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Elucidating the function of the type-III secreted effector proteins EspZ and NleC of the attaching and… Shames, Stephanie Rochelle 2011

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ELUCIDATING THE FUNCTION OF THE TYPE-III SECRETED EFFECTOR PROTEINS EspZ AND NleC OF THE ATTACHING AND EFFACING BACTERIAL PATHOGENS by Stephanie Rochelle Shames  B.Sc., University of Western Ontario, 2006  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in The Faculty of Graduate Studies (Microbiology and Immunology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) December, 2011 © Stephanie Rochelle Shames, 2011  Abstract Enteropathogenic and enterohemorrhagic Escherichia coli (EPEC and EHEC, respectively) are attaching and effacing (A/E) bacterial pathogens that cause diarrheal disease. EPEC causes severe infantile diarrhea in developing countries whereas EHEC infection results in severe bloody diarrhea worldwide. These pathogens employ a typeIII secretion system (T3SS) encoded on the locus of enterocyte effacement (LEE) pathogenicity island (PAI) to inject a panel of effector proteins directly into infected host cells where they subvert host cell functions. The roles of many type-III secreted (T3S) effectors remain to be elucidated.  Here, we have elucidated functions for the T3S  effectors EspZ and NleC. EPEC infection causes host cell cytotoxicity and death. We demonstrated that EspZ enhances host cell survival during EPEC infection. Removal of espZ from the EPEC genome (∆espZ) exacerbated host cell cytotoxicity. We found that EspZ interacts with host CD98 and contributes to protection against EPEC-mediated cytotoxicity by enhancing phosphorylation of focal adhesion kinase (FAK).  Further investigation  revealed that EPEC ∆espZ infection caused a severe decrease in host mitochondrial inner membrane potential (∆ψm) concurrently with host cell lysis. We also found that EspZ localizes to host cell mitochondria and interacts with the translocase of inner mitochondrial membrane (TIM) 17B. These studies are the first to demonstrate EspZ function. Many non-LEE encoded (Nle) effector proteins impact innate immune signaling and we thus examined the contribution of the T3S zinc-metalloprotease NleC to this phenotype. We identified the host acetyltransferase p300 as a target of NleC and show that NleC ii  causes decreased abundance of p300 in cellular nuclei. We further demonstrate that overexpression of p300 antagonizes repression of interleukin (IL)-8 secretion by EPEC and that small interfering ribonucleic acid (siRNA) knock-down of p300 dampens IL-8 secretion by EPEC ∆nleC-infected cells. Thesis work has identified a target of NleC and provided the first example of a bacterial virulence factor targeting the acetyltransferase p300. The work presented in this thesis provides novel insight into the function of two T3S effector proteins, EspZ and NleC. The mechanistic insight gained by these studies has thoroughly enhanced the understanding of these important virulence factors and their contribution to A/E pathogen infection.  iii  Preface The work presented in this dissertation was produced in part through collaborative efforts. Chapter 2 has been published as “The pathogenic E. coli type III effector EspZ interacts with host CD98 and facilitates host cell prosurvival signaling” in the journal Cellular Microbiology. My co-authors for this manuscript contributed as follows: Drs. W. Deng and P.R. Hardwidge performed yeast-two hybrid experiments presented in Table 2.5; Dr. J.A. Guttman made HeLa-EspZ and MDCK-EspZ stable cell lines; Ms. Y. Li, Dr. C.L. de Hoog, and Dr. L.J. Foster performed SILAC experiments and data analyses.; Dr. W. Deng created the EPEC ∆espZ strain.  All of the remaining experiments and data  collection were my own work. The manuscript was written by myself and my supervisor, Dr. B. Finlay. Chapter 3 has published as “The type-III secreted effector EspZ localizes to host mitochondria and interacts with the translocase of inner mitochondrial membrane (TIM) 17B” in the journal Infection and Immunity.  My co-authors on this manuscript  contributed as follows: Dr. W. Deng first cloned TIM17b cDNA from the HeLa cell cDNA library and Dr. M.A. Croxen made the ∆espZ/espZ and ∆espZ/espZHA chromosomally complemented strains used in this study. All of the experiments and data collection presented in this chapter were my own work. The manuscript was written by myself and my supervisor, Dr. B. Finlay. A portion of Chapter 4 has been published as “The attaching/effacing bacterial effector NleC suppresses epithelial inflammatory responses by inhibiting NF-κB and p38-MAP  iv  Kinase activation” in the journal Infection and Immunity in collaboration with Dr. B. Vallance, Mr. H.P. Sham and Dr. M.A. Croxen. Data in Fig. 4.1 was originally acquired by Mr. H.P. Sham and Dr. B.A. Vallance; however, I performed the experiment to generate the figure presented in this thesis. Dr. M.A. Croxen generated the ∆nleC/nleC and ∆nleC/nleCHA chromosomally complemented EPEC strains. A version of Fig 4.3 has been published in the article “The pathogenic