susceptibility to mechanical, chemical, enzymatic, and osmotic damages. Therefore, animal cells have adapted to such challenges and developed potent mechanisms to reseal and repair their membrane.

1.2 Plasma membrane damage under physiological and pathological conditions

Under normal physiological conditions, muscular, cardiovascular, and intestinal tissues are subject to substantial mechanical and sheer stresses, especially during exercise

19-21. Active individuals reduce the risk of developing type II diabetes, cardiovascular disease, and obesity, however, immense forces are exerted on our muscular and cardiovascular systems during exercise leading to ruptures of the plasma membrane of

7 different tissues, including muscle fivers 22. Additionally, during intense exercise, the cardiovascular system undergoes hyperemia, i.e. increased blood flow to the active tissues, resulting in increased blood pressure and sheer stress on the inner lining of the vasculature 23. In healthy individuals, the aforementioned forces are mitigated by the production of nitric oxide, which induces vasodilation to increase vessel surface area thereby reducing the effect blood pressure 24. However, individuals with hypertension or atherosclerosis have limited capacity to compensate for increased blood pressure and sheer stress derived from hemodynamic forces, resulting in vascular endothelial rupture and disruption of vascular integrity 23. One perplexing observation that distinguished vascular endothelial cells to surrounding tissues was that cardiac endothelial cells are highly enriched in caveolae, flask-shaped membrane dense invaginations enriched with the membrane-bound proteins caveolin-1 and cavin. It was proposed that the high concentration of caveolae at the apical surface of vascular endothelial cells act by repairing damaged regions of the plasma membrane via endocytosis or act as shock and stretch absorbers that take the blunt of the sheer and mechanical forces during high cardio output. Indeed, it was shown that in the absence of caveolin, the continuous endothelial layer exhibits distended morphologies and the cells lining the vessels are highly prone to plasma membrane damage during increased hemodynamic stress 25. Plasma membrane repair also plays a major role in the context of certain genetic diseases such as Duchene- muscular dystrophy, in which mutations in the dystrophin gene lead to defective membrane resealing processes and atrophy of muscular tissue. Dystrophin is a stabilizing protein that connects the actin cytoskeleton to the plasma membrane of striated muscle

8 cells, termed the sarcolemma, and with the extracellular matrix 9. In the absence of this stabilizing protein and potential mechano-sensor, rigorous activity results in the destruction of the sarcolemma, which cannot be tolerated because the resealing mechanisms cannot keep up with the damage, causing necrosis, weakening of the tissue, and development of cardiomyopathies 26. Individuals with hypertension and artherosclerosis are also prone to cardiomyopathies because the increased systolic pressure and non-linear flow of blood can cause differential sheer stress on the vessels and thus compromise the endothelium 19, 23. In extreme events, a loss of blood supply to an organ due to an embolus, termed ischemia, can be detrimental to the downstream organ(s) due to a lack of continuous oxygen flow. Ischemia induces anaerobic respiration and a buildup of waste products resulting in decreased cytosolic pH, release of oxidizing compounds such as superoxide radicals and hydrogen peroxide that can degrade lipids within the plasma membrane, and loss of ionic homeostasis leading to cell necrosis 27.

Consequently, re-establishment of blood supply or reperfusion, induces sudden sheer stress to the already compromised tissue, and an inflammatory response compounds the level of damage by the invasion of leukocytes that release a myriad of proteases and lytic enzymes that further compromise the plasma membrane of surrounding tissues 28.

Additionally, the oxidizing extracellular environment is a continuous source of free- radicals that oxidize lipids in the plasma membrane thus compromising membrane integrity 29. All of these frequent sources of damage occur on a daily basis, and can be compounded during inflammatory and infectious conditions. The propensity of various tissues to become infected is in part due to pathogen-mediated perforation of target cell

9 plasma membranes as exemplified with pore-forming toxins 30. Inflammation of the infectious or non-infectious etiology, can also cause cell damage as dramatic physiological changes occur during inflammation including an increase in temperature, swelling of surrounding tissues, leukocyte extravasation and migration, and release of lytic enzymes such as perforin and the membrane attack complex 31, 32.

It is evident that plasma membrane resealing and repair is a fundamental process necessary to maintain healthy tissues in physiological and pathological conditions.

Understanding the intricate mechanisms involved in plasma membrane damage recognition and subsequent resealing/repair processes is thus paramount to resolving numerous diseases, providing preventative therapeutics for individuals susceptible to vessel or tissue failure, and facilitating repair processes in individuals recuperating from traumatic injuries.

1.3 Models to Study Plasma Membrane Resealing

Understanding how cells reseal their plasma membrane required the development of various experimental approaches that mimic physiologically relevant damage, biochemical assays to assess membrane damage and resealing, and biological systems in which to observe and quantify the subsequent resealing processes. At the beginning of the plasma membrane repair field, preliminary in vivo and ex vivo studies involved erythrocytes, squid giant axons, X. laevis oocytes, and sea urchin embryos exposed to hypotonic solutions, electroporation, transection, or microneedle puncture techniques to induce cell damage 2, 3, 33, 34. To read out the extent of damage, researchers quantified the 10 subsequent cell permeability to K+ and Na+ ions via flame photometry, tracked the spread of heavy isotope ions (45Ca2+) from the site of injury, or used fluorescent probes to measure the influx of Ca2+. These studies provided the fundamental basis that the plasma membrane can withstand large, seemingly insurmountable forms of damage of several micrometers in size. However, the capacity to study the repair processes required the development of more refined and sensitive approaches that could more precisely damage membranes, accurately quantify the resealing process, and visualize protein and vesicular transport in real time.

Many in vivo and ex vivo models have been developed to replicate the physiological forms of stress that result in plasma membrane rupture, such as induction of sheer and mechanical stress through scratch and bead abrasion assays or via contraction and stretch-based assay 35. The damaged cells can then be incubated in the presence or absence of extracellular Ca2+ in a cell culture medium containing membrane impermeant dyes such as propidium iodide, quickly washed, and analyzed by flow cytometry. A similar technique called syringe loading replicates the sheer stress that endothelial cells are exposed to during high cardio output. Simply, adherent or non-adherent cells are passed through a 30-gauge hypodermic needle multiple times in the presence of propidium iodide and analyzed via flow cytometry. Conditions in which cells exhibit high fluorescence intensities correspond to repair-defective conditions whereas low fluorescence intensities correspond to efficient resealing. Alternately, damaged cells release catalytically active cytosolic or lysosomal proteins such as lactate dehydrogenase

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(LDH) and β-hexosaminidase, which retain activity for hours after release. Assays can then test enzymatic activity as a read out for the degree of damage and repair.

A more consistent approach of membrane damage was developed as the scratch method or bead abrasion could be inconsistent in the degree of damage elicited. As such, adherent cells can be plated on a contractile or stretchable substrate and exposed to repetitive contraction and stretching conditions that challenge the integrity of the plasma membrane. The cells can then be labeled with fluorescent dextrans or membrane impermeable dyes for a given time period that permits resealing, and imaged using a fluorescence microscope. This approach provides a means to replicate physiological forms of damage and analyze the resulting repair efficiencies in a moderate throughput fashion. However, most of the described assays lack spatiotemporal resolution.

Advancements in microscopy have significantly improved the capacity to visualize the repair dynamics in real time. Individual cells expressing fluorescently tagged proteins or labeled with lipophilic dyes such as FM1-43 can be damaged via microneedle puncture and visualized using high-speed wide-field fluorescence or confocal microscopy. More recently, researchers have turned to the use of two-photon laser ablation techniques in coordination with high-speed laser scanning confocal microscopy to study membrane repair processes at the single cell level. In this technique, a region of the cell membrane can be ablated using pulsed lasers that form holes with distinct and consistent diameters.

The capacity to control the damage size was revolutionary as researchers could then study the effect of wound size on repair dynamics and mechanics in real time, and

12 directly visualize the events as compared to the indirect measurements performed on the mechanical, sheer, and contraction/stretch methods 36. The downside of laser ablation, however, is that it involves an extreme localized temperature increase that could denature surrounding repair-relevant proteins. This temperature increase also induces compressive thermoelastic stress due to an increase in volume of the irradiated area, which is not physiologically relevant 37.

One last method consists of forming holes in the plasma membrane using pathogen-derived pore-forming toxins. Pore-forming toxins compose the largest family of bacterial virulence factors that act by forming stable proteinaceous pores in membranes in order to disrupt cell integrity and promote infection 38. This strategy is also used by the immune system-derived pore-forming proteins such as perforin and the membrane attack complex, which are used to perforate infected cells and pathogens, respectively, as a means to clear the body of an infection 39-41. Researchers have heavily relied on the use of bacterial pore-forming toxins belonging to the family of cholesterol dependent cytolysins

(CDCs) including listeriolysin O from Listeria monocytogenes, streptolysin O from

Streptococcus pyogenes, and pneumolysin from Streptococcus pneumoniae. CDCs, such as listeriolysin O, are versatile tools to study different aspects of plasma membrane biology because they bind to cholesterol, which is present in high concentrations in all animal cells, oligomerize into pores of consistent diameter (about 30 nm), and are physiologically relevant in the context of infectious disease. CDCs exhibit the same ternary structure (Figure 1.2) with four domains: domains 1-3 are intertwined beta sheets

13 that transition into beta hairpins upon insertion into the plasma membrane to form the pore and domain 4, which binds to the host cell membrane via three hydrophobic loops, contains a threonine-leucine pair for cholesterol recognition, and a highly conserved undecapeptide sequence necessary for the conformational change coordinated with domain 3 upon pre-pore to pore conversion. Efforts to understand the structural properties of CDCs illustrated that they are secreted or released as monomers or dimers, which then bind to cholesterol in lipid membranes where they oligomerize into a pre-pore structure that undergoes a conformational change to form stable proteinaceous pores

(Figure 1.3). CDC variants trapped at different stages of pore formation have been developed to demonstrate the mechanism of oligomerization and final insertion to form the transmembrane pore.

Figure 1.2 Cholesterol dependent cytolysin crystal structures (Left to right) Crystal structures of Listeriolysin O, Streptolysin O, and Pneumolysin. This figure was adapted from (left to right) Köster et al., Feil et al., and Marshall et al.42- 44

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Among the CDCs, listeriolysin O (LLO) presents unique characteristics: an acidic triad (Glu-247, Asp-320, and Glu-208) within domain 3 promotes pH- and temperature- dependent irreversible denaturation due to protein unfurling at neutral pH and temperatures >30˚C 44-47. Additionally, LLO contains a unique N-terminal consisting of a polyproline type II helix that is involved in inter- and intramolecular interactions 44.

Lastly, the N-terminal contains a PEST-like sequence that was suggested to regulate LLO degradation within the host cell cytosol, however, it was found that wild type LLO and a

ΔPEST mutant exhibit similar half-lives 46. It was recently discovered that the PEST-like sequence is recognized by the mammalian adaptor protein 2 (AP2), which acts to promote clathrin-mediated endocytosis of LLO that is secreted from within the host cell cytosol thus limiting host membrane perforation 45.

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Figure 1.3 Listeriolysin O crystal structure (Top) The listeriolysin O crystal structure as a monomer, a linear array, and a pore consisting of 36 monomers. Domains 1-4 are color coated red, yellow, green, and blue respectively. The N-terminal region colored purple contains a polyprolene type II helix, and a PEST-like sequence unique to LLO that is not conserved across other CDC pore- forming toxins. (Middle) A model of CDC insertion into a cholesterol-rich membrane based on the structure of PFO. Monomers (red and yellow) concentrate at the surface by binding to cholesterol (blue) and oligomerize into an arc formation. Interaction of the leucine-threonine pair with cholesterol initiates a conformational change in domains 1-3 leading to the formation of an alpha-helical “dagger” (purple) that partitions the surrounding membrane for the insertion of the PFT. (Bottom) is a transmission electron micrograph of LLO in monomer form, arcs, pre-pore, and pore arrangements. Scale bar = 50 nm. This figure was adapted from (top, middle, bottom) Köster et al., Rossjohn et al., and Vadia et al. 44, 48, 49

CDCs have been instrumental in the identification of lysosome exocytosis as a repair mechanism, characterization of mitogen activated protein kinases (MAPK) signaling in

16 response to K+ efflux, and recognition that pore-forming toxins can mediated the internalization of evolutionarily distinct pathogens 48, 50-55. In addition, pore forming toxins that create smaller pore sizes (1-2 nm in diameter) have been used to study the effect of pore size on cellular signaling and membrane resealing. Studies using the PFTs

Aerolysin from Aeromonas hydrophila (1.5-2 nm diameter), α-toxin from Staphylococcus aureus (1-2 nm diameter), and Cry5B from Bacillus thuringiensis (2 nm diameter) have been critical in understanding the role of K+ in mitogen-activated protein kinase (MAPK) signaling and subsequent transcriptional changes using human cell lines and animal models including Caenorhabditis elegans and Drosophila melanogaster 54, 56, 57.

Combined together, each model of damage has been influential in the identification of various mechanisms of plasma membrane repair.

1.4 Plasma Membrane Repair Models

Different experimental cellular and damage models were used leading to several mechanistic events, some of which may be universal, while others are potentially cell- and/or damage-specific. At this time there is no available unifying repair model, nor cell- type or damage-dependent repair model. The discovery of plasma membrane repair mechanisms originated from preliminary studies using squid giant axons and X. laevis oocytes, which revealed that extracellular Ca2+ influx through the plasma membrane breach results in intraluminal vesicle exocytosis, which facilitates resealing 1, 2. These findings, which were corroborated using other animal models including starfish oocytes and sea urchin eggs 4, not only identified Ca2+ influx as a universally conserved signal of a plasma membrane breach, but initiated an entire field dedicated to understanding the 17 mechanisms involved in plasma membrane repair downstream of the influx of extracellular Ca2+.

Vesicle exocytosis and Spontaneous Fusion

The preliminary studies of membrane repair utilized oocytes, which contain a population of cortical secretory granules that are primed for exocytosis upon fertilization- mediated damage, but this left the question of whether or not this mechanism of repair is similar in somatic cells. Upon damage to the plasma membrane, the hydrophobic acyl chains of the phospholipids at the open edge are exposed to the aqueous environment, causing a thermodynamically-driven repositioning of the phospholipids to create a rounded-off ledge. Studies using liposomes and erythrocytes illustrated how red blood cells permeabilized by incubation in hypotonic solution could spontaneously reseal. The model proposed that the increased lipid disorder at the edge of the wound drives the edges of the breach towards each other resulting in spontaneous fusion. However the conditions in which this event could experimentally occur (medium containing 1/10th the physiological extracellular concentration of ions, T = 0˚C) were not physiologically relevant 58-62. This model, feasible in cells lacking an underlying cytoskeletal structure and damaged with perforations in the nanometer range, fails to explain how nucleated cells combat tension forces produced by the cytoskeleton and in physiologically relevant damaging conditions in which micrometer-wide lesions are repaired. An alternative mechanism had to be at play.

The Patch Hypothesis

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Fusion of intracellular vesicles prior to resealing were observed in the X. laevis oocyte and sea urchin egg model, which incited an idea that came to be known as the patch hypothesis 63. It was proposed that the intracellular granules could fuse together to create a large vesicular body that would then fuse with the plasma membrane surrounding the breach creating a patch. However, the mechanisms by which these vesicles could directly translocate to and fuse with the damaged region were unknown. Studies using microneedle injection, 2-photon laser ablation and confocal/spinning disc microscopy, have been able to visualize the depolymerization of subcortical actin by action of calpains, recruitment of the fast-acting Ca2+ sensors annexins, S100A11, and dystrophin to the damaged region to stabilize the damaged membrane, and MG53/TRIM72 and dysferlin form a temporary protein mesh signaling network to designate the region for vesicle fusion mediated by synaptotagmins 64-66. Calpains are Ca2+-activated cysteine proteases that function to cleave the subcortical cytoskeleton in order to dissociate the damaged region of the plasma membrane from the underlying structural components of the cell thereby relieving membrane tension and opening a direct line for vesicle exocytosis 65, 67. Consequently, the surrounding actin cytoskeleton forms a contractile ring around the perimeter of the wound in order to limit wound extension by coordination with annexins and promote wound closure 68. The family of annexins consist of 12 isoforms (A1-A11, A13) that bind to phospholipids in a Ca2+-dependent manner to stabilize the plasma membrane and promote endo- and exocytic events 69. Annexins A1,

A2, and A6 and the Ca2+ binding protein S100A11 were found to be recruited to the plasma membrane in order to stabilize the damaged plasma membrane with the

19 surrounding actin cytoskeleton at the wound perimeter and mediate vesicle fusion events to promote resealing. 64, 68, 70, 71. Dystrophin, MG53/TRIM72, and dysferlin are muscle- specific proteins involved in the stabilization of the sarcolemma (plasma membrane of striated muscle cells) with the extracellular matrix and organization of a membrane repair complex that facilitates vesicle docking and fusion 72. All of these proteins function at the site of plasma membrane damage to stabilize the region, but the “patching” is finalized by the vesicular fusion events juxtaposed to the breach73. Upon influx of extracellular

Ca2+, vesicles are quickly transported to the membrane damage via the actin and microtubule motors myosin II and kinesin, respectively, and on their way frequently fuse together and with the plasma membrane via Rab3a, Rab10, and synaptotagmin VII, a

Ca2+-sensitive integral protein that binds to soluble N-ethylmaleimide-sensitive factor attachment protein receptors (SNAREs) on adjacent membranes promoting membrane fusion events 74-76. In accordance with these findings, using high speed confocal microscopy, it was evidenced that vesicles quickly exocytose after membrane perforation, fuse together to form a patch and upon fusion with the damaged region, frequently rupture outwards upon proximity to the breach producing a single bilayer that can fuse with the open edge of the breach and ultimately re-establishing a continuous bilayer.

Lastly, the recruitment of annexin A1, dysferlin, and activated Rho GTPases RhoA and

Cdc42 form concentric circles around the breach to facilitate constriction of the actin contractile ring around the damaged region during the final stages of patching. This patching mechanism was evidenced in X. laevis oocytes because the large cell provided the capacity to perform z-plane sectioning that could discern these events. However,

20 animal somatic cells such as endothelial and epithelial cells are not primed with secretory granules and the cytosolic space is spatially confined relative to the oocyte (oocyte volume is 6 orders of magnitude greater than that of human cells), which leaves the question of whether the patch hypothesis is a generalized mechanism or only seen in oocytes.

Lysosomal exocytosis followed by endocytosis

It was believed that only certain types of secretory cells exhibit regulated vesicular exocytosis to promote signaling (neurotransmitters, hormones) 77, 78. However, further studies revealed that many more cell types exhibit regulated exocytosis, especially in response to plasma membrane damage induced Ca2+ influx in which vesicles exocytose to promote membrane resealing 79, 80. Researchers were thus curious as to the nature and identity of the vesicle population. Using a combination of immunofluorescence labelling of the luminal domains of proteins found in specific intracellular compartments and chemical inhibition of vesicular trafficking, lysosomes were identified as the population of vesicles that exocytose upon membrane perforation 51, 52, 66, 81, 82. Further studies using the pore-forming toxin streptolysin O (SLO) to damage the plasma membrane of mammalian cells revealed that Ca2+-influx through the proteinaceous pores induced the exocytosis of lysosomes, and release of the lysosomal hydrolase acid sphingomyelinase

(ASM) into the extracellular milieu 51, 52, 66, 81, 83. A recent finding illustrated that lysosomal positioning and exocytosis is dependent on Rab3A, a membrane-associated protein involved in vesicular trafficking during exocytosis, in coordination with the non- muscle myosin heavy chain IIA, an actin motor that transports cargo along the actin 21 cytoskeleton 83. Upon lysosomal exocytosis, extracellular ASM hydrolyzes outer leaflet sphingomyelin into phosphocholine and ceramide, reducing the surface area of the outer leaflet of the plasma membrane, which is accommodated by the subsequent invagination of the membrane. These invaginated regions are then populated by the membrane-bound caveolin, which act as scaffolding proteins to localize signaling molecules and drive the endocytosis of SLO pores 51, 81, 84, 85. Follow up studies illustrated that endocytosed SLO is then trafficked to multivesicular bodies/lysosomes via the endosomal sorting complex required for transport (ESCRT) machinery and degraded 84. The ASM-dependent process was also proposed to repair large mechanical membrane lesions produced by glass bead abrasion and photon-ablation indicating that this mechanism of repair is not exclusive to the repair of the plasma membrane perforated by stable proteinaceous pores 86.