Escherichia coli type III secreted protease NleC degrades the host acetyltransferase p300” in the jorunal Cellular Microbiology. I performed all of the experiments presented in this chapter. Chapter 5 has been published as “The pathogenic Escherichia coli type III secreted protease NleC degrades the host acetyltransferase p300” in the journal Cellular Microbiology. My co-authors contributed to this work as follows: Dr. A.P. Bhavsar aided in designing and performing the protease cleavage experiment for Fig 5.11; Dr. M.A. Croxen made the ∆nleC/nleCE184A strain; Dr. W. Deng made the EPEC ∆nleC strain; Dr. C.L. de Hoog, Ms. Y. Li, and Ms. R. Bidshari, who worked under my supervision, helped me with SILAC experiments, and data analysis and LC-MS/MS were preformed by Dr. L.J. Foster; Ms. R.J. Law performed the experiment leading to Fig. 5.10A under my guidance and produced recombinant NleC for the experiment presented in Fig. 5.11; and Ms. S.H.C. Mak worked under my guidance to confirm the NleC-p300 interaction by co-immunoprecipitation presented in Fig 5.2. All remaining experimental design, execution and data collection presented in this chapter were a result of my own work. The manuscript was written by myself and my supervisor, Dr. B. Finlay.  v  Publications arising from graduate work Shames, S.R. and Finlay, B.B. Bacterial effector interplay: a new way to view effector function. Trends in Microbiology. Submitted manuscript # TIMI-D-11-00189 Shames, S.R., Croxen, M.A., Deng, W., and Finlay, B.B. (2011) The type-III system secreted effector EspZ localizes to host mitochondria and interacts with the translocase of inner mitochondrial membrane 17B. Infection and Immunity. 79(12): 4784-4790 Shames, S.R., Bhavsar, A.P. Croxen, M.A., Law, R.J., Mak, S.H.C., Deng, W., Bidshari, R., de Hoog, C.L., Foster, L.J., and Finlay B.B. (2011) The pathogenic Escherichia coli type III secreted protease NleC degrades the host acetyltransferase p300. Cellular Microbiology 13(10): 1542-1557 Sham, H.P.†, Shames, S.R.†, Croxen, M.A., Ma, C., Chan, J., Khan, M.A., Wickham, M.E., Deng, W., Finlay, B.B., and Vallance, B.A. (2011) The attaching/effacing bacterial effector NleC suppresses epithelial inflammatory responses by inhibiting NF-κB and p38MAP Kinase activation. Infection and Immunity. 79(9): 3552-3562 † - These authors contributed equally to this work. Shames, S.R. and Finlay, B.B. (2011) Proteolytic cleavage of NF-κB p65: a novel mechanism for subversion of innate immune signaling by pathogenic E. coli. Frontiers in Microbiology. 2 doi: 10.3389/fmicb.2011.00038 Shames, S.R. and Finlay B.B. (2010) Breaking the stereotype: virulence factor-mediated protection of host cells in bacterial pathogenesis. PLoS Pathogens. 6(9): e1001057 Shames, S.R., Deng, W., Guttman, J.A., de Hoog, C.L., Li, Y., Hardwidge, P.R., Sham, H.P., Vallance, B.A., Foster, L.J., and Finlay, B.B. (2010) The pathogenic E. coli type III effector EspZ interacts with host CD98 and facilitates host cell prosurvival signalling. Cellular Microbiology. 12(9): 1322-1339 Shames, S.R., Auweter, S.D., and Finlay, B.B. (2009) Co-evolution and exploitation of host cell signaling pathways by bacterial pathogens. International Journal of Biochemistry and Molecular Biology 41(2): 380-38  vi  Table of Contents Abstract.............................................................................................................................. ii Preface............................................................................................................................... iv Table of Contents ............................................................................................................ vii List of Tables .................................................................................................................. xiii List of Figures................................................................................................................. xiv List of Symbols and Abbreviations ............................................................................. xvii Acknowledgements ......................................................................................................... xx Dedication ....................................................................................................................... xxi Chapter 1. Introduction: virulence strategies of attaching and effacing pathogens... 1 1.1 Manifestations of A/E pathogen infection ................................................................ 1 1.1.1 EPEC.................................................................................................................. 1 1.1.2 EHEC ................................................................................................................. 3 1.1.3 Citrobacter rodentium ....................................................................................... 5 1.2 Type-III secreted effector proteins............................................................................ 6 1.2.1 LEE-encoded T3S effector proteins................................................................... 6 1.2.2 Non-LEE encoded T3S effectors ..................................................................... 12 1.3 Subversion of host cell death pathways .................................................................. 22 1.3.1 Apoptosis ......................................................................................................... 22 1.3.2 Necrosis............................................................................................................ 24 1.4 Subversion of innate immunity............................................................................... 25 1.4.1 Epithelial cells.................................................................................................. 26  vii  1.4.2 Immune cells.................................................................................................... 29 1.5 Summary of thesis................................................................................................... 31 Chapter 2. The A/E pathogen effector EspZ interacts with host CD98 and facilitates host cell pro-survival signaling. ..................................................................................... 34 2.1 Introduction............................................................................................................. 34 2.2 Experimental procedures ........................................................................................ 36 2.2.1 Tissue culture, bacterial strains, transfection, and infection conditions .......... 36 2.2.2 Generation of CD98-FLAG fusions................................................................. 37 2.2.3 Generation of pcDNA3::HA2 and pcDNA3::HA2espZ constructs .................. 38 2.2.4 EPEC ∆espZ mutant generation....................................................................... 39 2.2.5 Green fluorescent protein (GFP) fusion to EHEC EspZ.................................. 39 2.2.6 Y2H assay ........................................................................................................ 40 2.2.7 SILAC .............................................................................................................. 42 2.2.8 Lactate dehydrogenase (LDH) release assay ................................................... 44 2.2.9 Cell lifting assay .............................................................................................. 44 2.2.10 Caspase-3 activity assay ................................................................................ 45 2.2.11 siRNA ............................................................................................................ 45 2.2.12 Immunoblotting.............................................................................................. 46 2.2.13 Immunoprecipitation...................................................................................... 47 2.2.14 Confocal and wide field microscopy ............................................................. 47 2.2.15 Statistical analyses ......................................................................................... 48 2.3 EPEC ∆espZ causes enhanced detachment and death in HeLa cells and increased cell death in MDCK cells.............................................................................................. 48  viii  2.4 HeLa and MDCK cells expressing EspZ are protected from EPEC-mediated killing ....................................................................................................................................... 51 2.5 EspZ does not antagonize STS-induced apoptotic cell death ................................. 54 2.6 EspZ interacts with host CD98 ............................................................................... 55 2.7 CD98 contributes to EspZ-mediated protection from cytotoxicity......................... 64 2.8 EspZ influences AKT and FAK phosphorylation during EPEC infection ............. 66 2.9 Knock-down of CD98 decreases EspZ-mediated phosphorylation of FAK.......... 68 2.10 Discussion ............................................................................................................ 70 Chapter 3: EspZ localizes to host mitochondria and interacts with host translocase of inner mitochondrial membrane (TIM) 17B ............................................................. 76 3.1 Introduction............................................................................................................. 76 3.2 Experimental procedures ........................................................................................ 77 3.2.1 Tissue culture, bacterial strains, primers, transfection and infection conditions ................................................................................................................................... 77 3.2.2 Generation of espZ and espZHA complemented strains .................................. 78 3.2.3 Generation of TIM17B-FLAG fusion.............................................................. 78 3.2.4 JC-1 ∆ψm assay ................................................................................................ 80 3.2.5 LDH release assay............................................................................................ 