Ectocytosis/Microvesicle shedding

Although the ASM-mediated pathway can explain how proteinaceous CDC pores are removed from the plasma membrane, it was also observed that PFT-damaged cells exhibited outward-facing bleb morphologies and microvesicle shedding, indicating a potentially distinct mechanism of repair that did not exclusively involve lysosomal- mediated endocytosis 87. Studies using monomer, oligomeric, and pore-competent SLO, perfringolysin (PFO), and intermedilysin (ILY), all of which belong to the CDC family and characteristically bind to membranes via cholesterol (SLO and PFO) or other receptors such as CD59 in the example of ILY, showed that various cell types including adrenal (Hek293), epithelial (HeLa), fibroblast (3T3), monocyte (THP1), and bone marrow derived macrophages, shed oligomeric and pore-competent PFTs while the 22 monomeric forms are internalized and degraded 88. Additionally, a study using the unrelated PFT α-toxin from Staphylococcous aureus showed that although both monomeric (1mer) and pore-competent α-toxin (7mer) are endocytosed and trafficked to late endosomes, only 1mers were degraded whereas 7mers were further trafficked via multivesicular bodies back out of the cell in exosomes, termed toxosomes 89. This mechanism of toxin release was cell type dependent as HaCaT keratinocytes and Cos7 fibroblasts readily endocytosed α-toxin and retained cell viability whereas Huh7 hepatic epithelial cells were incapable of endocytosing α-toxin and succumbed to necrosis. There is no clear answer as to which mechanism is at play, or if both are at play, and why different cell types respond differently. The various observations made may be due to differences in the location, degree, and extent of damage, and the cell type, which was considered in the observation that HEK293 adrenal cells and SH-SY5Y neuroblastoma cells exhibit bleb morphology and microvesicle shedding in regions distal to the cellular body 87. Conversely, inhibition of lysosomal exocytosis resulted in the lysis of SH-SY5Y cells, but not HEK293, likely due to cell-specific tolerances of self-inflicted wounding during membrane contraction in response to SLO-treatment 87. This lays claim that microvesicle shedding and lysosomal exocytosis play different roles in repair, the former promoting the release of mature pores whereas the latter facilitating the repair of subsequent self-inflicted damage. Follow up studies of microvesicle shedding aimed to identify the mechanism whereby the membrane blebs are released. The endosome sorting complex required for transport (ESCRT) machinery has been suggested to promote the shedding of damaged regions of the membrane through exocytosis of intraluminal

23 vesicles within multivesicular bodies and/or direct release of microvesicles containing intact pores. As such, HeLa cells damaged via detergents (Saponin, Digitonin), pore- forming toxins (LLO), sheer stress, and laser ablation exhibited the recruitment of

ESCRT machinery to the site of injury, which promoted outward blebbing of the plasma membrane and shedding of blebs 90.

It is clear that the mechanisms of plasma membrane repair may follow different paths (Figure 1.4) depending on cell type, tolerance to damage, and form of damage. To add, the use of different cell models and damage have led to the description of several mutually non-exclusive models of repair, some of which occur under non-physiological conditions (spontaneous fusion) and some that seem to be a generalized mechanism

(endocytosis, exocytosis and shedding of vesicles), but mechanistically play different roles depending on the model system and form of damage.

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Figure 1.4 Plasma membrane repair models A breach in the plasma membrane results in the influx of extracellular Ca2+ causing the exocytosis of vesicles around the breach to reduce the membrane line tension produced from the underlying actin cytoskeleton. Conversely, vesicle-vesicle fusion forms a “patch” that fuses with the membrane boundary to seal the breach. Pore formation on the plasma membrane leads to the influx of Ca2+ and the subsequent exocytosis of lysosomes releasing acid sphingomyelinase into the extracellular milieu. Conversion of sphingomyelin to ceramide results in ceramide-driven invagination and caveolin- mediated endocytosis of the mature pore. The pore-containing vesicle is then shuttled to multivesicular bodies (MVBs) via ESCRT machinery and are either degraded in late endosomes/lysosomes or released as exosomes. Conversely, pores are explicitly shuttled into membrane blebs that are shed from the cell body as microvesicles via ESCRT machinery, in a process termed ectocytosis, while remaining pore forming toxin monomers enter the ASM-mediated endocytic pathway where they are shuttled into MVBs for degradation. Note that not all proteins identified in plasma membrane repair, including Rab proteins, Myosin/Kinesin, and the exocyst complex are depicted.

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The above figure illustrates the various models of repair, each brought to light by the use of distinct cell models (oocytes, myocytes, epithelial, endothelial, neuronal cells) and in vitro techniques to mimic physiologically damages (bead abrasion, microneedle injection, laser ablation, and pore-forming toxin). It is with these techniques that we now have an understanding of the genetic diseases associated with defective plasma membrane repair processes such as Limb girdle muscular dystrophy, Duchene muscular dystrophy, and

Miyoshi myopathy, and it is the hope that these models will pave the way for the development of therapeutics that can resolve such devastating diseases. Lastly, the role of plasma membrane repair has extended into the field of infectious disease as efforts to understand repair mechanism with the use of pore-forming toxins initiated a new perspective of plasma membrane damage and repair in the context of host-pathogen interactions.

1.5 Signaling and repair mechanisms in response to plasma membrane damage in the context of host-pathogen interactions

In the field of infectious disease, it is widely known that evolutionarily distinct pathogens utilize a multitude of virulence factors to damage host cell membranes in order to perpetuate their infectious lifecycles 91-103. Such mechanisms include secretion of phospholipases and pore-forming proteins that target with more or less specificity the plasma membrane of host cells. The subsequent cellular responses vary depending on the type and degree of damage. Numerous pathways are activated when the plasma membrane is disrupted including signaling pathways for membrane resealing and

26 additional pathways including the induction of the pro-inflammatory signaling via mitogen-activated protein kinase (MAPK) signaling, activation of the inflammasome and caspase-1, transcriptional reprogramming, translational arrest, and metabolic alterations.

These types of responses only occur in conditions in which the damaged cells have the capacity to repair their membranes. This has sparked interest in how the plasma membrane is repaired and whether the repair machinery on its own and/or repair- independent signaling pathways can facilitate an infection. For example, the eukaryotic parasite Trypanosoma cruzi is proposed to mechanically injure the host cell plasma membrane via flagellar motility while the human adenovirus HAdV-C2 perforates the host plasma membrane via its pore-forming peptide protein VI. These mechanisms of damage were proposed to co-opt the ASM-mediated membrane repair pathway for the efficient internalization of the pathogen into the target cell 50, 53. Additionally, due to the complexity of the human microbiome, pathogen-mediated host plasma membrane damage could promote secondary infections. Normal human keratinocytes exposed to α- toxin expressing Staphylococcus aureus or exposure to sublytic doses of the pore- forming toxin alone were more efficiently infected with Vaccinia virus and Herpes simplex virus type I, both of which exhibit endocytosis-mediated entry and require cholesterol/sphingomyelin domains for their entry process 104-107. Lastly, it was previously recognized that extracellular Listeria monocytogenes could damage target host plasma membranes via its pore-forming toxin listeriolysin O causing Ca2+-influx and ultimately dynamin- and actin-dependent internalization of the pathogen 55, 108. It is thus

27 conceivable that pathogen-induced host plasma membrane damage and the subsequent signaling events provide a mechanism for pathogen entry into target host cells.

1.6 Intracellular Infectious cycle of Listeria monocytogenes

The Gram-positive bacterium Listeria monocytogenes has long been used as a model facultative intracellular pathogen. Investigations of the intracellular lifecycle of L. monocytogenes in professional phagocytes and cells that are normally non-phagocytic provided significant findings in cellular biology as well as the innate and adaptive immune responses. Upon ingestion of contaminated food, L. monocytogenes encounters a sudden environmental shift including a rise in temperature and osmolarity, low pH from stomach fluid, and exposure to immune defenses such as alpha-defensins, to name a few

109. All of these environmental conditions act as cues that transition L. monocytogenes to a pathogenic bacterium. Most notably, a rise in temperature to 37˚C results in the transcription and translation of the virulence transcription factor PrfA, which regulates the expression of key virulence factors including the surface adhesins internalin A (InlA) and InlB, the pore-forming toxin listeriolysin O (LLO), two phospholipases (PI-PLC and

PC-PLC), and the actin nucleating factor ActA 110, 111. The covalently attached InlA and electrostatically bound InlB act as ligands for the mammalian receptor E-cadherin and c-

MET/HGFR, respectively, which promote receptor-mediated endocytosis of the bacterium into endosomes 112-115. Upon entry into an early endosome, L. monocytogenes secretes its pH- and temperature sensitive pore-forming toxin (PFT) LLO to perforate and break open the acidifying endosomal compartment resulting in the release of the bacterium into the cytosol where it utilizes polarly-situated ActA to recruit and nucleate 28 host actin to promote its intracellular motility 116-119. This motility frequently results in

Listeria-containing membrane protrusions that push into neighboring cells resulting in the spread and encapsulation of the bacterium into a double-membrane vacuole 120, 121. The two phospholipases, in conjunction with LLO, act to degrade and perforate the double membrane vacuole and release the bacterium into the cytosol thus perpetuating its intracellular lifecycle (Figure 1.5)

Figure 1.5 The Listeria monocytogenes intracellular lifecycle 122 Internalins InlA and InlB, listeriolysin O (LLO), actin assembly-inducing protein (ActA), phosphatidylinositide-specific and phosphatidylcholine-specific phospholipases (PlcA and PlcB).

It was previously thought that LLO activity was restricted to the acidifying endosomal compartment where it acts to release encapsulated L. monocytogenes into the host cell cytosol, and that cytosolic LLO is quickly denatured and degraded by the proteasome 46, 123, 124. Though its structure is conserved amongst all of the CDC PFTs,

LLO exhibits some unique qualities including an N-terminal PEST-like sequence that regulates cytosolic LLO activity, an acidic triad that optimizes PFT activity at low pH,

29 and temperature sensitivity that causes the loss of pore-forming activity due to unfolding and aggregation after incubation at 37˚C and neutral pH 47, 125. It was believed that these attributes strictly limit the formation of LLO pores to acidified endosomal/phagosomal compartments, thus preventing unintended damage to the host cell during its intracellular lifecycle. However, further studies found that both extracellular and cytosolic L. monocytogenes continuously secretes LLO that perforates the host plasma membrane resulting in various host cell responses such as histone modification, mitochondrial fragmentation, alterations of post-translational modifications (SUMOylation), induction of pro-inflammatory responses, cellular senescence, and disruption of immune cell functions 126-132. Additionally, recent findings from our lab suggest that the L. monocytogenes adhesins InlA acts to bind L. monocytogenes to the host plasma membrane where localized secretion of LLO permits controlled, sublytic perforation of the host plasma membrane leading to the endocytosis of L. monocytogenes 133. This brought the question of whether or not the LLO-mediated entry of L. monocytogenes also follows the ASM-dependent plasma membrane resealing process that facilitates the internalization of other pathogens. In addition, it was recently discovered that the N- terminal PEST-like sequence binds to the adaptor protein AP-2 to promote clathrin- mediated endocytosis of membrane bound LLO as a means of membrane repair 45.

Together, these findings warranted the following research project to dissect the mechanism whereby extracellularly secreted LLO modulates host cell signaling for the efficient internalization of L. monocytogenes and to determine if this entry pathway is equivalent to the ASM-mediated entry pathway utilized by T. cruzi and Adenovirus.

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Moreover, we would like to elucidate the mechanisms involved in repair of mammalian cells wounded by LLO and their relation to the AP-2 induced membrane endocytosis.

1.7 Implications and Perspective

The multiple mutually non-exclusive models of repair have provided the template of the intricate signaling processes involved in plasma membrane resealing and repair

(Figure 1.4), but there is still much more to be discovered. Though these models have been partially realized through various experimental approaches and model systems, to date there is no unifying pathway that completely describes step-by-step how a cell recognizes various forms (mechanical or pore-forming protein) and sizes of plasma membrane damage, what protein(s) are the direct sensor of cytosolic Ca2+ that initiate repair signaling, the step-by-step protein-protein interactions that are implemented to reseal the damaged membrane, and if resealing processes exhibit hierarchal implementation or are cell type specific.

Since the identification of dystrophin as the culprit behind Duchene muscular dystrophy (DMD), various approaches have been taken to reduce the symptoms associated with this disease. Just recently a group was able to introduce mini-dystrophin in a Duchene muscular dystrophy mouse model via viral gene therapy 134, which showed enough promising results for Pfizer to initiate a stage 1b human clinical trial with hopes of reducing the symptoms of the disease. Findings in the field of membrane repair stretch even further to cancer biology in which it was found that cancer cell metastasis requires

Annexin A2 and S100A11-mediated plasma membrane repair, which represents potential 31 targets to limit tumorigenesis 71. Further identification and characterization of proteins involved in plasma membrane resealing could open the doors to a myriad of targets that can be pharmacologically targeted to improve the welfare of individuals suffering from diseases associated with plasma membrane repair deficits such as muscular myopathies, diabetes and complications due to ischemia-reperfusion injury. The following research goals were 1) to characterize the cellular signaling in response to plasma membrane damage in the context of infection by L. monocytogenes and to establish the contribution of membrane resealing in L. monocytogenes internalization, and 2) to identify novel proteins involved in plasma membrane repair in order to complete our understanding of the plasma membrane resealing processes, with the hope of unifying the various repair models and providing a complete pathway from which to base new research and development.

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2 Chapter 2. Plasma Membrane Damage, A Novel Mechanism of Pathogen Entry

2.1 Introduction

Intracellular pathogens use a large repertoire of virulence factors to subvert host cell machineries, thereby ensuring their life cycle and propagation within the infected host. An initial and indispensable step is the internalization of the pathogen into host cells 135, 136.

The present study elucidated a signaling pathway underlying a unique mechanism of pathogen uptake by host cells, in which pathogens damage the host cell plasma membrane to promote their internalization.

The process of pathogen internalization into host cells can be passive from the perspective of the pathogen when a professional phagocyte uses its phagocytic receptors to engulf the pathogen 137-139. Alternatively, the uptake process can be active when pathogens express virulence factors that hijack the host cell endocytic pathways 136, 140-142.

Although there is a high diversity of pathogens, they evolved common strategies to actively promote their internalization into host cells. Some pathogens use surface invasins that specifically activate a host receptor-mediated endocytic pathway 142-145. Other pathogens use a secretion apparatus that injects effectors into the host cell cytosol, which directly subvert signaling pathways that orchestrate the endocytic uptake of the pathogen 137, 146-148.

Recent studies propose an additional invasion strategy that involves host cell plasma membrane perforation. Indeed, unrelated pathogens such as the eukaryotic parasite Trypanosoma cruzi, the bacterial pathogen Listeria monocytogenes, and the adenovirus HAdV-C2 perforate the host cell plasma membrane to promote their internalization into non-phagocytic cells 48, 50, 53, 55. Some pathogens express several 33 invasion factors and use multiple mechanisms to promote their internalization.

Collectively, cooperation between these factors may increase the efficiency of host cell invasion. For example, to enter host cells Salmonella uses the surface invasin Rck and injects effectors via a type III secretion machinery (T3SS) 148-150. Adenoviruses can be internalized into host cells as a result of host receptor-mediated endocytosis and/or of plasma membrane injury 50, 151, 152.

A particularity of the Gram-positive foodborne pathogen L. monocytogenes is its ability to infect a large diversity of cells including cells that are normally non-phagocytic such as enterocytes, hepatocytes, cytotrophoblasts, and neurons 111. Therefore, it is not surprising that this pathogen uses multiple invasins and invasion strategies. L. monocytogenes expresses the surface invasins InlA and InlB to promote its internalization into cells that express the internalins receptors, E-cadherin and c-Met, respectively 112, 145.

Additionally, the secreted pore-forming toxin listeriolysin O (LLO) perforates the host cell plasma membrane to promote L. monocytogenes internalization into epithelial cells 48, 55.

LLO is a member of the MACPF/CDC (membrane attack complex perforin/cholesterol- dependent cytolysins) superfamily, which includes numerous eukaryotic pore-forming proteins and the largest family of bacterial pore-forming toxins, the CDCs 41, 118. CDCs form large β-barrel pores across cholesterol-containing membranes and are major virulence factors produced by Gram-positive pathogens. The CDC LLO and intermedilysin O (ILO), which is secreted by Streptococcus intermedius, induce bacterial internalization into human hepatocytes 48, 153.

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Although numerous pathways for host receptor- and T3SS effector-mediated pathogen internalization have been extensively studied, less is known about the molecular machineries that link plasma membrane perforation to pathogen internalization. Recent studies have proposed that the pathway involved in resealing the damaged plasma membrane is directly responsible for pathogen uptake 50, 53, 154. Animal cells are frequently wounded due to numerous physical and biological stressors. For example, cells that compose contractile tissues, including those present in skeletal muscles or the intestinal tissue, are subject to mechanical stress. Cells throughout the body may also be exposed to pore-forming toxins produced by pathogens or by membrane-damaging agents released by the immune system. Owing to the frequency of these traumatic events and the inherent fragility of lipid bilayers, animal cells are able to rapidly reseal their plasma membrane to avoid necrosis or programmed cell death pathways. Several resealing mechanisms have been proposed including endocytosis of the damaged membrane 51, 155. It was proposed that the endocytic machinery that promotes membrane resealing is hijacked by pathogens such as T. cruzi and adenovirus to gain entry into host cells 50, 53.

The present studies were designed to 1) identify key signaling events activated downstream from LLO-induced host cell plasma membrane perforation that mediate L. monocytogenes internalization and 2) delineate the potential involvement of membrane resealing in this signaling pathway. Together, our studies identified a novel endocytic pathway of L. monocytogenes and support a model for damage-dependent pathogen uptake that is independent of membrane resealing.

Abbreviations 35

ASM – acid sphingomyelinase CDC – cholesterol-dependent cytolysin cPKC – conventional protein kinase C FRET – fluorescence resonance energy transfer LLO – listeriolysin O LLOpL – listeriolysin O prepore lock mCFP – monomeric cyan fluorescent protein mCit – monomeric citrine MyrPalm-CKAR – myristoylated and palmitoylated C kinase activated reporter NC – negative control PBD – p21 activated kinase binding domain PI – propidium iodide

For all movies refer to https://www.molbiolcell.org/doi/10.1091/mbc.e17-09-0561

2.2 Results

Rac1 is required for LLO-mediated internalization of L. monocytogenes We previously showed that LLO pore formation was sufficient to induce F-actin– dependent L. monocytogenes internalization into epithelial cells 48. Because the Rho

GTPases Rac1, Cdc42, and RhoA are master regulators of F-actin remodeling during bacterial internalization, we first established whether they were involved in the LLO- mediated entry pathway 156-158. Expression of Rac1, RhoA, and Cdc42 was knocked down in human hepatocytes (HepG2) via treatment with silencing RNAs (siRNAs) for 48 h. The resulting knockdown efficiencies were assessed in each experiment by Western blot analysis of Rac1, RhoA, or Cdc42 in nontreated and siRNA-treated cells (Figure 2.1A).

For each experiment, Rac1 protein level was decreased by 50–85% relative to negative control cells (NC) treated with NC siRNA, and RhoA and Cdc42 protein levels were decreased by more than 87% relative to NC cells (Figure 2.1A). Additionally, because these Rho GTPases can have overlapping activities, such as Rac1 and Cdc42 in the formation of lamellipodia 159, we also verified that silencing of one Rho GTPase did not

36 result in a compensatory overexpression of the other two (Figure 2.1B). Densitometry analysis of the Western blots indicated that cell treatment with Rho GTPase-specific siRNAs does not result in compensatory expression of the two other Rho GTPases when compared with cells treated with NC siRNA (Table A.1).

Figure 2.1: Rac1, RhoA, and Cdc42 knockdown efficiencies (A and B) Western blot analysis of Rac1, RhoA, and Cdc42 in nontreated HepG2 cells and in HepG2 cells treated with a negative control siRNA (NC) or with Rac1-, RhoA-, or Cdc42-specific siRNAs. (A) Western blot analysis of cell lysates (nondiluted, 1:2, 1:4, and 1:8 diluted) using anti-Rac1, -RhoA, -Cdc42, and –actin antibodies. (B) Western blot analysis of nondiluted cell lysates to verify the absence of compensatory expression of Rac1 (top), RhoA (middle), or Cdc42 (bottom) in cells treated with Rac1-, RhoA-, and Cdc42-siRNAs. All Western blots are representative of at least three independent experiments.

To determine the importance of these Rho GTPases for LLO-dependent internalization, nontreated and siRNA-treated cells were incubated with wild-type L. monocytogenes (WT, 10403s) or its isogenic LLO-deficient mutant (Δhly) for 30 min at 37

37˚C, and bacterial internalization (entry) efficiency was measured by immunofluorescence microscopy 160. In all treatment conditions, entry efficiencies of the

LLO-deficient bacteria were less than 14% relative to entry of WT bacteria in NC siRNA- treated cells, further verifying that L. monocytogenes internalization is LLO dependent in

HepG2 cells 48. Also, there was no significant difference in bacterial entry between the nontreated and NC siRNA-treated cells, showing that cell transfection did not alter bacterial uptake (Figure 2.2A).

Figure 2.2: Role of Rac1 in LLO-dependent entry of L. monocytogenes. (A and B) Nontreated HepG2 cells and HepG2 cells treated with negative control (NC), Rac1-, RhoA-, and Cdc42-siRNAs were incubated with WT (10403s) or the LLO deficient (Δhly) L. monocytogenes (A), or with BSA or BSA/LLO-coated beads (B), at a multiplicity of infection 20 (MOI 20) for 30 min at 37˚C. (C) HepG2 cells expressing mCit-Rac1 or dominant negative mCit-Rac1N17 were incubated with BSA/LLO-coated beads at MOI 5 for 30 min at 37˚C. Cells were then fixed and bacteria or beads were fluorescently labeled to enumerate the total number of bacteria (Nt) and the number of extracellular bacteria associated with host cells (Ne) (A, B, C). Entry efficiency was measured as 100 × [(Nt)-(Ne)]/(Nt) and results are expressed as the percentage entry ± SEM relative to NC-siRNA-treated cells incubated with WT (A) or BSA/LLO-coated polystyrene beads (B) (n ≥ 3). In C, results are expressed as the percentage entry ± SEM relative to mCit-Rac1 expressing cells incubated with BSA/LLO-coated beads (n = 4). Statistics are as follows: *p < 0.05; **p < 0.01; ***p < 0.005.