80 3.2.6 Confocal microscopy ....................................................................................... 81 3.2.7 Immunoprecipitation........................................................................................ 82 3.2.8 Immunoblotting................................................................................................ 82 3.2.9 Isolation of cellular mitochondria.................................................................... 83 3.2.10 siRNA ............................................................................................................ 84  ix  3.2.11 T3S assay ....................................................................................................... 84 3.2.12 Statistical analysis.......................................................................................... 85 3.3 Chromosomal insertion of espZ can complement the ∆espZ knock-out and preserve type-III secretion of other effector proteins .................................................................. 85 3.4 EspZ protects EPEC-infected cells from severe loss of inner mitochondrial membrane potential (Δψm)............................................................................................ 87 3.5 EspZ interacts with TIM17B .................................................................................. 89 3.6 EspZ localizes to mitochondria............................................................................... 90 3.7 EspZ mitochondrial localization is independent of TIM17B ................................. 93 3.8 siRNA knock-down of TIM17B dampens EspZ-mediated survival during EPEC infection ........................................................................................................................ 94 3.9 Discussion .............................................................................................................. 97 Chapter 4. The non-LEE encoded effector NleC dampens host NF-κB signaling and IL-8 secretion during EPEC infection......................................................................... 100 4.1 Introduction........................................................................................................... 100 4.2 Experimental procedures ...................................................................................... 101 4.2.1 Tissue culture, bacterial strains and infection conditions .............................. 101 4.2.2 Generation of EPEC ∆nleC............................................................................ 101 4.2.3 Generation of ∆nleC/nleC and ∆nleC/nleCHA complemented strains .......... 102 4.2.4 NF-κB activity assay...................................................................................... 104 4.2.5 IL-8 enzyme-linked immunosorbent assay (ELISA)..................................... 105 4.2.6 Nuclear isolation and western blot analysis................................................... 105 4.2.7 Statistical analyses ......................................................................................... 106  x  4.3 NleC reduces IL-8 secretion during EPEC infection............................................ 106 4.4 EPEC ∆nleC infection results in nuclear p65 accumulation................................. 107 4.5 NleC contributes to repression of NF-κB signaling ............................................. 109 4.6 Discussion ............................................................................................................. 110 Chapter 5. NleC interacts with and degrades the host acetyltransferase p300....... 113 5.1 Introduction........................................................................................................... 113 5.2 Experimental procedures ...................................................................................... 114 5.2.1 Tissue culture, bacterial strains, transfection and infection condition........... 114 5.2.2 p300 truncations............................................................................................. 115 5.2.3 GFP and HA2 epitope tag fusions to EHEC nleC .......................................... 115 5.2.4 Generation of NleCE184A-GFPN and NleCE184A-FLAG ................................. 116 5.2.5 Recombinant protein purification .................................................................. 117 5.2.6 SILAC ............................................................................................................ 119 5.2.7 Immunoprecipitation...................................................................................... 120 5.2.8 Confocal microscopy ..................................................................................... 120 5.2.9 Isolation of cellular nuclei ............................................................................. 121 5.2.10 In vitro cleavage assay ................................................................................. 122 5.2.11 Immunoblotting............................................................................................ 122 5.2.12 siRNA .......................................................................................................... 123 5.2.13 IL-8 ELISA .................................................................................................. 123 5.2.14 Statistical analysis........................................................................................ 124 5.3 NleC interacts with host p300 by SILAC ............................................................. 124 5.4 NleC co-immunoprecipitates with endogenous p300 ........................................... 126  xi  5.5 The NleC-p300 interaction is dependent on the TAZ1 domain of p300 .............. 127 5.6 Global acetylation levels in Caco-2 cells are increased during infection with EPEC ∆nleC .......................................................................................................................... 130 5.7 NleC facilitates decreased nuclear p300 levels in HeLa cells .............................. 131 5.8 The metalloprotease domain of NleC contributes to decreased nuclear p300...... 136 5.9 NleC facilitates p300 degradation in vitro ............................................................ 139 5.10 Overexpression of p300 enhances IL-8 secretion during WT EPEC infection .. 140 5.11 Knock-down of p300 dampens IL-8 secretion from cells infected with EPEC ∆nleC .......................................................................................................................... 142 5.12 Discussion ........................................................................................................... 143 Chapter 6. Discussion ................................................................................................... 149 6.1 Protection of host cells from cytotoxicity in bacterial pathogenesis .................... 149 6.2 Interplay between A/E T3S effectors in host cells as a mechanism for highly regulated cytotoxicity?................................................................................................ 154 6.3 Subversion of innate immune signaling by bacterial pathogens: a recurring theme ..................................................................................................................................... 155 6.4 Uncovering roles for bacterial effector proteins ................................................... 159 References...................................................................................................................... 163 Appendices..................................................................................................................... 184 Appendix  A.  Summary  of  proteins  identified  by  MS  as  specifically  immunoprecipitated with HA2EspZ in SILAC experiments ...................................... 184 Appendix B. Peptides of E1A-associated protein p300 identified in NleC SILAC experiment................................................................................................................... 187  xii  List of Tables Table 2.1 Bacterial strains used in this study.................................................................... 37 Table 2.2 Oligonucleotide primers used in this study....................................................... 41 Table 2.3 Plasmids used in this study ............................................................................... 43 Table 2.4 Summary of proteins identified by MS as specifically immunoprecipitated with HA2EspZ in SILAC experiments...................................................................................... 58 Table 2.5 Proteins identified in Y2H screen for interaction with EspZ............................ 58 Table 3.1 Bacterial strains used in this study.................................................................... 79 Table 3.2 Oligonucleotide primers used in this study....................................................... 79 Table 3.3 Plasmids used in this study ............................................................................... 79 Table 4.1 Bacterial strains used in this study.................................................................. 103 Table 4.2 Oligonucleotides used in this study ................................................................ 103 Table 4.3 Plasmids used in this study ............................................................................ 105 Table 5.1 Bacterial strains used in this study.................................................................. 116 Table 5.2 Oligonucleotides used in this study ................................................................ 116 Table 5.3 Plasmids used in this study ............................................................................. 117 Table 5.4 Summary of proteins identified by MS as specifically interacting with GSTHA3-NleC in SILAC experiments .................................................................................. 126  xiii  List of Figures Fig 2.1 EPEC ∆espZ infection influences host cell lifting and death ............................... 50 Fig 2.2 MDCK cell adherence following 8 h infection .................................................... 