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Knocking down Rac1 reduced entry of WT bacteria by 51.3 ± 3.2% (p < 0.01), whereas knocking down RhoA or Cdc42 did not significantly affect entry (Figure 2.2A). To further demonstrate the role of Rac1 in the LLO-mediated entry pathway in the absence of any other virulence factors, we measured the entry of 1-µm polystyrene beads coated with purified recombinant LLO. Beads were covalently coated with bovine serum albumin

(BSA) followed by a noncovalent adsorption of LLO to mimic the release of LLO by bacteria 48. BSA-coated beads were used as a negative control. Nontreated and cells treated with NC-, Rac1-, RhoA-, and Cdc42-siRNAs were incubated with BSA- or BSA/LLO- coated beads for 30 min at 37˚C and bead entry efficiency was measured by immunofluorescence microscopy. Relative to the NC siRNA treatment, Rac1-siRNA- treated cells exhibited a 61.8 ± 0.1% decrease in LLO-coated bead entry (p < 0.005). There was no statistically significant difference between RhoA and Cdc42 knockdown conditions relative to the NC siRNA-treated cells (Figure 2.2B).

On guanosine nucleotide exchange, the Rho GTPases undergo a conformational change into an active GTP-bound state, which permits their reversible association with downstream effectors such as p21 activated kinases (PAK) 161. We assessed whether Rac1 activation was required for the LLO-mediated entry pathway by utilizing a Rac1 dominant negative variant (Rac1N17) that is unable to bind GTP. HepG2 cells were transfected to express the fluorescent chimeras (monomeric Citrine) of native Rac1 (mCit-Rac1) or the dominant negative Rac1 variant (mCit-Rac1N17) 162. Entry of BSA/LLO-coated beads into transfected, fluorescent cells expressing mCit-Rac1N17 was 74.6 ± 0.1% lower than cells 39 expressing the native form of Rac1 (p < 0.05) (Figure 2.2C). Collectively, these data show that Rac1, but not RhoA or Cdc42, plays a crucial role in the LLO-mediated entry pathway of L. monocytogenes.

Rac1 activation by LLO is pore and Ca2+ influx dependent

To confirm that LLO is sufficient to activate Rac1, we monitored the spatiotemporal dynamics of Rac1 activation in living cells using a fluorescence resonance energy transfer– (FRET) based fluorescence microscopy method 162, 163. HepG2 cells cotransfected to express the FRET-pair mCit-Rac1 and monomeric cyan fluorescent protein–p21binding domain (mCFP-PBD) were imaged in an atmosphere-controlled, wide-field fluorescence microscope. In this system, Rac1 activation leads to the formation of the complex mCit-Rac1/mCFP-PBD, resulting in the generation of a FRET signal that can be analyzed quantitatively using the FRET stoichiometry method 162, 163 (Figure A.1 and Supplemental Movie S1). Cells that coexpressed mCFP-PBD and mCit-Rac1 were imaged for 5 min before the addition of physiologically relevant concentrations of LLO

(0.1 nM). Importantly, this toxin concentration is nonlethal and sufficient to induce the entry of LLO-deficient L. monocytogenes and BSA-coated beads when LLO is added exogenously to the cell culture medium 48. LLO-exposed cells exhibited elevated levels of

Rac1 activation, as evidenced by transient FRET flashes that paralleled the appearance and disappearance of membrane ruffles in successive waves at different cellular locations

(Figure 2.3A and Supplemental Movie 1). Because Rac1 was activated only within discrete cell regions in a transient manner, we quantified Rac1 FRET in regions of standard size that were positioned where Rac1 activation was observed or positioned arbitrarily if Rac1 40 activation was absent. Two regions of standard size were positioned on each cell and were analyzed whether Rac1 FRET fluctuations were observed. Data show that Rac1 activation was detected in 76.61% of cells exposed to LLO (Table 2.1; corresponding statistical analyses in Table A.2).

Toxin - LLO LLOpL LLOpL - LLO - LLO (0.1 nM) (10 nM) Medium (CaCl2) + + + + + + - - Time (min) 20 20 20 20 10 10 10 10 NC 91 75 164 148 91 75 125 173 FRET+ cells 3 57 9 12 0 56 8 10 % FRET 2.43 76.61 5.88 7.22 0 74.83 4.78 5.68 Table 2.1 LLO induces Rac1 activation in a Ca2+- and pore-dependent manner Cells were exposed, or not, to 0.1 nM LLO, 0.1 nM LLOpL, and 10 nM LLOpL in cell culture medium with (+) or without (−) CaCl2. Time indicates the total duration of imaging. In Ca2+-free medium (−) cells were imaged for only 10 min due to cell damages caused by LLO. NC is the number of analyzed cells. FRET+ cells is the number of FRET-positive cells (exhibiting at least twofold increase in FRET). % FRET is the percentage of FRET- positive cells (n > 3).

Rac1 activation was defined as an at least twofold increase in FRET efficiency within the analyzed region (FRET-positive region). In contrast, only 2.43% of control cells (not exposed to LLO) were positive for Rac1 FRET. Rac1 activation kinetics measured in all regions and kinetics of Rac1 activation averaged across all regions are presented in Figure

2.3 B–D.

41

Figure 2.3: FRET stoichiometry to monitor Rac1 activation by LLO. HepG2 cells coexpressing mCFP-PBD and mCit-Rac1 were imaged on the microscope stage at 37˚C. Phase-contrast (PC), and fluorescence images (IA, ID, and IF) were acquired every 20 s for 20 min. LLO was added after 5 min of imaging (T0). (A) PC, overlay of PC and IA, IA, ID, IF, and FRET (EAVE presented in pseudocolor scale). Scale bar = 10 μm. From top to bottom: time points 0, 3, and 5 min after addition of LLO. (B, C) For each imaged cell, two regions of standard size (305 pixels) were applied at locations where FRET variations were observed. If no FRET variation was observed, then the regions 42 were arbitrarily positioned. The y-axis indicates the fold change in EAVE and the z-axis represents all individual regions of control cells (182 regions, 91 cells) (B), and all FRET-positive regions from the LLO-exposed cells (92 regions, 57 cells) (C), in which FRET positive is defined as a greater than twofold increase in FRET efficiency (n = 3). (D) Kinetics of the averaged EAVE of control (182) and LLO-exposed (92) regions. The black vertical line indicates the time of addition of medium alone (control) or 0.1 nM LLO.

This analysis showed that at the cell population level, Rac1 was activated ∼40 s after addition of LLO with a maximal amplitude at 80–120 s for a duration of 2 min.

Statistical analyses of the FRET kinetics presented in Figure 2.3 are presented in Table

B.3. To establish the role of LLO pore formation in Rac1 activation, cells were exposed to an LLO variant (listeriolysin O prepore lock, LLOpL) that binds to the host cell plasma membrane and forms a prepore complex but is unable to transition into the transmembrane

β-barrel pore 48, 164. Indeed, LLOpL was shown to be inactive at concentrations exceeding

4 10 nM as compared with native LLO (EC50 = 1 nM), but its pore-forming activity was restored to the native LLO level upon treatment with 4 mM dithiothreitol (DTT) (Figure

A.2) 118. Treatment with 0.1 and 10 nM LLOpL failed to activate Rac1 in HepG2 cells

(Table 2.2; statistical analyses in Table A.2). Thus, as reported for LLO-dependent L. monocytogenes entry and membrane ruffling, Rac1 activation requires LLO pore formation 48. Formation of the transmembrane pore leads to an influx of extracellular Ca2+, which is required for LLO-mediated L. monocytogenes entry 55. To prevent Ca2+ influx, cells were analyzed in Ca2+-free medium. Because the influx of Ca2+ is indispensable for membrane resealing, for in the absence of extracellular Ca2+ cell integrity is rapidly

43 compromised upon damage, FRET analysis was limited to time points 0–5 min post–LLO exposure 55. In the absence of extracellular Ca2+, LLO-induced Rac1 activation occurred in

5.68% of cells compared with 74.83% in the presence of extracellular Ca2+ (Table 2.2; statistical analyses in Table A.3). Collectively, these data establish that LLO is sufficient to activate Rac1 within membrane ruffles in a pore- and Ca2+-influx–dependent manner.

LLO-mediated Rac1 activation results in Arp2/3-dependent F-actin remodeling The subcortical F-actin cytoskeleton acts as a dynamic scaffold that promotes the formation of membrane ruffles and the formation of the phagocytic cup 165. We used a live- cell fluorescence imaging approach to visualize F-actin rearrangement in Tomato-β-actin- expressing HepG2 cells. We could clearly visualize the formation of F-actin bundles within membrane ruffles in cell exposed to LLO (Figure 2.4 and Supplemental Movie 2). We observed by live-cell phase-contrast imaging that 85% of cells exposed to LLO initiated membrane ruffling as early as 84.8 ± 26.7 s (average ± SD; >100 cells, n = 8) following exposure to LLO. Additionally, LLO-induced membrane ruffling was frequently followed by the formation of large endocytic vacuoles reminiscent of macropinosomes

(Supplemental Movie 3).

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Figure 2.4: LLO induces F-actin rearrangement. HepG2 cells expressing Tomato-β-actin were imaged on the microscope stage at 37˚C. Phase-contrast (PC, column 1), and fluorescence images were acquired every 20 s for 20 min. LLO was added after 5 min of imaging (T0). LLO induces dynamic F-actin rearrangement (red arrows) within membrane ruffles. Regions highlighted in column 2 are shown in column 3. Scale bar = 10 μm.

F-actin polymerization can involve two types of actin nucleators: formins and the

Arp2/3 complex 166. Because Arp2/3 is most commonly involved in remodeling F-actin at

45 bacterial entry sites, we focused on establishing the role of the Arp2/3 complex in HepG2 cells exposed to LLO in the presence or absence of the Arp2/3 inhibitor CK-666 167.

Inhibition of Arp2/3 abolished LLO-induced membrane ruffling, as assessed by live cell phase-contrast imaging (0% of cells exhibited membrane ruffling; >100 cells, n = 3) as compared with control cells (85%, >100 cells, n = 3; Supplemental Movies 4 and 5).

Therefore, LLO induces Arp2/3-dependent de novo F-actin remodeling and subsequent membrane ruffling. To establish whether Rac1 activation occurs upstream from Arp2/3- dependent F-actin remodeling, cells were transfected to express fluorescent chimeras of the native (mCit-Rac1)- or dominant-negative (mCit-Rac1N17)-Rac1 or fluorophore alone

(mCit). In cells expressing the fluorophore alone (126 cells, n = 3) or mCit-Rac1 (112 cells, n = 3) 75.4% and 58.04% of cells exhibited membrane ruffling in response to LLO, respectively. However, cells expressing mCit-Rac1N17 (114 cells, n = 3) exhibited substantial decrease in membrane ruffling upon LLO exposure (6.14% ruffling). As expected, nonfluorescent neighboring cells that did not express mCit-Rac1N17 exhibited membrane ruffling, further supporting the hypothesis that Rac1 activation is a prerequisite for LLO-induced membrane ruffling (Supplemental Movie 6). Importantly, we also established that the Arp2/3 complex is required for L. monocytogenes entry into HepG2 cells using the Arp2/3 inhibitors CK-636 and CK-666. Figure 2.5 shows the dose- dependent inhibition of L. monocytogenes entry by the Arp2/3 inhibitors, indicating that the same machinery that is driving LLO-mediated F-actin–dependent remodeling is also necessary for L. monocytogenes internalization. Collectively, our data indicate that LLO-

46 mediated entry involves Rac1 activation upstream from Arp2/3-dependent F-actin remodeling.

Figure 2.5: Role of Arp2/3 in LLO-dependent entry of L. monocytogenes. Nontreated and cells treated with 0.2% DMSO or the Arp2/3 inhibitors CK-636 and CK- 666 were infected with WT L. monocyto•genes at MOI 20 for 30 min at 37˚C. Cells were fixed, and bacteria were fluorescently labeled to enumerate the total number of bacteria (Nt) and the number of extracellular bacteria associated with host cells (Ne). Entry efficiency was measured as 100 × [(Nt) - (Ne)]/(Nt), and results are expressed as the mean percentage entry ± SEM relative to the DMSO control. Statistics are as follows: *p < 0.05; ***p < 0.005 (n = 3).

A conventional PKC is required for LLO-mediated entry of L. monocytogenes The conventional Ca2+-responsive PKC alpha (PKCα) has been shown to be involved in Rac1 translocation to the plasma membrane of cells treated with thapsigargin or ionomycin, which, similarly to LLO, cause an increase in cytosolic Ca2+ 168. To determine whether a PKC is involved in L. monocytogenes internalization into HepG2 cells, we measured L. monocytogenes entry into cells pretreated with broad spectrum

(Gö6983) and conventional (GF109203x) PKC inhibitors. Treatment with 0.5 µM Gö6983 or 1 µM GF109203x resulted in a 59.9 ± 1.5% (p < 0.05) and 61.5 ± 1.9% (p < 0.01) 47 decrease in L. monocytogenes entry, respectively, relative to the vehicle control, indicating a role for a conventional PKC (Figure 2.6).

Figure 2.6: Role of a conventional PKC in LLO-dependent entry of L. monocytogenes. Nontreated and cells treated with 0.2% DMSO, the broad spectrum (Gö6983) or conventional (GF109203x) PKC inhibitors, were infected with WT L. monocytogenes at MOI 20 for 30 min at 37˚C. Cells were chemically fixed, and bacteria were fluorescently labeled to enumerate the total number of bacteria (Nt) and the number of extracellular bacteria associated with host cells (Ne). Entry efficiency was measured as 100 × [(Nt) - (Ne)]/(Nt), and results are expressed as the mean percentage entry ± SEM relative to the DMSO control. Statistics are as follows: *p < 0.05; **p < 0.01 (n = 3).

A conventional PKC is activated by the influx of extracellular Ca2+ leading to Rac1 activation We then focused on demonstrating the activation of a PKC by LLO. We transfected

HepG2 cells with fluorescent chimeras of two conventional (α, βII) and two novel (ε, δ)

PKC isoforms. HepG2 cells exposed to LLO exhibited dynamic translocation of GFP-

PKCα (translocation was observed in 70.8% of transfected cells; 250 cells, n = 7; Figure

2.7 and Supplemental Movie 7), GFP-PKCε (63.63%; 22 cells, n = 1), and GFP-PKCβII

(37.5%; 24 cells, n = 1), but not of GFP-PKCδ (0%; 65 cells, n = 2) or GFP alone (0%; 32

48 cells, n = 1; Supplemental Movie 7) to regions of plasma membrane ruffles as early as 40 s after addition of LLO.

Figure 2.7: LLO induces PKCα translocation to the plasma membrane. HepG2 cells expressing GFP-PKCα were imaged on the microscope stage at 37˚C. Phase-contrast (PC), and fluorescence images were acquired every 20 s for 20 min. LLO was added after 5 min of imaging (T0). LLO induces GFP-PKCα translocation to regions

49 of dynamic plasma membrane ruffles and around newly formed vesicles. Regions highlighted in column 2 are shown in column 3. Scale bar = 10 μm.

We further established that the influx of extracellular Ca2+ was indispensable for

PKC translocation, as observed in cells exposed to LLO in Ca2+-free medium (0% of GFP-

PKCα translocation, 23 cells, n = 1; Supplemental Movie 8). Importantly, cells treated with the conventional PKC inhibitor GF109203x did not undergo F-actin–dependent membrane ruffling in response to LLO exposure (3.5% of cells exhibited membrane ruffling, >100 cells, n = 3), suggesting that a Ca2+-responsive conventional PKC was involved in Rac1 activation. To demonstrate that LLO activates a PKC, we used the PKC FRET biosensor myristoylated and palmitoylated c kinase activity reporter (MyrPalm-CKAR), which has a

20% dynamic range and was designed to specifically assess PKC activity at the plasma membrane 169, 170. In the nonphosphorylated state, MyrPalm-CKAR exists in a constrained conformation that places the N-terminal mCFP in close proximity to the C-terminal mCit leading to FRET, but phosphorylation of the interconnecting sequence by any PKC isoform relaxes the sensor leading to a loss of FRET. We tested whether addition of the

Ca2+ ionophore, ionomycin, which is known to activate PKCs, would lead to pronounced phosphorylation of the biosensor. As expected, addition of ionomycin resulted in a sudden drop in FRET efficiency followed by a quick recovery (Figure A.3). This result is consistent with the notion that Ca2+ influx leads to activation of PKC at the plasma membrane followed by dephosphorylation of the biosensor thereafter, as previously observed in mouse oocytes 171. HepG2 cells expressing MyrPalm-CKAR and exposed to

50

LLO exhibited an initial drop in FRET, that is, MyrPalm-CKAR phosphorylation, 40 s after addition of LLO, which overlays with the kinetics of PKCα translocation and Rac1 activation (Figure 2.8, Supplemental Movie 9, and statistical analyses in Table A.4).

Figure 2.8: A PKC is activated by LLO. HepG2 cells expressing MyrPalm-CKAR were imaged on the microscope stage at 37˚C. Phase-contrast (PC) and fluorescence images (IA, ID, and IF) were acquired every 20 s for 20 min. LLO was added after 5 min of imaging (T0 and black vertical line). (A) A representative cell exhibiting LLO-induced MyrPalm-CKAR phosphorylation. From left to right: PC, overlay of PC + IA, IA, ID, and IF, and FRET (in pseudo-color scale). From top to bottom: time points 0, 1, and 5 min after addition of LLO. Scale bar = 10 μm. (B) Data are presented as the average fold change in EAVE ± SEM (n = 3, 24 cells in DMSO control, 26 cells in GF109203x, 28 cells in DMSO +LLO, and 26 cells in GF109203x +LLO).

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The magnitude of change in FRET efficiency in LLO-exposed cells is 65% lower than observed in ionomycin-treated cells, but the duration of PKC activity is prolonged, which may be due to smaller and prolonged Ca2+ fluxes in LLO-exposed cells. Importantly, LLO- induced MyrPalm-CKAR phosphorylation was inhibited by cell pretreatment with the conventional PKC inhibitor GF109203x (Figure 2.8). We then established whether a conventional PKC was involved in Rac1 activation. Rac1 activation was measured by

FRET stoichiometry as presented in Figure 2.3. We found that Rac1 was no longer activated by LLO in cells pretreated with the conventional PKC inhibitor GF109203x

(Table 2.2; statistical analyses in Table A.5). Collectively, our data show that a conventional PKC links Ca2+ influx to Rac1 activation and Arp2/3-dependent F-actin remodeling for bacterial internalization.

Toxin - LLO LLO Medium DMSO DMSO GF109203x

Time (min) 20 20 20 NC 115 92 136 FRET+ cells 1 50 26 % FRET 0.87 54.3 19.12 Table 2.2: Rac1 is activated downstream of a conventional PKC Cells were exposed to DMSO (0.2%), 0.1 nM LLO, or GF109203x. Time indicates the total duration of imaging. NC is the number of analyzed cells. FRET+ cells is the number of FRET-positive cells (exhibiting at least a twofold increase in FRET). % FRET is the percentage of FRET-positive cells (n > 3).

Inhibition of the LLO-mediated entry pathway facilitates membrane resealing We previously showed that cells exposed to physiological concentrations of LLO undergo rapid Ca2+-dependent membrane resealing 55. To investigate if LLO-mediated L. 52 monocytogenes entry pathway is part of the membrane resealing machinery, we measured the efficiency of membrane resealing in cells pretreated with the inhibitors that prevent

LLO-induced L. monocytogenes entry 172. HepG2 cells were incubated with inhibitors of conventional PKC, Arp2/3, or F-actin and exposed to LLO. As a control, inhibition of membrane resealing was achieved by incubating cells in Ca2+-free medium. These experiments were performed in the presence of the plasma membrane-impermeant nucleic acid–binding probe propidium iodide. Propidium iodide flows through damaged cell membranes, leading to a strong, quantifiable fluorescence signal, whereas cells with intact plasma membranes exclude the probe or are minimally labeled. We found that HepG2 cells treated with the conventional PKC inhibitor, the Arp2/3 inhibitor, or the F-actin inhibitor and exposed to LLO, resealed their plasma membrane more efficiently than cells treated with LLO (+ vehicle) in the absence of inhibitors (Figure 2.9).

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Figure 2.9: The LLO-dependent entry pathway is dispensable for membrane resealing. HepG2 cells were plated in a 96-well plate and incubated in the presence of DMSO (control), GF109203x, cytochalasin D (CD), or CK-666 in Ca2+-containing medium (M1) or in DMSO in Ca2+-free medium (M2, control for membrane repair). Cells were exposed to 0.25, 0.5, or 1 nM LLO, and the fluorescence intensity of propidium iodide (PI) was measured in the plate reader every 5 min for 30 min at 37˚C. Experimental triplicates were performed for each condition, and three independent experiments were performed. Data are presented as the average PI intensity ± SEM. Statistical analyses were performed as described under Materials and Methods and are presented in the text box (n = 3).

Indeed, statistical analyses revealed that the presence of the inhibitors improved cell integrity (p < 0.0001). Importantly, LLO pore-forming activity was not altered by any of these inhibitors (Figure A.2). In conclusion, LLO pore formation was followed by F-actin–- independent membrane resealing and by F-actin–dependent bacterial entry. Both processes

54 were initiated by the influx of extracellular Ca2+, but involved distinct signaling pathways thereafter.

2.3 Discussion

This study identified a signaling pathway activated by the pore-forming toxin listeriolysin O (LLO) to promote the internalization of the bacterial pathogen L. monocytogenes into human hepatocytes. LLO is an exotoxin that forms large pores across the host cell plasma membrane causing a rapid influx of extracellular Ca2+. Ca2+ influx is then involved in the translocation and activation of conventional protein kinase C (cPKC) at the plasma membrane. Subsequently, the cPKC is required for Rac1 activation upstream of Arp2/3-dependent de novo assembly of the F-actin cytoskeleton for L. monocytogenes internalization (Figure 2.10).

Figure 2.10: Model for LLO-mediated L. monocytogenes entry. L. monocytogenes secretes the pore-forming toxin listeriolysin O (LLO), which binds to the cholesterol-rich plasma membrane and oligomerizes into a prepore complex. The prepore complex then transitions into a transmembrane pore, which permits the influx of extracellular Ca2+. A rise in cytosolic Ca2+ results in the translocation and activation of a 55 conventional PKC at the plasma membrane. Activated PKC promotes the activation of the Rho GTPase Rac1, which transduces the signal down to induce Arp2/3-mediated F- actin remodeling at the plasma membrane leading to L. monocytogenes entry. The influx of extracellular Ca2+ also induces a membrane resealing pathway.

This is the first identification of a signaling pathway activated downstream from plasma membrane perforation by a pore-forming toxin leading to the internalization of a bacterial pathogen. Because pore-forming toxins are virulence factors produced by a large diversity of pathogens, perforation of the host cell plasma membrane is likely a ubiquitous mechanism by which pathogens hijack signaling to enter host cells.

Importantly, this work also shows that the identified pathway is dispensable for efficient resealing of the plasma membrane. Although membrane resealing is required for L. monocytogenes internalization, for in its absence cell integrity is lost, the signaling pathway directing F-actin remodeling at the bacterial entry site is distinct from the membrane resealing pathway.

Link between pathogen internalization and resealing of the injured plasma membrane

Studies on the eukaryotic parasite T. cruzi and the adenovirus HadV-C2 proposed that pathogens damage the host cell plasma membrane and co-opt the subsequent membrane repair machinery to promote their endocytosis 50, 53. It is thought that flagellar- mediated motility of T. cruzi induces mechanical damages and that the adenovirus perforates the cell by uncoating of the membrane damaging protein-VI. Both studies support the hypothesis that cell injury leads to the influx of extracellular Ca2+, which

56 promotes plasma membrane resealing via the following sequence of events: Ca2+- dependent exocytosis of lysosomes, release of the lysosomal enzyme acid sphingomyelinase (ASM), conversion of sphingomyelin into ceramide by ASM on the outer leaflet of the plasma membrane, and ceramide-mediated endocytosis 154. This pathway leads to the endocytosis of the damaged membrane together with the pathogen

51. Furthermore, it was established that resealing is independent of F-actin, dynamin, and clathrin, but requires caveolin-rich membrane microdomains 85, 154. While the evidence supports that membrane resealing is required for pathogen internalization, it is unclear whether the membrane resealing process is directly responsible or whether it is a prerequisite for pathogen uptake. In the latter hypothesis, resealing would be necessary to maintain cell homeostasis and allow for a distinct pathway to promote uptake of the pathogen. Our studies favor the latter hypothesis. Indeed, we identified signaling events required for LLO-dependent L. monocytogenes internalization that were dispensable for membrane resealing. Inhibition of cPKC, Arp2/3, and F-actin assembly improved the efficiency of cell resealing (Figure 2.9) without altering LLO pore formation (Figure

A.2). Similar to previous studies, we found that resealing of cells damaged by LLO is independent of F-actin, clathrin, and dynamin 48. Therefore, the resealing machinery downstream from LLO perforation is expected to proceed via the same pathway as described for T. cruzi and adenovirus, but LLO-induced L. monocytogenes endocytosis does not directly involve the resealing machinery. In accordance with our findings, studies have also identified roles for cPKC and F-actin reorganization in entry of adenovirus and T. cruzi into nonprofessional phagocytes 173-176. Whether or not T.

57 cruzi and adenovirus utilize the resealing machinery or the signaling pathway described in this work to gain entry into cells, pathogen-mediated host plasma membrane damage represents an invasion mechanism broadly used by unrelated pathogens.

Role of F-actin remodeling in L. monocytogenes uptake and in membrane resealing

Internalization of most pathogens requires F-actin remodeling for the formation of the endosome that contains the pathogen. This was demonstrated for most, if not all, bacterial pathogens, including L. monocytogenes invasion of professional phagocytes and normally nonphagocytic cells as well as for the endocytosis of viruses, such as the adenovirus and for T. cruzi invasion of nonphagocytic cells 173, 177, 178. Similarly, the

LLO-dependent L. monocytogenes entry pathway is dependent on de novo F-actin assembly 48. A point of convergence is that an intact F-actin cytoskeleton is not required for resealing the plasma membrane during pathogen uptake 53, 55, 82. Plasma membrane resealing is thought to require initial disassembly of cortical F-actin to 1) clear the way for docking and fusion of lysosomes with the plasma membrane and 2) decrease surface tension, which prevents enlargement of the plasma membrane wound 63. In accordance with this model, our data show that inhibiting signaling events that favor F-actin assembly improves plasma membrane resealing (Figure 2.9). Thus, how does one explain that cells undergo de novo F-actin assembly in response to perforation by LLO if F-actin assembly is dispensable for resealing? Similarly to our observation, membrane damage in Xenopus laevis oocytes was followed by Rho GTPase activation and F-actin assembly

179. In this example, F-actin assembly was proposed to facilitate repair of very large mechanical wounds. It is possible that injured cells respond to wounding by a sequence 58 of events that include initial disassembly of the actin network followed by its reassembly, regardless of the nature of the wound. We propose that F-actin reassembly 1) repairs large mechanical wounds but is dispensable for repairing smaller injuries caused by toxin pores and 2) restores the cortical F-actin network. Alternatively, one can propose that the reassembly of the F-actin network is due to an additional property of LLO.

LLO is sufficient to activate cPKC-, Rac1-, and Arp2/3-dependent F-actin remodeling

LLO is a multifunctional toxin that affects transcriptional regulation, protein ubiquitination, and mitochondrial dynamics 54, 118, 180, 181. The present studies identified

LLO as a regulator of the F-actin cytoskeleton. To monitor the spatiotemporal dynamics of signaling events downstream from LLO pore formation, we used a highly sensitive live-cell FRET-based approach that quantitatively detects small changes in protein activation in single cells 162, 163. This approach revealed that the kinetics of PKC activation overlay and potentially precede Rac1 activation, a fact that we confirmed with pharmacological inhibitors (Figures 2.3 and 2.8). Interestingly, Rac1 activation proceeded in a wavelike pattern, reminiscent of calcium waves. This could be observed because we used a fluorescent chimera of native Rac1. However, the MyrPalm-CKAR biosensor is artificially anchored to the plasma membrane, which allows us to monitor the kinetics of PKC activation at this location, but not native PKC translocation 169, 170, 182. To assess PKC translocation, we used cells expressing fluorescent chimeras containing the membrane-binding domains of the PKC isoforms PKCα, PKCβII, and PKCε. Fluorescent cPKC domains were translocated to regions of the membrane where membrane ruffles

59 and macropinosomes were formed similarly to the Rac1 activation pattern (Figure

2.7 and Supplemental Movie 7). The present work demonstrates for the first time the activation of a conventional PKC by LLO. Previous studies reported the activation of the calcium-independent PKCε (novel PKC) 183. Because activation of conventional and novel PKC requires production of diacylglycerol (DAG), LLO may also activate host phospholipases, as previously suggested 183. Our finding that Ca2+ influx leads to cPKC activation upstream to Rac1 is reminiscent of previous work using the calcium ionophore ionomycin 168. In that study, a cPKC responsive to Ca2+ influx phosphorylated a RhoGDI facilitating Rac1 activation 168. Thus, the influx of Ca2+ through LLO pores could suffice to activate Rac1 in a similar manner. This idea is supported by the dependence of PKC and Rac1 activation on Ca2+influx and the fact that the prepore-locked form of LLO

(LLOpL) was inactive. In accordance with our results, the Streptococcus pneumoniae CDC pore-forming toxin pneumolysin induces Ca2+-dependent F-actin remodeling in a Rho GTPase (Rac1, Cdc42, and RhoA)-dependent manner 184. Our FRET studies established that LLO also activates Cdc42 (Figure A.3), but to a lower extent than

Rac1, and we showed that Cdc42 (and RhoA) are both dispensable for the LLO-mediated entry pathway.

Why does L. monocytogenes use multiple invasins and invasion strategies?

Listeria monocytogenes infects multiple nonphagocytic cell types, including enterocytes, hepatocytes, cardiac myocytes, neurons, cytotrophoblasts, endothelial cells, and so on. It is therefore not surprising that this pathogen evolved to express multiple strategies to invade host cells. The major L. monocytogenes invasin, InlA, acts by 60 attaching the bacterium to the cell adhesion molecule E-cadherin but not all cell types infected by L. monocytogenes express E-cadherin 113, 185, 186. The second invasion protein

InlB acts via activating c-Met and was shown to cooperate with InlA to potentiate the bacterial internalization pathway 114, 187-189. Although the initial stage of activation of bacterial internalization differs for each invasin (InlA binds to E-cadherin, InlB activates c-Met, LLO forms pores) they all converge toward activation of Rho GTPases and the

Arp2/3 complex leading to F-actin– and dynamin-dependent internalization 142. More recently it was shown that the InlB internalization pathway involves a PKC similarly to our finding with LLO 190. InlA, InlB, and LLO are coexpressed and may cooperate to activate a more robust endocytic signaling in normally nonphagocytic cells.

Alternatively, the invasins may promote L. monocytogenes invasion in a cell-type– dependent manner.

Overall, this work confirms the novel concept that pathogens use host cell plasma membrane damage as a strategy to induce their endocytic uptake into normally nonphagocytic cells, thereby ensuring the first step of their intracellular life cycle. Unlike previous studies, this work proposes that the plasma membrane resealing pathway is a prerequisite but is not directly responsible for pathogen uptake. We propose that in the example of L. monocytogenes, a Ca2+-dependent pathway that is distinct from plasma membrane resealing is activated downstream from cell injury to restore the cortical actin cytoskeleton, which is hijacked for pathogen internalization.

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2.4 Materials and Methods

Reagents and inhibitors

Cells were pretreated for 1 h with the following pharmacological inhibitors

(Sigma-Aldrich): kanamycin (30 μg/ml), ampicillin (100 μg/ml), chloramphenicol (5

μg/ml), CK-636 and CK-666 (50–150 μM), GF109203x (0.05–1.0 μM), and Gö6983 (0.5

μM). Cells were pretreated for 10 min with cytochalasin D (1 μM; Sigma-Aldrich).

Antibodies include the following: anti–L. monocytogenes rabbit polyclonal antibodies

(GeneTex); goat anti-rabbit secondary antibodies conjugated to Alexa488 and Alexa568

(Molecular Probes); anti-BSA rabbit antiserum (B1520; Sigma-Aldrich); mouse monoclonal antibodies directed against Rac1, RhoA, and Cdc42 (clones ARC03, ARH04, and ACD03, respectively; Cytoskeleton); rabbit anti-actin (clone A2103; Sigma-Aldrich); and secondary goat anti-mouse immunoglobulin G (IgG) and goat anti-rabbit IgG antibodies conjugated to horseradish peroxidase (Cell Signaling). FluoSpheres carboxylate modified microspheres (1 μm) (Molecular Probes), BSA (Santa Cruz

Biotechnology), and 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride

(Thermo-Fisher) were used to prepare the beads for LLO coating. Lipofectamine

RNAiMax and Lipofectamine 2000 (Life Technologies) were used to transfect siRNA and plasmid DNA, respectively, using the manufacturer’s protocols. Silencer select siRNA for Rac1, RhoA, Cdc42, and NC were used at 60–90 pg per well in a six-well format (Life Technologies).

Listeriolysin O purification

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Escherichia coli BL21(DE3) containing pET29b hly, which encodes a C-terminal

6His-tagged listeriolysin O missing the signal sequence (N-terminal 25 amino acids), or pET29b hly (G80C, S215C), which encodes a C-terminal 6His-tagged listeriolysin O missing the signal sequence (N-terminal 25 amino acids) and contains two amino acid substitutions (G80C, S215C) to produce the prepore locked variant LLOpL, were grown on Luria broth (LB) agar containing 30 μg/ml kanamycin at 37˚C overnight. Bacterial cultures were grown overnight at 37˚C in LB-supplemented kanamycin (30 μg/ml). The overnight culture was subcultured in 0.5 l LB supplemented with kanamycin and grown to OD600 = 0.6 at which point isopropyl β-d-1-thiogalactopyranoside (1 mM final concentration) was added to induce expression of the 6His-tagged listeriolysin O and the culture was agitated at room temperature for 5 h. The culture was pelleted at 6000 × g for

10 min. All buffers used for the LLOpL purification lacked beta-mercaptoethanol (BME) and dithiothreitol (DTT). Cell pellets were suspended in lysis buffer (50 mM phosphate, pH 8.0, 1 M NaCl, 5 mM BME, and protease inhibitor) and lysed using a French press.

Cell lysate was centrifuged and the supernatant was applied to Ni-NTA Agarose (Qiagen) for 2 h at 4°C. The agarose was washed with (50 mM phosphate, pH 6, 1 M NaCl, 20 mM imidazole, 5 mM BME, 0.1% Tween-20, 10% glycerol) followed by two washes with (50 mM phosphate, pH 6, 1 M NaCl, 20 mM imidazole, 5 mM BME). The protein was eluted with (50 mM phosphate, pH 6, 1 M NaCl, 500 mM imidazole, 5 mM BME) and dialyzed in 50 mM phosphate, pH 6, 0.5 M NaCl, 5 mM DTT.

Bacterial cultures

63

Wild-type L. monocytogenes (strain 10403s) and its isogenic hly deletion mutant were gifts from Dan Portnoy (University of California, Berkeley). Bacterial cultures were grown overnight at 37˚C under agitation in brain and heart infusion (BHI; BD

Biosciences). Escherichia coli XL1blue and DH5α were grown at 37˚C under agitation in

LB supplemented with kanamycin (30 μg/ml), ampicillin (100 μg/ml), or chloramphenicol (5 μg/ml).

Plasmid construction and purification

Escherichia coli XL1Blue harboring the plasmids pmECFP-C1, pmEYFP-C1, pECFP-YFP, pmECFP-PBD, pmEYFP-Rac1, pmEYFP-Rac1-T17N, and pmEYFP-

Rac1-H40Y/Q61L were prepared as described previously 162, 163. Sanger sequencing with primers A and B was used to verify insert sequences (Table B.6). To obtain the constitutively active form of Rac1 (pmEYFP-Rac1-Q61L), pmEYFP-Rac1-H40Y/Q61L was mutated via site-directed mutagenesis using primers C and D. Sanger sequencing with primers A and B was used to verify that the proper mutation was made. Escherichia coli DH5α containing pcDNA3-hPKCβII-YFP, pcDNA3-hPKCε-HA1-GFP, and pcDNA3-hPKCδ-HA1-GFP and E. coliXL1Blue containing pEGFP-PKCα were gifts from Alexandra Newton (University of California, San Diego) 182. Escherichia coli containing pcDNA3 constructs was grown in LB supplemented with 100 µg/ml ampicillin while E. colicontaining pEGFP constructs was grown in LB supplemented with 30 µg/ml kanamycin. Escherichia coli DH5α containing pcDNA3 MyrPalm-CKAR was grown in LB supplemented with 100 µg/ml ampicillin (Addgene) 170. Plasmids pEGFPN1 and pEGFP-PKCα were submitted for Sanger sequencing using primers E and 64

G and E, F, and G, respectively (Table B.6). The GenElute Endotoxin-free Plasmid

Midiprep Kit was used to purify endotoxin-free plasmids according to the manufacturer instructions (Sigma-Aldrich).

Mammalian cell culture, silencing, and transfection

The human hepatocyte carcinoma cell line HepG2 (ATCC HB-8065) was grown in MEM with Earle’s salts and 2 mM l-glutamine supplemented with 10% heat- inactivated fetal bovine serum (FBS; Lonza), 0.1 mM nonessential amino acids, 1 mM sodium pyruvate, 100 U/ml penicillin, and 100 μg/ml streptomycin (Invitrogen). Cells were grown at 37˚C in a 5% CO2 humidified cell culture incubator. Cells were seeded at

1 × 104 cells/well in a 24-well tissue culture plate, 4 × 105 cells/dish in No. 1.5, 10-mm glass diameter MatTek dish for live cell imaging, or 5 × 105 cells/well in six-well tissue culture plates containing 22 × 22 mm No. 1.5 cover glass (Gold Seal). To silence the expression of the Rho GTPases, cells were transfected in antibiotic-free medium using

Lipofectamine RNAi Max (Invitrogen) with silencer select siRNA for Rac1, RhoA,

Cdc42, or negative control siRNA. The media was replaced 24 h after seeding and incubated for an additional 24 h. To monitor Rho GTPase activation by FRET microscopy, cells were seeded at 4 × 105 cells in 35 mm No. 1.5, 10-mm-diameter glass

MatTek dishes (MatTek P35GC-1.5-10-C). Twenty-four hours after seeding, cells were transfected with 2500 μg total plasmid DNA in antibiotic- and serum-free medium using

Lipofectamine 2000 (ThermoFisher Scientific). Serum was added to a final 10% after 1 h, and then the cell culture media was replaced with antibiotic-free medium after an additional 1 h. 65

Western blot analysis for Rac1, RhoA, and Cdc42 in HepG2 expression

After 48 h of treatment with, or without (control, nontreated), the siRNAs, HepG2 cells were washed once with PBS, pH 7.4, and lysed in 100 μl boiling reduced sample buffer. Cell lysates were boiled for 5 min and equivalent to 1.5 × 105 (nondiluted), 7.5 ×

104 (1:2 dilution), 3.75 × 104 (1:4 dilution), and 1.88 × 105 (1:8 dilution) cells were loaded into a 10% SDS–PAGE. All samples were subjected to SDS–PAGE and Western blot analysis using Rac1, RhoA, Cdc42, and actin antibodies, followed by a goat anti- mouse IgG conjugated to HRP or goat anti-rabbit IgG conjugated to HRP. Western blots were developed using the Amersham ECL Select Detection Reagent (GE Healthcare Life

Sciences) and imaged using a ChemiDoc XRS+ system and ImageLab software (Bio-

Rad). Once the conditions for knockdown were established, at least three independent experiments were performed to confirm efficient silencing, and Western blot analyses were performed in each experiment to verify knockdown efficiencies.

Densitometry analysis

Nontreated and siRNA-treated HepG2 cell lysates were processed for Western blot analysis. Densitometry analysis using ChemiDoc XRS+ system and ImageLab software (Bio-Rad) was performed to assess band intensities (verified for nonsaturation), and all values were normalized relative to NC siRNA-treated sample. The ratio of Rho

GTPase to actin was then calculated to compare the Rho GTPases expression levels

(Table B.1).

Bacterial invasion assay

66

Overnight cultures of L. monocytogenes were diluted 1:20 in BHI and grown at

37˚C to mid–log phase at an OD600 = 0.8. Bacteria were washed 3 times in phosphate- buffered saline (PBS), pH 7.4, diluted into MEM at a multiplicity of infection (MOI) 20 and added to the mammalian cells that were seeded at 5 × 105 cells/well in a 24-well plate and cultured for 48 h. Cell culture plates were centrifuged at 520 × g for 3 min at room temperature and incubated for 30 min at 37˚C. Infected cells were washed with MEM twice followed by two additional washes in PBS and fixed with 4% paraformaldehyde

(PFA) in PBS, pH 7.4, for 15 min at room temperature. Cells were washed three times with 0.1 M glycine in PBS and blocked in 0.1 M glycine in PBS/10% FBS for 1 h. For labeling, extracellular bacteria were labeled with the primary anti–L. monocytogenes antibody and secondary anti-rabbit antibody conjugated to Alexafluor-488, and total bacteria were labeled, after cell permeabilization with 0.5% TX-100 (in PBS), with a primary anti–L. monocytogenes rabbit polyclonal antibody and a secondary anti-rabbit antibody conjugated to Alexafluor-568. Cells were preserved in Prolong Gold Antifade

Mountant medium containing 4’,6 diamidino-2-phenylindole, dihydrochloride (DAPI).

To quantify the numbers of bacteria, 40 sets of images (DAPI, Alexa-488, Alexa-568, and phase-contrast) were randomly acquired using a Metamorph programmed, motorized stage and the 20× objective. Metamorph imaging and analysis software were used to determine the number of extracellular (Ne) and total (Nt) bacteria such that the efficiency of bacterial entry was measured as the percentage of intracellular bacteria (Ni) using the equation Ni = (Nt - Ne)*100/Nt.

Polystyrene beads invasion assay 67

FluoSpheres carboxylate-modified microspheres (1 µm) (blue fluorescent

365/415) were washed with 15 mM NaoAc, pH 5, three times; incubated with 5 mg/ml

BSA in 15 mM NaoAc, pH 5; and treated with 0.008 g 1-ethyl-3-(3- dimethylaminopropyl) carbodiimide hydrochloride and 0.1 N NaOH at 4°C overnight to covalently coat the beads with BSA. The following day, 2.5 M glycine was added for a

30-min incubation followed by two washes in 0.33× PBS, pH 7.4 120. Beads were washed twice with ice-cold buffer A (20 mM HEPES, pH 7.5, 50 mM NaCl, 1 mM NiCl2), suspended in ice-cold buffer B (20 mM HEPES, pH 7.5, 50 mM NaCl) and 5 μg LLO (or none) to adsorb LLO to the surface, and incubated on ice for 10 min. Two washes with buffer B removed excess LLO, and beads were diluted into ice-cold MEM to obtain MOI

5 or 20. Beads were added to mammalian cells in a six- or 24-well plate format followed by centrifugation at 520 × g for 3 min at room temperature and incubated for 30 min at

37˚C. Cells were washed with MEM twice followed by two washes with PBS and fixed with 4% paraformaldehyde (PFA) in PBS, pH 7.4, for 15 min at room temperature. Cells were washed three times with 0.1 M glycine in PBS, pH 7.4, and blocked in 0.1 M glycine in PBS/5% milk for 1 h. Extracellular beads were labeled using an anti-BSA rabbit antiserum followed by a secondary goat anti-rabbit IgG antibody conjugated to

Alexa-488. To quantify the number of intracellular beads, 40 sets of images (DAPI,

Alexa-488, and phase-contrast) were randomly acquired using a Metamorph programmed, motorized stage, and the 20× objective. Metamorph imaging and analysis software was used to determine the number of extracellular (Alexa-488, Ne) and total

68

(DAPI, Nt) beads such that the efficiency of bead entry was measured as the percentage of intracellular beads (Ni) using the equation Ni = (Nt - Ne)*100/Nt.

Microscope equipment

A motorized, atmosphere-controlled inverted wide-field fluorescence microscope

(Axio Observer D1; Carl Zeiss) equipped with a PZ2000 XYZ Series Automated Nano stage (Applied Scientific Instruments); a TempModule S1, Heating Unit XL S, and

Objective heater 22.5/34 S1 (D) (Carl Zeiss); 20X Plan Neofluar, 40X Plan Neofluar, and

100× Plan Apo objective lenses (NA = 0.5, 1.3, 1.4, respectively); Cascade II:512 electron-multiplying charge-coupled device (EMCCD) camera (Photometrics); ORCA-

FLASH4.0 V2 Digital complementary metal-oxide-semiconductor camera (Hamamatsu); and a Smart Shutter (Sutter Instrument) on the LED lamp. A Lambda 10-3 optical emission filter changer and DG-4 ultra-high-speed wavelength switching illumination system with a 300-W Xenon arc bulb (Sutter Instrument Company) equipped with excitation filters for CFP and FRET (S430/25x) and for YFP (S500/20x), emission filters for CFP (S470/30) and for YFP and FRET (S535/30), and a JP4 beamsplitter for CFP and

YFP (86002v2-spr) from Chroma Technology, were controlled with Metamorph

Microscopy Automation and Image Analysis Software (Molecular Devices).

FRET stoichiometry imaging

Fluorescence resonance energy transfer or FRET is a biochemical technique that assesses the physical proximity of fluorophores at distances ≤10 nm. On excitation, a donor fluorophore transfers its energy in a nonradiative manner to a lower energy

69 acceptor fluorophore leading to acceptor fluorescence. FRET stoichiometry utilizes donor emission at donor excitation wavelength (ID), acceptor emission at acceptor excitation wavelength (IA), and acceptor emission at donor excitation wavelength (IF) to quantify

FRET. FRET stoichiometry relies on four predetermined coefficients (α, β, γ, and ξ) that are characteristic of the microscope settings and fluorophores used 162, 163. The coefficients were determined using HepG2 cells expressing mCit alone (α = 0.072), mCFP alone (β = 0.584) and mCit-mCFP tandem (γ = 0.058 and ξ = 0.0156).

Phase-contrast (PC), ID, IA, and IF images were acquired with the Cascade II:512

EMCCD camera (Photometrics) at 200-ms exposure every 20 s over 20 min, in which no photobleaching was observed. Images were corrected for camera dark noise and adjusted with a binary mask, and FRET stoichiometry was applied to determine, pixel by pixel, EA and ED using the Metamorph software. The apparent FRET efficiency EA is proportional to the fraction of mCit in complex with mCFP, while EDis proportional to the fraction of mCFP in complex with mCit 163. The average apparent FRET efficiency, EAVE= (EA+ED)/2, corrects for variations in donor and acceptor expression levels and therefore normalizes FRET signals among different transfected cells 191.

FRET stoichiometry measurements between mCit-Rac1 and mCFP-PBD

HepG2 cells cotransfected to express mCit-Rac1 and mCFP-PBD, or the mentioned chimeras, were imaged for PC, ID, IA, and IF under 40× magnification. Cells were imaged in microscopy cell culture medium (150 mM NaCl, 1 mM CaCl2, 1 mM

2+ MgCl2, 5 mM KCl, 20 mM HEPES, 10 mM D + glucose) or Ca -free microscopy cell

70 culture medium. FRET stoichiometry was applied to each image, pixel by pixel, using ID, IA, and IF to create a calculated image corresponding to EAVE = (EA + ED)/2 or

FRET. As controls for Rac1 FRET analysis, we quantified the apparent FRET efficiency in cells that coexpressed mCFP-PBD plus the mCit fluorophore alone (FRET negative control), the native mCit-Rac1 (control for basal Rac1 activity), the dominant negative mCit-Rac1N17 (negative control for Rac1 activation), and the constitutively active mCit-

Rac1L61 (positive control for Rac1 activation). Additionally, cells separately expressing mCFP and mCit (FRET negative control), or the FRET pair mCit-mCFP tandem (FRET- positive control for the two fluorophores), were analyzed for FRET efficiency. Cells were placed on the microscope stage at 37˚C, and image series were collected every 20 s for 20 min (Supplemental Figure S1 and Supplemental Movie S1). In cells that coexpressed mCit-Rac1 and mCFP-PBD and exposed to LLO and/or inhibitors, FRET (EAVE) was calculated within two standard regions (305 pixels) that were positioned on each cell, at each time point. For each region, the average FRET value over the first 5 min of imaging

(before the addition of medium or LLO) was calculated to determine the baseline FRET within the analyzed region; all data points were then normalized relative to this value. In cells exposed to LLO and control nontreated cells, the regions were positioned where

Rac1 FRET variations were observed. If no FRET variation were observed, then the two regions were arbitrarily positioned at the cell periphery. A FRET-positive cell was determined as a cell that displayed at least a twofold increase in normalized FRET efficiency.

FRET measurements of the MyrPalm-CKAR biosensor 71

The C kinase activity reporter (CKAR) encoded in pcDNA3.1(+) is a chimeric protein containing an N-terminal mCFP fused to the Rad53p FHA2 domain, followed by an optimal substrate sequence for all PKC isoforms and a C-terminal mYFP (mCit). A

10-residue sequence from Lyn Kinase, which signals for myristoylation and palmitoylation, is located at the N-terminal end of the reporter resulting in the plasma membrane associated reporter MyrPalm-CKAR. In a nonphosphorylated state, the reporter exists in a clamped conformation resulting in FRET between mCFP and mCit.

On substrate sequence phosphorylation by a PKC, the FHA2 phosphopeptide domain binds the phosphorylated sequence resulting a conformational change moving mCit away from mCFP resulting in a loss of FRET. Importantly, this PKC biosensor has a dynamic range of 20% 169, 170. FRET stoichiometry was applied to each image, pixel by pixel, using HepG2 cells transfected to express MyrPalm-CKAR. Cells were washed once with microscopy cell culture medium (150 mM NaCl, 1 mM CaCl2, 1 mM MgCl2, 5 mM KCl,

20 mM HEPES, 10 mM d-glucose) and incubated in the same medium for 1 h at 37˚C prior to imaging. The ID, IA, and IF images were acquired with the Cascade II:512

EMCCD camera (Photometrics) at 200-ms exposure time every 20 s for 20 min under

100× magnification. FRET stoichiometry was applied to each image, pixel by pixel, using ID, IA, and IF to create a calculated image corresponding to EAVE = (EA + ED)/2 (or

FRET) and EAVE was averaged over the entire cell surface at each time point. The FRET values were normalized relative to EAVE averaged over the first 5 min before the addition of medium or LLO.

Membrane repair assay 72

HepG2 cells were seeded in a 96-well black-walled and clear bottom cell culture plate at 5 × 104 cells/well in 200 μl and incubated for 24 h. For Ca2+-containing conditions, cells were washed twice with 200 μl M1 (Hank’s balanced salt solution

[HBSS] supplemented with 0.5 mM MgCl2, 1.2 mM CaCl2, 10 mM HEPES, 25 mM glucose, pH 7.4) at 37˚C. For the Ca2+-free condition, cells were washed once with M2

(HBSS supplemented with 0.5 mM MgCl2, 10 mM HEPES, 25 mM glucose, pH 7.4) containing 5 mM EGTA and twice with M2 at 37˚C. M1 or M2 (100 µl) was added to the respective wells. Reagents were added to a separate 96-well plate maintained on ice: 100

μl of 120 μM PI in medium containing Ca2+ (M1) or not (M2), plus 100 μl of 4×- concentrated LLO (M1 or M2). The amount of 100 μl of medium containing the reagents were then transferred to the plate containing the cells. Additionally, as a positive control of plasma membrane damage, 0.1 and 1% Triton X-100 was added in place of LLO. As a negative control, M1 alone and M1 containing PI was added. HepG2 cells were exposed to 0.25, 0.5, and 1 nM LLO (final concentration) in M1 or M2. The kinetics assay was performed on a Spectra Max i3X Multi Mode Detection Platform (Molecular Devices) with the following parameters: 37˚C, fluorescence readings every 5 min for 30 min with an excitation/emission filter of 535/617 and 15/15 nm bandwidth 172.

Erythrocyte isolation and hemolysis assays

Human peripheral blood was obtained from healthy adult volunteer via venipuncture (Institutional Review Board protocol H0045). All donors provided a written informed consent. Erythrocytes were separated from plasma, leukocytes, and platelets by centrifugation on Polymorphprep (Axis-Shield) and erythrocytes were washed three 73 times with Alsever’s Solution (71.9 mM NaCl, 27 mM C6H5Na3O7⋅2H2O, 2.6 mM

C6H5Na3O7⋅H2O, 113.8 d-glucose).

Hemolysis assay (end point).

Erythrocytes (5 × 107 cells/ml) were suspended in PBS or in PBS containing 4 mM DTT. LLO or LLOpL were serially diluted in PBS or PBS containing 4 mM DTT in

10 μl in a 96-well cell culture plate, and 150 μl of erythrocytes in the respective media was added. Erythrocytes were incubated at 37˚C for 30 min and then centrifuged at 343

× g for 5 min. Supernatant (100 µl) from each well was transferred to a new 96-well cell culture plate, and OD = 540 nm was measured using the Spectra Max i3X Multi Mode

Detection Platform (Molecular Devices).

Hemolysis assay (kinetics).

Erythrocytes were suspended in ice-cold PBS containing DMSO vehicle control,

1 mM cytochalasin D, 1 mM GF109203x, or 150 μM CK-666 to a concentration of 5 ×

107 cells/ml. LLO (10 nM) in PBS containing DMSO vehicle control, 1 mM cytochalasin

D, 1 mM GF109203x, or 150 μM CK-666 was distributed in 10-μl aliquots in a 96-well cell culture plate that was placed on ice. Erythrocytes (150 μl) were added, and the plate was placed in the Spectra Max i3X Multi Mode Detection Platform (Molecular Devices).

The OD = 700 nm was measured every min for 30 min at 37˚C.

Statistics

In the entire article, “n” refers to the number of independent experiments. Statistical analyses were performed as follows. 74

Invasion assays

A linear mixed effects model was used to take account of the correlation among observations obtained from the same day and Holm’s procedure was used to control for multiple comparisons.

Rac1 FRET and MyrPalm-CKAR FRET.

The area under the FRET curve (AUC) between time points T0 (LLO addition),

2+ and T15 min (end of the movies) in cell culture with Ca , or T0 and T5 in cell culture without Ca2+, was calculated and analyzed using the Student’s t test for unequal variances and analysis of variance, respectively (Tables B.2 and B.3 for Rac1 FRET and Tables B.4 and B.5 for MyrPalm CKAR FRET). When comparing the percentage of FRET-positive cells for the Rac1 FRET (Tables B.2 and B.3), linear mixed effects models were used to account for repeated experiments. Holm’s procedure was used to adjust the p values 172 for all the above multiple comparisons. SAS9.4 was used for the analyses (SAS Institute).

Membrane repair.

Propidium iodide (PI) intensity values were first background corrected, log10- transformed, and averaged for a given condition for each time point for each experiment.

Negative values were set to 0 after transformations. The fixed effects of conditions with

DMSO or the inhibitors GF109203x, CK-666, and cytochalasin D, time (as a continuous variable), and LLO concentration (as a categorical variable) were tested with a linear mixed model to take into account the correlation among observations from the same independent replicates using the average log-transformed intensity values. A pairwise

75 comparison was made between the different LLO concentrations (0, 0.25, 0.5, and 1 nM) and their differences on log10 (intensity), and the same analysis was performed on the different inhibitor conditions (DMSO, 1 μM cytochalasin D, 1 μM GF109203x, and 150

μM CK-666) (Table B.5). Holm’s procedure was used to adjust the p values 172 for all above multiple comparisons. SAS9.4 was used for the analyses (SAS Institute).

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3 Chapter 3: A High-Throughput Assay to Quantify Membrane Repair Efficiency

3.1 Introduction

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in the plasma membrane damage. In response to these damages, the plasma membrane is rapidly resealed by complex molecular machineries to restore its barrier function and maintain cell survival. Despite

60 years of research in this field, we still lack a thorough understanding of the cell resealing machineries. With the goal of identifying cellular components that control plasma membrane resealing or drugs that can improve resealing, we developed a fluorescence-based high-throughput assay that measures the plasma membrane resealing efficiency in mammalian cells cultured in microplates. As a model system for plasma membrane damage, cells are exposed to the bacterial pore-forming toxin listeriolysin O

(LLO), which forms large 30-50 nm diameter proteinaceous pores in cholesterol- containing membranes. The use of a temperature-controlled multi-mode microplate reader allows for the rapid and the sensitive spectrofluorometric measurements in combination with bright field and fluorescence microscopy imaging of living cells.

Kinetic analysis of the fluorescence intensity emitted by a membrane impermeant nucleic acid-binding fluorochrome reflects the extent of membrane wounding and resealing at the cell population level, allowing for the calculation of the cell resealing efficiency.

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Microscopy imaging allows for enumeration of cells in each well of the microplate to account for potential variations in their number and allows for eventual identification of distinct cell populations. This high-throughput assay is a powerful tool expected to expand our understanding of membrane repair mechanisms via screening for host genes or exogenously added compounds that control plasma membrane resealing.

Mammalian cells are subject to mechanical, osmotic, and biochemical stresses resulting in the loss of plasma membrane integrity. Without rapid and efficient resealing, damaged cells would quickly succumb to programmed or necrotic death. Since the 1960s, efforts to understand plasma membrane resealing processes have been motivated by the devastating consequences associated with its dysfunctions. Indeed, diseases such as Limb-

Girdle Muscular Dystrophy, diabetes, and Chediak-Higashi Syndrome have been linked to deficient plasma membrane repair due to mutations in the gene encoding dysferlin, to production of advanced glycation end products, and to defects in the lysosomal trafficking regulator CHS1, respectively 192-197. However, to date our understanding of membrane resealing is still limited 63. Initial studies demonstrated that membrane resealing is initiated by the influx of extracellular Ca2+ through the damaged plasma membrane 2, 198, 199. Since then, several non-mutually exclusive Ca2+-dependent mechanisms have been proposed to reseal the plasma membrane of mammalian cells. The patch hypothesis proposes that in proximity to the wound, intracellular vesicles fuse with each other and with the damaged plasma membrane to act as a patch 200-203. A second model proposes that the calcium- dependent exocytosis of lysosomes at the wound site releases the lysosomal enzyme acid

78 sphingomyelinase, which converts sphingomyelin to ceramide in the outer leaflet of the plasma membrane. This sudden change in lipid composition results in ceramide-driven endocytosis of the damaged region 51, 52, 66. Lastly, the third major mechanism involves a role for the endosomal sorting complex required for transport (ESCRT) to promote the formation of outward-facing vesicles that bud off from the plasma membrane 90. Only a limited set of proteins were identified in these models and their machineries need to be further elucidated.

Here we describe a high-throughput assay that measures the plasma membrane resealing efficiency in adherent mammalian cells subjected to damage mediated by recombinant listeriolysin O (LLO) 172. LLO is a pore-forming toxin (PFT) secreted by the facultative intracellular pathogen Listeria monocytogenes 117, 118, 181 and belongs to the

MACPF/CDC (membrane attack complex, perforin, and cholesterol-dependent cytolysin) superfamily. The MACPF family constitutes mammalian pore forming proteins while

CDCs are bacterial pore-forming toxins mainly produced by Gram-positive pathogens 39.

CDCs are synthesized as water-soluble monomers or dimers that bind to cholesterol present in the plasma membrane and oligomerize into a prepore complex of up to 50 subunits that then undergo a conformational change to insert β-strands across the lipid bilayer forming a β-barrel pore spanning 30-50 nm in diameter 44, 204-206. These pores permit fluxes of ions and small cellular components in and out of the cell, though some studies proposed that smaller pores are also formed 207-209. Among the CDC toxins, LLO displays unique properties including its irreversible pH- and temperature-dependent aggregation 119, 210.

79

These characteristics are conducive to high-throughput assays as LLO can be added to the cell culture medium at 4˚C, a temperature permissive to its binding to cells, but not to formation of the pore complex. Initiation of pore formation can then be synchronized by raising the temperature to 37˚C, allowing for the efficient diffusion of toxin molecules in the plane of the membrane and formation of oligomers that then rearrange into pore complexes. The time necessary to form pores and the degree of cell damage will therefore depend on the amount of toxin bound to the plasma membrane. Furthermore, unbound, soluble LLO rapidly and irreversibly aggregates at 37˚C, which alleviates the need to wash away the unbound toxin and limits the extent of membrane damage over time. Lastly, because LLO binds to cholesterol and forms pores in cholesterol-rich membranes, this assay is amenable to a wide range of mammalian cells. It is important to keep in mind that

LLO affects host cell signaling mainly via pore formation, with a few exceptions in which pore-independent cell responses linked to the immune response can occur 211-217. Therefore, latter signaling activities may influence membrane repair.

This assay directly assesses cell wounding by measuring the incorporation of a cell impermeant fluorochrome (propidium iodide or TO-PRO-3) that passively enters wounded cells and becomes fluorescent only once it associates with nucleic acids. Hence the fluorochrome can be maintained in the cell culture medium throughout the experiment, allowing real-time analyses of cell wounding. The fluorescence intensity of the nucleic acid dye will increase with the concentration of toxin and, for a given concentration of toxin, will increase overtime until all pores are formed and cells are fully repaired or until

80 saturation is reached. The influx of extracellular Ca2+ through membrane pores is a key signal necessary for resealing. Therefore, the plasma membrane resealing efficiency can be indirectly evidenced by measuring cell wounding in culture medium containing Ca2+ versus Ca2+-free medium, for any given experimental condition allowing for the calculation of a repair efficiency coefficient (REFF). Because the fluorescence intensity of the nucleic acid-binding dye is also directly related to the cell concentration in each well, it is important to seed cells at the same concentration in all wells. It is also important to enumerate cells in each well before and after the assay to ensure that no cell detachment had occurred, which would complicate data interpretation. To enumerate cells in each well, cells expressing nuclear-localized histone 2B-GFP (H2B-GFP) were used in this assay.

Temperature-controlled, multi-mode, microplate readers combine rapid high-throughput analytics (using a 96 or 384-well plate format) of rapid fluorometric measurements of live cells at 37˚C with microscopy imaging. The latter can be used to enumerate cell number and observe eventual formation of distinct cell populations.

Ultimately, this assay is a powerful tool that provides users the ability to expand our knowledge in the complexity of membrane repair and to screen for host molecules or exogenously added compounds that can modulate membrane repair. The following protocol describes the necessary conditions to evaluate the effect of a drug or siRNA treatment on membrane resealing relative to the control using the following conditions:

Ca2+-containing medium (repair permissive) ± LLO and Ca2+-deplete medium (repair- restrictive) ± LLO.

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Abbreviations used: H2B – histone 2B TL – transmitted light PI – propidium iodide GFP – green fluorescent protein TP3 – TO-PRO-3 M1 – medium 1 (+Ca2+) M2 – medium 2 (no Ca2+) E – efficiency REFF – efficiency ratio

3.2 Protocol

1. Preparation

1.1 Cell Plating

NOTE: Human cervical epithelial cells, HeLa and HeLa expressing Histone 2B-GFP

(H2B-GFP), were used in this protocol, but this assay can be adapted to other mammalian cells 172.

1.1.1 Detach adherent cells from a 75 cm2 cell culture flask by washing the cells with 2 ml of Trypsin-EDTA 0.25% and replace the used trypsin with 2 ml of fresh Trypsin-EDTA

0.25%.

1.1.2 Incubate the cells at 37˚C for 5 min until the cells have rounded and detached from the flask.

1.1.3 Resuspend the cells in 8 ml of growth medium (DMEM containing 10% heat- inactivated fetal bovine serum, 100 U/ml penicillin, and 100 µg/ml streptomycin).

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1.1.4 Determine the cell concentration using a hemocytometer and 10 µl of cell suspension.

1.1.5 Dilute the cells in growth medium to a concentration of 2.5 x 105 cells/ml.

1.1.6 Pour the cell suspension into a sterile pipette basin and thoroughly mix the suspension using a 10 ml serological pipette.

1.1.7 Using a 12-multichannel micropipette and 200 µl tips, distribute HeLa cells (2.5 x

104 cells/100 µl/well) in triplicate (or quadruplicate) in a 96-well flat, clear bottom, black polystyrene tissue culture-treated plate.

Note: A plating arrangement is presented as an example in Figure 3.1.

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Figure 3.1 Experimental Design The flow diagram depicts a representative plate design configured to test the effect of seven test conditions in comparison to control non-treated cells. Additional controls should be included if appropriate, as for example drug vehicles. Cells are plated (plate 1) 24 h prior to the experiment. On the day of the experiment, cells in plate 1 are washed 84 with M1 or M2 medium pre-warmed at 37˚C and the plate is imaged (TL, GFP and PI fluorescence) pre-kinetic. During the 15 min of imaging, reagents are added on ice to plate 2. After imaging, plate 1 is immediately placed on ice for 5 min and 100 μl/well are then transferred from plate 2 to plate 1. Plate 1 is placed in the plate reader to run the kinetic assay at 37˚C for 30 min, followed by imaging (TL, GFP and PI fluorescence). Data are then analyzed to count cells and assess the repair efficiency in all experimental conditions. In large data sets, analysis can be automated. Also, the number of technical replicates can be increased to 4 in high-throughput screens.

1.1.8 Culture cells for 24 h in a humidified cell culture incubator at 37˚C and 5% CO2.

1.2 Stock solution preparation

1.2.1 Prepare 1 l of a 10x stock of buffer M (used to prepare M1 and M2) by adding 95 g of Hanks Balanced Salt Solution, 0.476 g MgCl2 (5 mM), and 23.83 g HEPES (100 mM) to 900 mL of water, adjust the pH to 7.4, and raise the volume to 1 l. Filter sterilize.

1.2.2 Prepare 50 ml of a 50x (1.25 M) stock of glucose by adding 11.26 g of D-(+)-

Glucose to a total of 50 ml of water. Filter sterilize.

1.2.3 Prepare 50 mL of a 100x (120 mM) stock of calcium by adding 0.666 g of CaCl2 to a total of 50 ml of water. Filter sterilize.

1.2.4 Prepare 50 ml of a 10x (50 mM) stock of ethylene glycol-bis(2-aminoethylether)-

N,N,N’,N’,tetraacetic acid (EGTA) by adding 0.951 g of EGTA to 40 ml water. Increase the pH to 8 using NaOH to dissolve the EGTA and then raise the volume to 50 ml. Filter 85

Sterilize.

1.2.5 For a single 96-well plate, prepare 50 ml of Medium 1 (M1, contains Ca2+), 50 ml of Medium 2 (M2, Ca2+-free), and 15 ml of Medium 2 supplemented with EGTA accordingly:

1.2.5.1 For M1, add 5 ml of 10x Buffer M, 0.5 ml of 100x CaCl2, and 1 ml of 50x glucose to 43.5 ml of water.

1.2.5.2 For M2, add 5 ml of 10x Buffer M and 1 ml of 50x glucose to 44 ml of water.

1.2.5.3 For M2/EGTA, add 1.5 ml of 10x Buffer M and 1.5 ml of 10x EGTA to 12 ml of water.

Note: All solutions containing propidium iodide (PI) should be prepared right before adding to cells.

1.3 Plate reader/Imaging cytometer settings.

NOTE: Use a multi-mode plate reader equipped with two detection units: a spectrofluorometer and an imaging cytometer. Limit the fluorescence exposure to avoid photobleaching the fluorophores.

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1.3.1 Pre-warm the plate reader to 37˚C before performing the assay.

1.3.2 Setup the parameters for the kinetic assay accordingly within the Settings mode:

1.3.2.1 Choose Monochromator, FL (fluorescence), and Kinetic for the Optical configuration, Read modes, and Read type, respectively.

1.3.2.2 Under Wavelength Settings, select a 9 and 15 nm excitation and emission bandpass, respectively. For assays using propidium iodide (PI), set the excitation and emission wavelengths to 535 and 617 nm, respectively.

1.3.2.3 Under Plate Type, select 96 Wells for the plate format and a pre-set plate configuration corresponding to a black-wall clear bottom plate.

1.3.2.4 Under Read Area, highlight the wells that will be analyzed throughout the kinetic.

1.3.2.5 Under PMT and Optics, preset the Flashes per read to “6” and check the box for

“read from bottom”.

1.3.2.6 Under Timing, insert 00:30:00 in the Total Run Time box for a 30 min kinetic assay, and insert 00:05:00 for the Interval. For each time point and one wavelength, reading time of a full 96-well plate is 30 s.

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1.3.2.7 Confirm the specified settings in the Settings Information to the right, and select

OK. Press Read to initiate the kinetic run.

1.3.3 Setup the imaging parameters accordingly within the Settings mode:

1.3.3.1 Choose Minimax, Imaging, and Endpoint for the optical configuration, read modes, and read type, respectively.

1.3.3.2 Under Wavelengths, select transmitted light, and either or both of the fluorescence boxes corresponding to excitation and emission wavelengths of 456/541 nm (GFP) and

625/713 nm (PI).

1.3.3.3 Use the same options for the Plate Type and Read Area as defined in steps 1.3.2.3 and 1.3.2.4.

1.3.3.4 Under Well Area Setting, select the number of sites within a well to be imaged. 12 sites corresponds to a full-well image.

1.3.3.5 Under the Image Acquisition Settings, select the exposure times for transmitted light, 541 (GFP), and 713 (PI). For GFP, image the entire well with an exposure time of 20 ms/image. For transmitted light (TL) and PI fluorescence acquire a single image of the

88 center of each well with exposure times of 8 and 20 ms, respectively.

1.3.3.6 Confirm the specified settings in the Settings Information to the right, and select

OK. The acquisition time for imaging the entire surface of each well (12 images/well) of a

96-well plate and for one wavelength is ~15 min/plate. Press Read to initiate imaging.

NOTE: The acquisition time of a single image/well of a 96-well plate requires ~2.5 min/plate for one wavelength. The Parameters described above correspond to the specific equipment of our laboratory. Spectrofluorometric measurements: A xenon flash lamp displaying 1.0 nm increment excitation wavelengths (250-850 nm) with an adjustable 9 or 15 nm bandpass, a photomultiplier tube detector with a >6 log dynamic range and an adjustable 15 or 25 nm emission bandpass. Imaging cytometer: An illumination light source capable of white light, 460 nm and 625 nm excitation wavelengths with a 20 nm bandpass, emission filters centered at 541 nm (108 nm bandpass) and 713 nm (123 nm bandpass), respectively, and a 4X objective coupled to a 1.25 megapixel 12-bit charge- coupled device camera.

2. Assay

Note: At the time of the assay, cells must be 70-90% confluent. During the wash steps, the medium should be removed from and applied to the side-wall of the well, not directly above the cells. Maintain the temperature of LLO <4˚C to prevent its aggregation until step 3.1.5.

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2.1.1 Prepare a stock of 30 µM PI in M1 and a stock of 30 µM PI in M2 pre-warmed at

37˚C.

2.1.2 Gently wash cells in plate 1 using a 12-multichannel micropipette and 200 µl tips, as follows:

2.1.2.1 For repair-permissive conditions, remove the growth medium and wash the cells twice with 200 µl/well M1 pre-warmed at 37 ˚C. Replace the medium with 100 µl/well of warm M1 containing 30 µM PI.

2.1.2.2 For repair restrictive-conditions, remove the growth medium and wash the cells once with 200 µl/well warm M2 containing 5 mM EGTA to chelate Ca2+, followed by one wash with 200 µl/well M2. Replace the medium with 100 µl/well warm M2 containing 30

µM PI.

2.1.2.3 After the growth medium has been washed and replaced with medium containing propidium iodide, directly move to step 2.1.3.

2.1.3 Image plate 1 under transmitted light, GFP and PI as detailed under 1.3.3 (pre- kinetic). This step takes 15-20 min.

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2.1.4 During the 15 min of step 2.1.3 prepare plate 2 using a 12-multichannel micropipette and 200 µl tips as follows:

2.1.4.1 Place a 96-well round bottom polypropylene microplate on ice. Configure the plate using an experimental design corresponding to plate 1 (Figure 3.1).

2.1.4.2 For repair-permissive conditions, add 100 µl/well of ice-cold M1 containing 60 µM

PI, followed by the addition of 100 µl/well of ice-cold M1 containing or not 4X LLO.

2.1.4.3 For repair-restrictive conditions, add 100 µl/well of ice-cold M2 containing 60 µM

PI, followed by the addition of 100 µl/well of ice-cold M2 containing or not 4X LLO.

2.1.5 After imaging plate 1 (step 2.1.3), immediately place plate 1 on ice using aluminum foil to separate the plate from direct contact with ice. Allow plate 1 to cool down for 5 min.

2.1.6 Using a 12-multichannel micropipette and 200 µl tips, transfer 100 µl from each well in plate 2 (step 3.1.1) to the corresponding wells in plate 1. To properly distribute the toxin in the media, insert the tips below this meniscus and gently eject the volume without introducing bubbles.

Note: Do not pipette up and down as this may inadvertently detach the cells.

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2.1.7 Leave the plate for an additional 1 min to allow the toxin to bind to host cells and immediately transfer plate 1 to the plate reader for the kinetic assay using the spectrofluorometer mode (step 1.3.2).

2.1.8 At the end of the kinetic assay, immediately image plate 1 (post-kinetic) using step

1.3.3.

3 Analysis: Cell enumeration

3.1. Determine the cell count based on the nuclear fluorescence using the microplate cell enumeration software.

3.1.1 Within “Settings” select “re-analysis”, and under the category section within the

“Image Analysis Settings” select “Discreet Object Analysis” using 541 as the wavelength for finding objects.

3.1.2 Within the Find Objects option, using the “draw on images” finding method, select

“Nuclei” under the settings tab, and press apply,

3.1.3 Press “OK” and the “read” button to initiate the cell counting algorithm.

3. Alternatively, if no such tool is available, use an image analysis software such as

ImageJ to enumerate cells.

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3.2.1 In ImageJ, open the image file as a stack.

3.2.2 Convert the stack to 8-bit greyscale images by clicking "Image” in the menu bar, hover over “Type” and select “8-bit”.

3.2.3 Subtract the background: Click “Image” in the menu bar, hover over “Adjust”, and select “Brightness/Contrast”. Adjust the minimum value to remove the background noise and select “Apply”.

3.2.4 Threshold to create binary images: Click “Image” in the menu bar, hover over

“Adjust”, and select “Threshold”. Select “Dark background”, adjust the minimum and maximum threshold values and click “Apply”.

3.2.5 In the case of overlapping nuclei, a “Watershed” tool can be used to segment nuclei.

Click “Process” in the menu, hover over “Binary” and select “Watershed”. This will automatically separate connected nuclei.

3.2.6 Analyze the masked images by applying user-specified criteria (size and circularity) to refine the identification of nuclei and to exclude cell debris.

3.2.6.1 Click on “Analyze” in the menu and click on “Analyze particles”. Set the desired

93 size (pixel^2) and circularity (a value of 1 is a perfect circle) ranges that are sufficient to include individual cells/nuclei.

3.2.6.2 In the “Show” dropdown box, select the option(s) desired, check “Summarize”, and click “OK” to obtain cell counts.

4. Analysis: Kinetic curves

4.1. Transfer the kinetic data from the plate reader software to an analytical data software.

4.2. For each experimental condition, average the fluorescence intensities of the replicates at each time point along with the corresponding standard deviation and standard error of the mean for each experimental condition.

4.3. For each experimental condition, trace the corresponding kinetic curve: PI intensity

(y-axis) versus time (x-axis).

4.4. To calculate the resealing efficiency of a given treatment condition, calculate the area under the curve (AUC) of the +LLO in M1 (AUC(M1)) and +LLO in M2 (AUC(M2)).

Use the approach suggested below to assess the efficiency (E) of resealing:

AUC(M1) AUC(M2) − AUC(M1) E = 1 − [ ] = AUC(M2) AUC(M2) 94

4.5. Perform a comparison between control and test treatment by determining the efficiency ratio (REff) indicated below:

Etest AUC(control, M2) AUC(test, M2) − AUC(test, M1) R = = ∗ [ ] EFF Econtrol AUC(test, M2) AUC(control, M2) − AUC(control, M1)

REFF = 1, test treatment has no effect on repair REFF < 1, test treatment inhibits repair REFF > 1, test treatment improves repair

4.6. Calculate the area under the curve using the following equation:

k−1 AUC = ∑i=1 (Intensityi+1 + Intensityi) × (Timei+1 − Timei)/2, where k is the total number of follow-ups.

3.3 Results

Cell Counting Accuracy

HeLa cells are frequently used as a model mammalian cell line to explore membrane repair mechanisms. When assessing membrane repair at the cell population level, it is important to plate cells at the same concentration in all wells for proper data interpretation. It is also important to verify at the time of the assay that cell numbers are equivalent across wells. HeLa cells that constitutively express histone 2B fused to GFP

(H2B-GFP) were introduced in this assay to automatically enumerate cells based upon detection of their fluorescent nuclei. To establish the accuracy in cell enumeration, twofold serial dilutions of HeLa H2B-GFP cells were plated in triplicate in a 96-well plate and cultured for 4 h. This time is sufficient for cell attachment and provides limited cell division. Full wells were imaged under transmitted light (TL) and GFP fluorescence 95 illuminations and cell counts were assessed based on GFP fluorescence using the plate reader analysis software (Figure. 3.2 A and B). The average cell counts ± standard deviation was plotted against cell seeding concentrations and a line of best fit indicated a

1.08:1 ratio of cell count to cell seeding, thus demonstrating the accuracy of the counting

(Figure. 3.2C). By imaging all of the wells prior to a kinetic assay one can ensure that cell numbers are consistent among all wells. Also, by imaging wells after the kinetic assay, one can establish if exposure to the toxin caused cell detachment. This information is critical for accurate interpretation of the kinetic data.

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Figure 3.2: Cell Counting accuracy HeLa cells expressing GFP-tagged Histone 2B were seeded in triplicates at the indicated concentrations. A. Cells were imaged at 37˚C under transmitted light (TL) and GFP fluorescence (12 images/well) and a cell detection algorithm was used to delineate individual nuclei (in purple). Scale bar = 1 mm. B. Higher magnification of TL, GFP, and cell detection (purple) images (zoomed 2X). Scale bar = 100 μm. C. Image analysis software was used to count the number of cells and the cell counts were plotted against the initial cell seeding concentration.

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Expression of GFP does not interfere with propidium iodide (PI) intensity measurements (IPI)

To ensure that H2B-GFP nuclear association does not interfere with PI incorporation or PI fluorescence intensity measurement (IPI), IPI was compared in HeLa and HeLa H2B-GFP cells that were exposed, or not, to 1 nM LLO (Figure. 3.3). In the absence of PI, there was a similar low level background fluorescence in HeLa and HeLa

H2B-GFP indicating that GFP does not bleed-through PI fluorescence emission filters. In the presence of PI, but in the absence of LLO, there was a similar basal PI fluorescence emission in both HeLa and HeLa H2B-GFP that did not change over time. This confirmed that expression of GFP does not affect measurement of PI fluorescence and also indicated that PI does not penetrate non-damaged cells over the time frame of the experiment. Addition of LLO resulted in an increase in PI fluorescence over time that was similar in both HeLa and HeLa H2B-GFP. This increase is due to cell wounding by

LLO combined with PI association with nucleic acids. Together, these results establish that expression of Histone-2B-GFP does not affect PI incorporation or measurement of its fluorescence.

Figure 3.3: Propidium iodide fluorescence measurement is not affected by Histone 2B- GFP expression 98

Histone 2B-GFP expressing and non-expressing HeLa cells were exposed, or not, to 1 nM LLO in the presence (solid lines) or absence (dashed lines) of 30 μM PI in Ca2+- containing medium (M1). The kinetic assay measured PI fluorescence intensities by spectrofluorometry every 5 min for 30 min at 37˚C. Data are the average PI fluorescence intensity (IPI) expressed in relative fluorescence units (RFU) ± SEM (n = 3 independent experiments, each performed in triplicates).

PI fluorescence does not interfere with GFP-based cell counting

Reciprocally, it was important to verify that PI nuclear incorporation in wounded cells does not interfere with GFP-based cell counting. Representative fluorescence images of HeLa and HeLa H2B-GFP exposed to 1 nM LLO in the presence of PI, showed that there was a marked accumulation of PI in wounded cells post kinetics, as expected (Figure. 3.4A). Imaging also revealed that PI could bleed-through GFP fluorescence detection (Figure. 3.4A and Table 3.1).

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Figure 3.4: Cell enumeration is unaffected by PI fluorescence. A. Representative pre- and post-kinetic images (TL, PI, and GFP) of cells exposed, or not, to 1 nM LLO in M1. Scale bar = 100 μm. B. Quantitative fluorescence microscopy analysis (IGFP ± SEM) revealed the increased GFP fluorescence measurement due to PI nuclear incorporation upon cell wounding by LLO (post-kinetic). C. GFP-based cell enumeration was unaffected by the increase in GFP intensity. Cell count per well was expressed as average ± SEM. (In B and C, black bars = pre-kinetic data, red bars = post- kinetic data. n = 3 independent experiments, each performed in triplicates. A two-tailed Student’s t-test was used to analyze quantitative fluorescence intensity and cell counts from acquired images **p<0.01)

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Imaging Cytometer Emission Channel Specifications Fluorophore Fluorophore Green channel Red channel ex/em (nm) (ex/em ± bandpass, nm) (ex/em ± bandpass, nm) GFP 488/510 PI 533/617 460/541 ± 20/108 625/713 ± 20/123 TO-PRO-3 642/661 Table 3.1 Fluorophore excitation/emission and imaging cytometer bandpasses. GFP, PI, and TO-PRO-3 excitation and emission peaks and the Green and Red channel excitation and emission bandpasses for the imaging cytometer.

This fluorescence crossover was best appreciated on the post-kinetic images of HeLa cells that do not express GFP, but still displayed green fluorescent nuclei (Figure 3.4A).

This crossover could also be evidenced by the measurement of GFP fluorescence intensity (IGFP) in HeLa H2B-GFP cells, which significantly increased post-kinetic in comparison to pre-kinetic (Figure 3.4B). Importantly, PI fluorescence crossover does not affect cell counting because the segmentation process involved in the enumeration of nuclei is unaffected by an increase in GFP fluorescence (Figure 3.4C).

Measuring plasma membrane resealing efficiency

In this section, we present the basic methodology used to measure the efficiency of membrane resealing. To evidence the process of membrane resealing, HeLa H2B-GFP cells were exposed, or not, to 1 nM LLO in the presence (M1) or absence (M2) of extracellular calcium (Figure 3.5). As expected, in the absence of LLO, IPI remained constant in M1 and M2. Addition of LLO in Ca2+-containing medium resulted in a steady increase in PI fluorescence intensity (IPI); whereas, in the absence of extracellular calcium

101 there was a significantly steeper increase in PI fluorescence, reflecting the absence of membrane resealing. To assess the resealing efficiency, which is defined as the capacity of cells to repair in M1 relative to M2 (Protocol 1.5.4.1), the area under the M1 and M2 curves (AUC) were determined and the efficiency of repair (E) was calculated to be

0.287.

Figure 3.5: Measuring plasma membrane resealing efficiency. Histone 2B-GFP expressing HeLa cells were exposed, or not, to 1 nM LLO in Ca2+- containing (M1) or Ca2+-free (M2) medium containing 30 μM PI. Kinetic data represent PI fluorescence intensity (IPI) in relative fluorescence units (RFU) ± SEM, measured for 30 min at 37˚C. n = 3 independent experiments, each performed in triplicate. The resealing efficiency was measured as indicated in protocol step 1.5.4.

An alternative to PI

PI has been ubiquitously used as a marker for plasma membrane damage.

However, there are other nucleic acid binding dyes that are also suitable for this assay.

For example, the membrane impermeant carbocyanine nucleic acid binding dye, TO- 102

PRO-3 (TP3), exhibits an emission spectrum reaching into the far red and has a high specificity for double stranded DNA. PI on the other hand binds both DNA and RNA 218,

219. Unlike PI, the excitation and emission spectra of TO-PRO-3 does not overlap with those of GFP allowing for better spectral resolution between the two fluorochromes.

Furthermore, TO-PRO-3 has an extinction coefficient nearly twice that of PI, which means that for their respective excitation wavelengths, this dye is more capable of absorbing energy than PI resulting in a stronger fluorescence emission. Quantitative fluorescence analysis of TO-PRO-3 and GFP images showed that this dye exhibits a large fluorescence dynamic range, does not significantly affect GFP fluorescence, and does not affect cell count (Figure 3.6A-D). Indeed, HeLa H2B-GFP cells incubated in M1 or M2 media containing TO-PRO-3 and damaged by 1 nM LLO exhibited a 4- and 5.5-fold increase in ITP3 relative to the non-damaged controls, respectively (Figure 3.7A). For comparison, cells exposed to 1 nM LLO in the presence of PI exhibited a 2.5- and 3-fold increase in PI fluorescence intensity in M1 and M2, respectively (Figure 3.5). Like PI,

TO-PRO-3 exhibits increasing fluorescence intensity with increasing LLO concentration in M1 (Figure 3.7A), and the resulting repair efficiency was calculated as described in protocol 1.5.4.1 (Figure 3.7B). We report that the cell resealing efficiency decreases as

LLO concentration increases. This phenomenon reflects the fact that cells decrease their capacity to reseal when excessive damages are caused.

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Figure 3.6: TO-PRO-3 as an alternative dye to assess membrane resealing. HeLa H2B-GFP cells were exposed, or not, to 0.5 nM LLO for 30 min at 37˚C in the presence of 1 μM TO-PRO-3 in Ca2+-containing (M1) or Ca2+-free (M2) medium. A. Images of HeLa H2B-GFP cells were acquired pre- and post-kinetic in M1 containing the dye. Scale bar = 100 μm. B and C. Integrated TO-PRO-3 and GFP fluorescence intensities were measured using the imaging cytometer and expressed in relative fluorescence units (RFU) ± SEM. D. GFP fluorescence images were processed to enumerate HeLa H2B-GFP cells (black bars = pre-kinetic data, red bars = post-kinetic data, n = 3 independent experiments, each performed in triplicates). A two-tailed Student’s t-test was used to analyze quantitative fluorescence intensity and cell counts from acquired images **p<0.01, ***p<0.001)

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Quality Assessment of the Membrane Repair Assay

A critical aspect of any assay is its robustness or capability to detect and resolve differences between the negative and positive controls. Signal variation between positive and negative controls must display sufficient dynamic range and reproducibility. In this membrane repair assay, the negative and positive controls are cells exposed to LLO in repair-permissive (M1) and repair-restrictive (M2) conditions, respectively. Two approaches were taken to assess the robustness of this assay for high-throughput analysis.

First, the Z-factor, or screening window coefficient, determines whether a given set of conditions provides a large enough dynamic range, while accounting for signal variability. Z-factors in the range 0

0.25 and 0.5 nM LLO produced Z-factor values of 0.3100813 and 0.137313, and β =

6.0672 and 4.803308, respectively, indicating that a window of LLO concentrations is suitable for the assay. As the concentration of LLO is increased beyond 1 nM, the gap in

105 the ITP3 kinetic curves between M1 and M2 conditions closes resulting in drastically reduced Z-factor and SSMD values (Figure 3.7B).

Figure 3.7: Effect of LLO concentration on the resealing efficiency, Z-factor, and SSMD. A. HeLa H2B-GFP cells incubated in M1 or M2 containing 1 μM TO-PRO-3 were exposed to increasing concentrations of LLO and subjected to the kinetic assay for 30 min at 37˚C. Data are expressed as TO-PRO-3 intensity (ITP3) in relative fluorescence units (RFU) ± SEM. B. The Z-factor and strictly standardized mean difference (SSMD) were calculated as a quality assessment for the robustness of the membrane repair assay, using the area under the curve (AUC) as a metric for the kinetic curves 220-222. The resealing efficiencies, were calculated as described in protocol and results sections (n = 3 independent experiments, each performed in triplicates).

Such high concentrations of LLO correspond to conditions in which the repair potential is outweighed by damage and thus negates the use of the assay in identifying factors

106 involved in membrane repair. This is further illustrated by the decrease in repair efficiency (E) as LLO concentration increases (Figure 3.7B). All data (Z-factor, SSMD, and E) were generated with 3 biological replicates and 3 technical replicates per experimental condition for assay validation. Together, these data show that this assay has the robustness expected for a high-throughput assay with LLO concentrations inferior to

1 nM for HeLa cells.

Proof of principle

Once the robustness of the assay was established, we performed additional experiments as a proof of principle that this assay has the sensitivity and resolution to identify a defect in repair. Also, we considered that high-throughput assays are used as a screening process to identify “hits” within large screens, which may involve less than 3 biological replicates. Thus it is pertinent that the experimental design provides the detection power to identify “hits” within a single assay. Therefore, under a high- throughput screen, the assay layout can be adjusted to accommodate 4 technical replicates to increase statistical power in a single biological replicate. Cells were plated in quadruplicate and were pre-treated 1 h prior to the assay with desipramine, a pharmacological inhibitor of the lysosomal protein acid sphingomyelinase (ASM), of which plays a role in plasma membrane repair 51, 66, 82. Importantly, treatment with desipramine did not affect cell counts throughout the assay allowing for the appropriate comparisons between desipramine-treated and non-treated cells (Figure 3.8A).

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Figure 3.8: Cell exposure to desipramine causes a defect in resealing. HeLa H2B-GFP cells (plated in quadruplicate) were pre-treated with 30 μM desipramine (or not) for 1 h at 37˚C and then exposed to 0.25 or 0.5 nM LLO in the presence of 1 μM TO-PRO-3 in Ca2+-containing (M1) or Ca2+-free (M2) medium. Cells were imaged (pre- and post-kinetic) and were subjected to the kinetic assay at 37˚C for 30 min. A. Cells were enumerated and data are expressed as average cell counts ± SEM. B. TO-PRO-3 fluorescence intensity (ITP3) is expressed in relative fluorescence units (RFU) ± SEM. C. Resealing efficiencies were calculated in the presence and absence of desipramine. A 108 mixed effects model was used on log-transformed intensity values assuming a random intercept for each technical replicate. To capture both a shift and change in shape of the kinetic curves, the main effect of treatment condition and the interaction effect between treatment condition and time were jointly tested for statistical significance. A two-tailed Student’s t-test was used to analyze cell counts from acquired images. The p-value was calculated using the mixed effects model. (4 technical replicates, one experiment).

Inhibition of ASM in desipramine-treated cells resulted in a defect in membrane resealing efficiency upon exposure to LLO, as illustrated by the decrease in E and REff (Figure 3.8B and 2.8C). Using a mixed effects model, a comparison of desipramine-treated to non- treated cells in M1 showed p-values of 0.0010 and 1x10-10, respectively. Together, data indicate that 0.25 and 0.5 nM LLO are appropriate concentrations to identify defects in repair in a high-throughput experimental setting with possible statistical analyses of a single experiment once the technical replicates are increased to four. Note that the statistical approach of the mixed effects model between quadruplicate will not account for potential variations across multiple biological replicates. Any significant findings using one biological replicate should be verified in additional experimental settings.

A high-throughput siRNA screen to identify proteins involved in plasma membrane repair

In order to identify proteins involved in plasma membrane repair, a siRNA-mediated knockdown approach was used. An initial optimization procedure was performed to determine the cell seeding concentration necessary to reach 70-90% confluence 72 h after 109 reverse transfection with 2 pmol of negative control siRNA in a 96-well format. In parallel, HeLa H2B-GFP cells were reverse transfected with 60 pmol of negative control sRNA, or Rac1-, RhoA-, Cdc42-, or CLTC (Clathrin heavy chain)-specific siRNA in a 6- well format to assess HeLa cells knockdown efficiency after 72 h. Cell seeding and amounts of reagents were increased proportionally to account for the ~30-fold increase in surface area from the 96- to the 6-well plate. A cell seeding of 7x103 cells/well resulted in

~6x104 cells/well (approximately 90% confluence) 72 h post-transfection in the 96-well plate. We achieved dramatic decreases in Rac1, RhoA, Cdc42, and CLTC expression as assessed by Western blot analysis (Figure 3.9). This is in accordance with the measured half-lives of Rac1, RhoA, Cdc42, and CLTC of 30, 31, 15, and 20 h, respectively 223-225.

For reference, 60% of the HeLa proteome exhibits a half-life of 20±5 h, while the entire

HeLa proteome exhibits a 50% turnover in ~24 h 226.

Lane 1 2 1 2 1 2 1 2

Rac1 RhoA Cdc42 CLTC Actin Actin Actin Actin

Figure 3.9 siRNA knockdown efficiency HeLa H2B-GFP cells were reverse transfected with negative control siRNA (lane 1) Rac1-, RhoA-, Cdc42-, and CLTC-specific siRNA (lane 2) for 72 hours. Cell lysates were obtained and subjected to Western Blot analysis. Actin was used as a loading control. N=1

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Rationale for selecting the library of siRNAs

siRNA targets were chosen based on the current models of plasma membrane repair mechanisms in order to identify intermediate proteins involved in the repair process. This involved identifying known signaling processes that branch out both upstream and downstream from the known mediators of plasma membrane repair. We also included most proteins know to play a role in endocytosis, exocytosis, intracellular trafficking, membrane fusion, and autophagy for potential discovery of novel repair mechanisms. As such, the library consists of 287 targets that play a role in exocytosis

(exocyst complex), endocytosis (clathrin, adaptor protein complex, dynamin, caveolin), vesicle trafficking (Autophagy, COP proteins, ESCRT machinery, Rab proteins), vesicle fusion (synaptotagmins and SNARE complex proteins), membrane stabilization

(Annexins, Flotillins, Septins), lysosomal function (Cathepsins, Acid sphingomyelinase), and cytoskeleton remodeling (Calpains, Caspases). Additionally, targets known to be involved in plasma membrane repair such as acid sphingomyelinase, annexins (A1, A2, and A6), and calpains were included to act as controls that would verify the capacity of the assay to identify defects in membrane repair.

Protocol

A high-throughput siRNA screen was performed to identify components of plasma membrane repair. HeLa H2B-GFP cells were reverse transfected with siRNA corresponding to 287 specific targets. Three 11-mer siRNAs per target were pooled in four wells (quadruplicates) and reverse transfected into HeLa H2B-GFP in order to

111 improve the efficacy of siRNA-mediated knockdown. HeLa H2B-GFP cells were treated with siRNA for 72 h and exposed to 0.5 nM LLO in the presence of 1 μM TO-PRO-3 in

Ca2+-containing (M1) and Ca2+-free (M2) media as described above (M1+LLO and M2

+LLO). In addition, each 96-well plate had 3 sets of control wells treated with negative control siRNA, with three seeding concentrations of 100%, 85%, and 70%, in order to account for potential variability in cell proliferation.

Results

The knockdown of proteins involved in membrane trafficking may have drastic effects on cell physiology including their ability to attach to the substratum. Also cell perforation by LLO could result in cell detachment, which events would interfere with data interpretation. To account for potential cell detachment pre- or post-exposure to

LLO, cells were imaged to allow their enumeration via image cytometry before and after the 30 min exposure to LLO (kinetic assay). Relative to the negative control at higher cell density (100% cell density), the knock down of 205 proteins resulted in a cell proliferation range of 65-105%, 28 proteins resulted in reduced cell proliferation (<65%), and 54 proteins resulted in increased proliferation (>105%), and one protein Cathepsin S, resulted in a dramatic 40% increase in proliferation (Figure 3.10). None of the siRNA treatment conditions exhibited cell detachment after exposure to LLO in the Ca2+- containing and in the Ca2+-free media. The average cell count ratio, defined as the cell count post-kinetic relative to the cell count pre-kinetic, across all siRNA treatment

112 conditions was 1.0061 ± 0.0065 and 1.0059 ± 0.0058 for M1 and M2 conditions, respectively (Figure 3.11A and B).

Figure 3.10 Cell proliferation. HeLa H2B-GFP cells were treated with pooled siRNA for 72 hours. Cell counts were determined by image cytometry and the average cell count ± standard deviation (n=4 technical replicates) was plotted relative to the 100% cell count negative siRNA control in increasing order.

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A. Cell count ratio of all siRNA treatment conditons in M1 1.2

1.1

1

0.9 Post:Pre Post:Pre Ratio

0.8

7 5 3

70 67 91 38 87 92 24 11

180 212 276 141 188 253 282 228 124 265 105 156 240 125 182 205 230 192 185 120 226 siRNA

B. Cell count ratio of all siRNA treatment conditions in M2 1.2

1.1

1

Post:Pre Post:Pre Ratio 0.9

0.8

7 5 3

70 67 91 38 87 92 24 11

180 212 276 141 188 253 282 228 124 265 105 156 240 125 182 205 230 192 185 120 226 siRNA

Figure 3.11 Cell count ratios. The cell count ratios ± standard deviation (n=4 technical replicates) were determined for all siRNA treatment conditions in M1 (A) and M2 (B).

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To identify siRNA treatment conditions that either positively or negatively affected membrane repair, the raw kinetic data was log transformed and the area under the curve (AUC) determined. From the four replicates per condition, no outliers were detected with the threshold of σ = ±1.5 i.e. variations no greater than 41% of the average.

The resulting AUCs were then compared relative to an approximated control using a best fit line between the 100%, 85%, and 70% control wells. Table 3.2 lists the significant hits

(Q<0.05) categorized by function. 49 out of 287 (17.07%) conditions were found to be significantly different from the controls, and of the 49 hits, 37/49 (75.51%) could be compared to an interpolated control (green). However, the remaining 12 exhibited cell counts <65% (10/12) or >105% (2/12) relative to the 100% control (red).

Table 3.2 Preliminary siRNA library hits The siRNA screen identified 49 proteins (Q<0.05) that when knocked down, inhibited (red, -) or improved (green, +) plasma membrane repair relative to the control. Conditions in which the cell counts are comparable to a control or interpolated control are highlighted in green while conditions with cell counts less than 65% or greater than 105% relative to the 100% control are highlighted in red. The list of 49 hits is categorized by function. From left to right: Gene name, Protein, cell count relative to the 100% control, knockdown effect on repair, Q value, and functionality. N/D = not determined.

Effect Relative on cell repair Gene Protein count (+/- ) Q Function

ANXA2 annexin A2 0.905434 - 0.048147 Plasma membrane

ANXA11 annexin A11 0.837058 - 0.014754 stabilization

adaptor-related protein complex Clathrin-mediated 1, sigma 1 endocytosis

AP1S1 subunit 1.0352 - 0.002259

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adaptor-related protein complex

AP2M1 2, mu 1 subunit 0.876 - 0.002259 adaptor-related protein complex 3, sigma 1

AP3S1 subunit 0.989 - 0.002259 adaptor-related protein complex 3, sigma 2

AP3S2 subunit 1.004 - 0.014754 adaptor-related protein complex

AP3M2 3, mu 2 subunit 1.15 - 0.046381 adaptor-related protein complex

AP4B1 4, beta 1 subunit 0.864517 - 0.027503 coatomer protein complex, subunit

COPG2 gamma 2 0.785 - 0.00742 coatomer protein complex, subunit

COPA alpha 0.0824 N/D 1.35E-08 coatomer protein complex, subunit

COPB1 beta 1 0.06903 N/D 1.35E-08 coatomer protein complex, subunit

COPG1 gamma 1 0.0867 N/D 0.00158 coatomer protein complex, subunit

COPZ1 zeta 1 0.1198 N/D 0.002259 clathrin, light

CLTB chain B 0.5413 + 0.012787

DNM2 dynamin 2 0.98027 + 0.006648

DNM3 dynamin 3 0.75336 - 0.022437

charged multivesicular Endosomal sorting

CHMP2A body protein 2A 0.7808 - 0.022437 complex required for chromatin transport modifying protein

CHMP4A 4A 0.8213 + 0.045144 116

chromatin modifying protein

CHMP6 6 0.64458 + 0.014754 vacuolar protein sorting 36 homolog (S.

VPS36 cerevisiae) 0.845404 + 0.014754 vesicle (multivesicular body) trafficking

VTA1 1 0.8919 + 0.032054

RAB2A, member RAS oncogene

RAB2A family 0.992846 + 0.04089 RAB4A, member RAS oncogene

RAB4A family 1.0702 + 0.006963 RAB7B, member RAS oncogene

RAB7B family 0.859883 + 0.014754 RAB30, member RAS oncogene

RAB30 family 0.917539 + 0.045144 Membrane transport RAB, member of and fusion RAS oncogene

RABL3 family-like 3 0.74782 - 0.015437 RAB9B, member RAS oncogene

RAB9B family 0.698374 +? 0.045058 RAB38, member RAS oncogene

RAB38 family 0.572099 - 0.019146 RAB13, member RAS oncogene

RAB13 family 0.7972 - 0.046381

synaptosomal- associated

SNAP25 protein, 25kDa 1.005 - 0.00079 Membrane fusion syntaxin 1A

STX1A (brain) 1.033 - 0.015569

STX4 syntaxin 4 0.90145 - 0.002259 117

STX5 syntaxin 5 0.8265 - 0.01504

STX12 syntaxin 12 0.7877 - 0.040635

STX19 syntaxin 19 0.920526 - 0.023366 syntaxin binding

STXBP1 protein 1 1.026 - 0.023725 syntaxin binding

STXBP3 protein 3 0.96156 - 0.03029 syntaxin binding

STXBP4 protein 4 0.591457 - 0.022437

SYT10 synaptotagmin X 0.751075 - 0.00742 synaptotagmin

SYT12 XII 0.8684 - 0.022035 synaptotagmin

SYT14 XIV 0.622293 - 0.00742

sphingomyelin phosphodiesterase Lysosomal protease

SMPD1 1, acid lysosomal 0.5285 - 0.023366 exocyst complex Exocytosis

EXOC6 component 6 1.146237 - 0.02636

heat shock 70kDa Protein folding

HSPA8 protein 8 0.673 + 0.04089

mannose-6- Anterograde phosphate trafficking Golgi to receptor binding Endosome/Lysosome

M6PRBP1 protein 1 1.153144 - 0.028155 prostaglandin E Tissue homeostasis

PTGES2 synthase 2 0.6988 - 0.00742 Sec23 homolog A

SEC23A (S. cerevisiae) 0.95 - 0.011708 SEC24 family, Anterograde member B (S. trafficking ER to

SEC24B cerevisiae) 0.958121 - 0.029911 Golgi lectin, mannose-

LMAN1 binding, 1 0.7354 + 5.30E-05

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3.4 Discussion

This assay measures the efficiency of membrane resealing at the cell population level with high-throughput capacity. It can be used to screen for cellular components or drug libraries that could affect membrane repair. The described assay used a 96-well plate format, but it can be adapted to 384-well plates for higher throughput. An advantage of this assay is its ability to obtain fluorescence measurements of adherent living cells in real time without the need for excessive cell processing such as cell detachment, fixation, or fluorescence labeling post-fixation. Multimode plate readers, such as the one used in this protocol, have sufficient sensitivity for rapid spectrofluorometric measurements at time intervals as low as 30 s (for a 96-well plate). The acquisition of fluorescence images provides additional information including cell enumeration and eventual changes in cell morphology, and allows for potential identification of distinct cell populations. The present assay does not claim to establish the kinetics of plasma membrane resealing per se, but to identify experimental conditions (pharmacological compounds or cellular components) that can affect the process of membrane resealing, either positively or negatively, within a high-throughput assay.

Several other experimental approaches have been developed to assess the membrane resealing mechanisms. For example, mechanical disruption by needle puncture, bead abrasion, and laser ablation have been used to model membrane damage due to mechanical stress. The measurement of wounding/repair has involved fluorescence microscopy or flow cytometry by quantifying the entry of fluorescent probes (FM 1-43,

119 propidium iodide, fluorescein-conjugated dextrans) or tracking fluorescent protein chimeras 73, 227-229. Each of these approaches has its own advantages; however, they are not amenable to high-throughput screening in living cells as presented in this assay.

The present assay was optimized to analyze the resealing efficiency of cells wounded by the pore-forming protein LLO, which forms large pores that permit the massive Ca2+influx provoked by mechanical ruptures of the plasma membrane. Although pore-forming toxins represent one form of plasma membrane damage, repair of large toxin pores and mechanical wounds were proposed to share common Ca2+-dependent pathways 66, 84. It is important to note that LLO interaction with cell membrane components such as cholesterol may affect cell signaling and thereby may influence membrane resealing mechanisms when compared to mechanical wounds. Our knowledge about membrane repair is still limited and further studies are required to establish if resealing of mechanical wounds differ from resealing following the formation of toxin pores. There are several advantages of using LLO. First, the initiation of membrane damage can be synchronized by raising the temperature from 4˚C to 37˚C thus allowing for multiple conditions to be tested at the same time. Second, the soluble form of LLO

(not bound to the cell membrane) irreversibly aggregates at neutral pH and 37˚C, thus limiting cytotoxic effects and abrogating the need to wash the cells. Finally, the degree of damage can be adjusted by varying the concentration of the toxin. However, a limitation of this assay is the temperature switch between 37˚C and 4˚C, which may affect the repair mechanism as vesicular transport and membrane fluidity, among other processes, are

120 influenced by temperature 230-232. It is important to verify that any pharmacological inhibitor included in the assay does not interfere with the formation of LLO pores by performing a hemolysis assay in the presence of the drug 233. Due to potential batch differences in LLO activity, it is important to prepare a stock of LLO that is large enough for the entire high-throughput screen 233.

We included the use of cells expressing the nuclear-localized Histone-2B-GFP chimera as a means to enumerate cells before and after the membrane repair assay via microscopic imaging followed by automated image analysis. Importantly, equivalent cell counts across conditions and unchanging cell counts before and after the kinetic assay are crucial as PI or TO-PRO-3 fluorescence intensities cannot simply be normalized. Indeed, a difference in cell count will result in differences in the degree of damage by a given concentration of LLO, which cannot be corrected for via fluorescence normalization due to variations in resealing efficiency (Figure 7). We showed that Histone-2B-GFP expression does not interfere with PI or TO-PRO-3 incorporation or fluorescence emission. Conversely, PI or TO-PRO-3 incorporation does not affect GFP-based cell enumeration. If other combinations of fluorophores are to be used, it would be necessary to assess potential spectral overlap between fluorochromes, as was performed in this work. Although PI has been extensively used to measure membrane damage, we show that TO-PRO-3 is an excellent substitute that exhibits higher fluorescence quantum yield resulting in a larger dynamic range suitable for characterizing resealing efficiency.

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Z-factor and SSMD confirmed that this assay has the robustness necessary to perform high-throughput analyses. The calculation of the resealing efficiency is a critical and reliable tool to identify potential “hits”. In addition, the mixed effects model can be used as a statistical tool to evaluate “hits” within a single assay. The experimental plan must include a minimum of three technical replicates if the screen can be repeated several times. Quadruplicates should be used if statistical tools, such as the mixed effects model, are to be included in a single experiment, whether or not it will be repeated. It is advised to perform the screen several times and it is required to validate the results by performing complementary experiments.

Plasma membrane repair is an absolutely necessary function for cell survival.

Efforts to decipher the molecular mechanisms involved in the repair of mechanically damaged membranes or membranes damaged by pore forming toxins have elucidated four mutually non-exclusive models, all of which require the influx of Ca2+. The initial spontaneous fusion model proposed that membranes with nanometer wide lesions could spontaneously fuse back together due to increased lipid disorder that drives the edges of the membrane back together to re-establish a thermodynamically favorable state.

However, this model was based on observations of red blood cells (RBCs) resealing after hypotonic shock in cold medium (~0˚C) with low ionic strength, 1/10th the ionic strength found in the physiological extracellular environment. This model was debunked as damaged RBCs in medium containing the physiological concentration of extracellular

Ca2+ (1.2 mM) failed to repair. Observations of vesicular exocytosis in X. laevis oocytes

122 and sea urchin eggs brought about the patch hypothesis in which exocytosing vesicles could fuse together and form a membranous patch that fuse with and seal the wound.

Indeed, this hypothesis was confirmed in X. laevis oocytes damaged via laser ablation, but left the question of whether this mechanism is relevant to mammalian cells. Indeed, mammalian cells exhibit vesicular exocytosis after wounding and the identity of these vesicles were later determined to be lysosomes. This brought about a new mechanism of membrane resealing in which epithelial cells damaged by the pore-forming toxin streptolysin O initiates lysosomal exocytosis and release of acid sphingomyelinase, which converts outer leaflet sphingomyelin to ceramide resulting ceramide-driven invagination of the plasma membrane and subsequent endocytosis of the wound. Conversely, a variety of cell types including epithelial, endothelial, adrenal, and neuronal cells exhibit vesicular shedding suggesting that damaged membranes may A) endocytose the damaged membrane, traffic that membrane to multivesicular bodies, but then the MVB refuses with the plasma membrane to release the intraluminal damaged vesicles (ectocytosis), or

B) the damaged membrane induces cellular swelling typically observed as blebs, that pinch off from the membrane releasing microvesicles containing the damaged region. All of these mechanisms evidence that a variety of cell types manage different forms of membrane damage in distinct manners. However, due to the lack of overlap in damage models and cell models, it is hard to elucidate or even connect these mechanisms of repair.

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We developed a high-throughput assay protocol that provides the capability to quantify plasma membrane resealing efficiency of a variety of adherent cell types exposed to the pore-forming toxin listeriolysin O. Unlike other models of membrane damage, we used LLO because of its tractability: useful in a high-throughput setting, degree of damage can be adjusted by changing the concentration, and can inflict damage on all cell types that contain cholesterol in their membrane. It is our hope that this assay will provide the platform for researchers to identify the myriad of proteins involved in plasma membrane in all cell types and thus fill in the gaps of the various repair models or even identify new mechanisms of repair. Using this approach, we performed a siRNA screen of 287 targets involved in membrane trafficking, fusion, and stabilization, endocytosis and exocytosis, autophagy, and lysosomal function. The initial screen identified 49/287 targets, that when knocked down, altered membrane resealing efficiency positively (17/49), or negatively (32/49). Confirmation studies are required, but this is the first time, to our knowledge, that a high-throughput assay was developed and used to study plasma membrane repair in real time.

Annexins

Annexins are Ca2+-sensitive membrane-associating proteins that facilitate vesicular trafficking and stabilization of the plasma membrane, especially under damaged-conditions. Annexins A1, A2, A5, and A6 are known to form a repair cap at the site of injury in muscle cells wounded by laser ablation 234, 235. Annexin A2 was also found to be involved in the repair of metastatic cancer cells 68. In accordance with these

124 findings, our screen identified annexin A2 as a positive regulator of plasma membrane repair. Additionally, we identified annexin A11 as a positive regulator of repair, though its function within the cell involves cell cycle progression and regulation of apoptosis 236.

If confirmed, this will be the first time annexin A11 has been implicated in plasma membrane repair.

Clathrin-mediated endocytosis

Cytosolic L. monocytogenes continuously secretes listeriolysin O, which actively perforates the host plasma membrane. And interestingly, cells heavily infected survive for days despite continuous plasma membrane damage. Recently, it was found that the host cell uses the clathrin adaptor protein 2 a2 subunit from the AP2 complex to mediate the endocytosis of LLO bound to the interior face of the plasma membrane via recognition of LLOs N-terminal PEST-like sequence 45. In fact, this screen identified adaptor protein (AP) subunits from four (AP1-4) of the five known AP complexes that mediate endocytic and secretory membrane trafficking. AP1-3 are involved in clathrin- mediated endocytosis and localize to the trans-golgi network/endosomes, plasma membrane, and endosomes, respectively. AP4 localizes to the trans-golgi network where it functions in anterograde transport of cargo 237. How plasma membrane perforation by

LLO from outside of the cell is recognized by the cytosolic adaptor protein complexes is yet to be determined. It is important though to be careful interpreting this data as knockdown of these proteins could drastically affect the composition of the plasma membrane and thereby alter the dynamics of LLO-mediated plasma membrane

125 perforation. Further work is necessary to verify the role of these adaptor proteins in plasma membrane repair.

Syntaxins and Synaptotagmins

Vesicular fusion events have been very well studied and characterized in the context of neuron biology in which pre-synaptic vesicles fuse with the plasma membrane to release neurotransmitters between neuronal axons 238. The mechanism whereby membrane fusion occurs requires the coordinated action of the plasma membrane- associated proteins SNAP-25 and synatxin-1 with the vesicle associated protein synaptobrevin, also known as VAMP 239. These proteins form the SNARE complex that zipper together to bring the vesicle membrane and plasma membrane within close proximity to induce the fusion of the two membranes. Our screen identified SNAP-25 and Syntaxin-1 as positive regulators of plasma membrane repair, along with syntaxins 4,

5, 12, and 19, syntaxin binding proteins 1, 3, and 4, and synaptotagmins 10, 12, and 14. It is not surprising that these proteins were found to be involved in plasma membrane repair as others have shown the involvement of synaptotagmin VII, SNAP-23, and syntaxin 4 involved in Ca2+-regulated exocytosis of lysosomes 12, 240.

These preliminary findings have confirmed some of the known proteins involved in plasma membrane repair and added a slew of previously unidentified proteins that act positively to promote resealing. Interestingly, knockdown of of the ESCRT and Rab proteins resulted in improved repair, which was unexpected. Identification of how these

126 proteins may be acting as negative regulators of repair will provide insight into the intricate signaling processes involved in the repair models. Again, these preliminary findings will need to be confirmed through a secondary screen with a complimentary analysis of the efficient knockdown of gene expression and decrease in protein under the given conditions. But, these findings provide a positive outlook for the characterization of the signaling processes that occur with respect to the existing models of repair

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Chapter 4: Future Directions and Final Comments

Pathogen-induced host plasma membrane damage represents a novel mechanism for the internalization of intracellular pathogens of diverse pathogens as is the case for

Trypanosoma cruzi, Listeria monocytogenes, and Adenovirus 241-244. In contrast to the

ASM-mediated plasma membrane repair process that facilitates the efficient entry of

Trypanosoma cruzi and Adenovirus into HeLa epithelial cells, perforation of hepatocyte epithelial cells by LLO induces a kinetically and mechanistically distinct Ca2+-influx dependent signaling pathway involving the activation of a conventional protein kinase C and the Rho GTPase Rac1 to induce Arp2/3-dependent F-actin assembly and Listeria monocytogenes invasion. In addition to these findings, it was previously illustrated that

K+ efflux coincident with Ca2+ influx is a prerequisite for the LLO-mediated internalization of Listeria 55. Indeed, HepG2 cells exposed to LLO in the presence of extracellular concentrations of K+ that equal K+ intracellular concentration, to prevent K+ efflux, failed to exhibit Rac1 activation (preliminary data). How K+ efflux affects this pathway remains to be determined.

Cellular homeostasis is dependent on the maintenance of cytosolic osmolyte (Na+, K+, Cl-

, and Ca2+) concentrations. An imbalance due to exposure to a hypotonic environment leads to cell swelling, which could lyse the cell if not for the regulated volume decrease

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(RVD) response 245-247. In fact cells exposed to high concentrations of LLO exhibit the formation of large plasma membrane blebs, which are slowly retracted over time, which is reminiscent of the RVD process 248, 249. Cellular swelling due to hypotonicity induces a rise in cytosolic Ca2+, which initiates RVD processes by activating Ca2+-dependent K+ channels to compensate for the increased cytosolic cations. To maintain electroneutrality, the volume-regulated anion channel (VRAC) is activated by binding of ATP and Ca2+ to induce the efflux of Cl- ions 250. Interestingly, it has been suggested that mitogen- activated protein kinase signaling (Ras-Raf-Mek-Erk) and Rho/Rho kinase signaling regulate that activation of VRAC 251, 252. In accordance with these findings, it was shown that LLO perforation of HT29 human colon epithelial cells resulted in a K+-efflux dependent activation of the MAPK p38, Erk1/2, Mek1/2, and Msk1/2 54. Based on these findings, we postulate that LLO-mediated perforation of the plasma membrane leads to an initial influx of Ca2+ and efflux of K+, which is halted by membrane repair processes, but leads to cellular swelling. The efflux of K+ induces an RVD response via MAPK signaling involving Rho kinases, which is integrated into the activation of Rac1. Future work will determine if LLO perforation of the plasma membrane facilitates the endocytosis of Listeria monocytogenes by integrating various cellular responses: membrane repair, PKC activation, and the regulated volume decrease response.

The preliminary siRNA screen identified 49/287 hits in which plasma membrane repair was inhibited (32/49), improved (13/49), or inconclusive (4/49). A second siRNA screen will be performed, which will aid in the confirmation of hits and potentially identify

129 targets that were missed in the first screen. This secondary screen will be adjusted such that conditions that have lower or higher cell counts after 72 h of siRNA treatment will be plated with control wells containing similar cell counts after 72 h such that comparisons can be made with interpolated data. Findings from these two screens will be further confirmed by a directed screen of the significant hits (Q<0.05) and submitted to Ingenuity

Pathway Analysis, which will identify links between the identified proteins and their potential role(s) in the various models of repair. As with all siRNA screens, the efficiency of knockdown comes into question. To assess the knockdown efficiency, HeLa H2B-GFP cells will be treated with siRNAs corresponding to the significant hits along with a set of randomly chosen siRNAs, and the cell lysates will be submitted for dot blot analysis. It is plausible that membrane repair signaling exhibits redundancy due to overlapping functions of similar proteins, as seen with the annexins 235. One approach to assess redundancy within a small family of proteins is to siRNA knockdown various combinations of genes and determine the effect on plasma membrane repair. Lastly, siRNA knockdown of a particular protein could lead to a myriad of downstream effects such as alterations to protein folding, post-translational modifications, and protein functionality and signaling, which could result in misleading conclusions. To assess the involvement of identified proteins in membrane repair, one approach would be to express fluorescently tagged proteins and use high-resolution confocal microscopy to visualize their localization to damaged regions of the plasma membrane (2 photon laser ablation or microneedle injection). Alternatively, FLAG-tag or Myc-tag proteins can be visualized

130 via immunofluorescence microscopy at different intervals after plasma membrane perforation.

It is likely that phenotypically and functionally different cell types exhibit hierarchal mechanisms of plasma membrane repair due to the unique expression profiles that provide different tolerances to plasma membrane damage. Myocytes frequently incur damage to the plasma membrane from repetitive contraction and flexion of the musculature. As such, these cells uniquely express MG53/TRIM72 to promote the trafficking and fusion of vesicles to efficiently reseal the damage region 253. Conversely, neuronal cells regularly undergo Ca2+ transients during potentiation and depolarization leading to the fusion of presynaptic vesicles and release of neurotransmitters into the synaptic cleft. Unique to neurons is the high expression of synaptotagmins that mediate vesicle fusion events 238, 254. Thus, plasma membrane damage to neurons may preferentially utilize a synaptotagmin-dependent pathway to repair the membrane, but could also be distinguished from a normal Ca2+ transient.

A future direction of this project would identify components of plasma membrane repair in other cell types including endothelial cells, myocytes, macrophages, lymphocytes, and neuronal cells and to characterize the mechanisms by which the different cell types have adapted to plasma membrane damage.

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In the context of infection by L. monocytogenes, infected cells can harbor large numbers of cytosolic bacteria which release LLO. Although multiple mechanisms have been shown to decrease LLO activity in the host cell cytosol, it is likely that these mechanisms are not sufficient to fully detoxify infected cells. As a consequence, some level of perforation still occurs. For example, it was proposed that intracellular LLO perforates the host cell plasma membrane of the forming protrusion to facilitate its capture by adjacent cells during cell-to-cell spreading of the bacterium. If this is correct, the constant repair of the plasma membrane of infected cells would be necessary to maintain cell survival and consequently for efficient cell-to-cell spread of bacteria. Once we confirm the identified set of proteins involved in the repair of infected cells, we will evaluate the role of membrane repair in the survival of infected cells and cell-to-cell spreading.

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Appendix A. Supplemental Data Chapter 2

Figure A.1 Rac1 FRET controls. HepG2 cells expressing mCit + mCFP (FRET negative control), mCit + mCFP-PBD (FRET negative control), mCit-Rac1N17 + mCFP-PBD (negative control for Rac1 activation), mCitRac1 + mCFP-PBD (basal Rac1 FRET activity), mCitRac1L61 + mCFP-PBD (positive control for Rac1 activation), or the tandem mCit-mCFP (positive control for mCit and mCFP FRET pair) were imaged on the microscope stage at 37˚C. Phasecontrast (PC), and fluorescence images (IA, ID, and IF) were acquired every 20 s for 20 min. (A) Representative FRET efficiency images presented in pseudo-color (Scale bar = 10 μm). EAVE was calculated over the entire cell surface and is expressed as the mean ± standard deviation by FRET stoichiometry. 13-35 cells were analyzed per experimental condition (n=1).

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Figure A.2 Hemolysis Assays. (A) End point analysis. Human erythrocytes were exposed to varying concentrations of LLO (0.2-100 nM) or LLOpL (0.1-100 nM and 27-14,000 nM) in the presence or absence of 4 mM dithiothreitol (DTT) for 30 min at 37˚C. Experiments involved technical triplicates and results are expressed as percent erythrocyte lysis ± S.E.M. relative to Triton X-100 lysed control cells, and PBS-incubated non-lysed control cells (n = 3). (B) Kinetic analysis. Human erythrocytes were treated with DMSO (negative control), Triton X-100 + DMSO (positive control), 1 μM GF109203x, 150 μM CK-666, 1 μM cytochalasin D (CD) and exposed to 0.2-100 nM LLO. The kinetics of lysis were assessed every min for 30 min at 37˚C. Results are expressed as percent lysis ± standard deviation relative to the DMSO control (n = 1).

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Figure A.3 Ca2+ influx triggers PKC activation at the plasma membrane. HepG2 cells expressing MyrPalm-CKAR were imaged on the microscope stage at 37˚C. Phase-contrast (PC) and fluorescence images (IA, ID, and IF) were acquired every 20 s for 20 min. EAVE was calculated over the entire cell surface and data are presented as the average fold change in EAVE. Medium, DMSO (0.1%) or Ionomycin (1 μM) were added after 5 min of imaging (T0). (11 cells in medium control, 7 cells in DMSO control, and 19 ionomycin-treated cells, n = 1).

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Figure A.4 LLO activates the Rho GTPase Cdc42. HepG2 cells co-expressing mCFP-PBD + mCit-Cdc42 were imaged on the microscope stage at 37˚C. Phase-contrast (PC), and fluorescence images (IA, ID, and IF) were acquired every 20 s for 20 min. LLO was added after 5 min of imaging (T0). The apparent FRET efficiency (FRET) was calculated. For each imaged cell, two regions of standard size were applied at locations where FRET variations were observed. If no FRET variation was observed, the regions were randomly positioned. (A and B) Data are expressed as the normalized apparent FRET efficiency. The y-axis indicates the fold-increase in FRET efficiency and the z-axis represents all regions analyzed. (A) 78 regions corresponding to 39 control cells, and (B) 84 regions, corresponding to 61 cells, that exhibited >2-fold increase in FRET upon exposure to LLO. (C) Summary of FRET data collected (n = 3). Time indicates the duration of imaging. NC corresponds to the number of analyzed cells. FRET+ cells correspond to the number of FRET positive cells. % FRET corresponds to the percentage of FRET positive cells.

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Non-treated NC Rac1 RhoA Cdc42 Rac1 0.75 1 0.16 1.15 1.19 Actin 0.97 1 1.09 1 1.21 Ratio Rac1:Actin 0.77 1 0.15 1.11 0.98

RhoA 0.83 1 1.08 0 1 Actin 1.04 1 1 1.02 1.2 Ratio RhoA:Actin 0.8 1 1.08 0 0.83

Cdc42 0.76 1 0.84 0.71 0.08 Actin 0.87 1 0.8 0.71 0.86 Ratio Cdc42:Actin 0.87 1 1.05 1 0.09 Table A.1 Densitometry analysis for Figure 3.1 B Densitometry analyses of Rho GTPases expression in siRNA-treated cells, corresponding to representative Western blots from Figure 1B (n=4).

Condition 1 Condition 2 LLO Time (0.1 Time LLO or Mean % Adjusted CaCl2 (min) nM) %FRET CaCl2 (min) LLOpL %FRET difference p-value + 20 - 2.43 + 20 LLO 76.61 -74.18 0.0005 LLOpL (0.1 + 20 + 76.61 + 20 nM) 5.88 70.74 0.0096 LLOpL + 20 + 76.61 + 20 (10 nM) 7.22 69.39 0.0096 LLOpL (0.1 + 20 - 2.43 + 20 nM) 5.88 -3.45 1.0000 LLOpL + 20 - 3.33 + 20 (10 nM) 7.22 -4.79 1.0000 + 10 - 0 + 10 LLO 74.83 -74.83 <0.0001 - 10 - 4.78 - 10 LLO 5.68 -0.91 0.861 + 10 - 0 - 10 - 4.78 -4.78 0.8056 + 10 + 74.83 - 10 LLO 5.68 69.14 <0.0001

Table A.2 Statistics for Table 3.1. Statistical analyses compared experimental conditions 1 to experimental conditions 2 and were performed on % FRET (n≥3). 152

Control LLO NC AUC NC AUC 91 13.323 91 13.323 95% confidence interval 11.753-14.894 17.608-24.882 Mean difference, 95% confidence interval 7.922, 4.359-11.485 Adjusted p-value 0.0040

Table A.3 Statistics for Figure 3.3D. NC: number of cells. AUC: area under the curve was used for statistical analysis (n≥3).

GF109203x GF109203x DMSO control control DMSO +LLO +LLO NC AUC NC AUC NC AUC NC AUC 24 13.899 26 14.156 28 13.343 26 14.077 95% confidence interval 13.738-14.060 13.806-14.506 12.986-13.700 13.804-14.349

Unadjusted Adjusted Condition 1 Condition 2 p-value p-value DMSO +LLO DMSO control 0.0004 0.0018 DMSO +LLO GF109203x +LLO <0.0001 0.0003 DMSO +LLO GF109203x control <0.0001 0.0002 DMSO control GF109203x +LLO 0.1052 0.2104 DMSO control GF109203x control 0.0296 0.0888 GF109203x +LLO GF109203x control 0.4383 0.4383 Table A.4. Statistics for Figure 3.8. AUC: area under the FRET efficiency curves, NC : number of cells. The first part of the table presents the AUC values for different experimental conditions. The second part presents the statistics performed on AUC and comparing conditions 1 to conditions 2 (n = 3).

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Condition 1 Condition 2 Time Time % Adjusted Medium LLO %FRET Medium LLO %FRET (min) (min) difference p-value DMSO 20 - 0.87 DMSO 20 + 54.34 -53.47 0.0009 DMSO 20 + 54.34 GF109203x 20 + 19.12 35.22 0.0028 DMSO 20 - 0.87 GF109203x 20 + 19.12 -18.25 0.0185

Table A.5 Statistics for Table 3.2. Statistics compared experimental conditions 1 to experimental conditions 2 and were performed on % FRET (n = 3).

Primer Sequence A 5’ GCAGAGCTGGTTTAGTGAAC 3’ B 5’ CAGTTATCTAGATCCGGTGG 3’ 5’ C CCTGGAGAATATATCCCTACTGTCTTTGACAATTATTCTGCCAATG TTATGG 3’ 5’ D CCATAACATTGGCAGAATAATTGTCAAAGACAGTAGGGATATATT CTCCAGG 3’ E 5’ GCAGAGCTGGTTTAGTGAACC 3’ F 5’ CAAATGTGGTATGGCTGATTATG 3’ G 5’ CGTCCAGCTCGACCAGGATG 3’

Table A.6 Primers used in this study. Sanger sequencing primers were used to confirm the sequences of mCit, mCFP, mCit- Rac1, mCit-Rac1N17, mCit-Rac1L61, and mCFP-PBD (A and B). Site directed mutagenesis primers (C and D) were used to convert mCit-Rac1 H40Y/Q61L to mCit- Rac1 Q61L. Primers used to confirm the sequence of GFP-PKCα and empty vector GFPN1 (E, F and G).

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