51 Fig 2.3 Coomassie Brilliant Blue staining of secreted proteins from various EPEC strains ........................................................................................................................................... 52 Fig 2.4 Ectopic expression of EspZ protects against cytotoxicity .................................... 53 Fig 2.5 EspZ does not protect HeLa cells from Caspase-3 dependent apoptosis ............. 55 Fig 2.6 SILAC screen for EspZ interacting proteins ........................................................ 57 Fig 2.7 EspZ and CD98 both localize at the host cell plasma membrane ........................ 60 Fig 2.8. CD98 localizes at sites of bacterial attachment. .................................................. 61 Fig 2.9. HA2EspZ co-immunoprecipitated with CD98-FLAG........................................ 62 Fig 2.10 The first 42 amino acid residues of EspZ are not required for interaction with CD98 ................................................................................................................................. 63 Fig 2.11 LDH release of mock infected HeLa and HeLa-EspZ cells treated with nt siRNA or CD98 siRNA ................................................................................................................ 65 Fig 2.12 CD98 contributes to EspZ-mediated protection from EPEC-induced cytotoxicity ........................................................................................................................................... 65 Fig. 2.13 EspZ enhances phosphorylation of AKT and FAK during EPEC infection ..... 67 Fig 2.14 Influence of CD98 knock-down on EspZ-mediated signaling ........................... 69 Fig 2.15 Schematic model for EspZ function during EPEC infection.............................. 75 Fig 3.1 Chromosomal insertion of espZ preserves type-III secretion of effectors and translocators from EPEC................................................................................................... 86 Fig 3.2 Infection with EPEC ∆espZ causes severe loss of ∆ψm........................................ 88  xiv  Fig 3.3 TIM17B interacts with EspZ ................................................................................ 90 Fig 3.4 EspZ localizes to host cell mitochondria.............................................................. 92 Fig 3.5 The EspZ-TIM17B interaction is not required for mitochondrial localization of EspZ .................................................................................................................................. 94 Fig 3.6 siRNA knock-down of TIM17B dampens the ability of EspZ to protect HeLa cells from EPEC-mediated cell lysis................................................................................. 96 Fig 4.1 NleC contributes to repression of IL-8 secretion during EPEC infection .......... 107 Fig 4.2 p65 translocates to nuclear fractions in EPEC ∆nleC-infected Caco2 cells....... 108 Fig 4.3 Infection of Caco-2 cells with EPEC ∆nleC/nleC results in similar NF-κB activity levels to WT EPEC ......................................................................................................... 110 Fig 5.1 Representative mass spectra for peptides recovered from SILAC experiments 125 Fig 5.2 NleC co-immunoprecipitates with endogenous p300......................................... 127 Fig 5.3 The TAZ1 domain of p300 is required for interaction with NleC...................... 129 Fig 5.4 NleC does not co-immunoprecipitate with the C-terminus of p300................... 130 Fig 5.5 Global acetylation levels in Caco-2 cell cytosol is increased during infection of Caco-2 cells with EPEC ∆nleC....................................................................................... 131 Fig 5.6 NleC facilitates decreased nuclear p300 levels .................................................. 133 Fig 5.7 Ectopic expression of NleC decreases nuclear p300 protein levels in HeLa cells ......................................................................................................................................... 135 Fig 5.8 Increasing concentration of HA2NleC causes decreased p300 protein levels in Caco-2 cells..................................................................................................................... 135 Fig 5.9 NleCE184A interacts with endogenous p300 ........................................................ 137 Fig 5.10 The metalloprotease domain of NleC contributes to decreased nuclear p300 . 139  xv  Fig 5.11 NleC cleaves p300 in vitro ............................................................................... 140 Fig 5.12 Over-expression of p300 causes enhanced IL-8 secretion during EPEC infection ......................................................................................................................................... 141 Fig 5.13 siRNA knock-down of p300 dampens IL-8 secretion during EPEC ∆nleC infection .......................................................................................................................... 143 Fig 6.1 Strategies evolved by bacterial pathogens to restrain virulence......................... 153  xvi  List of Symbols and Abbreviations Δψm A/E Ac-K APC ARF Arp2/3 ATCC ATP BFP BI-1 BSA CBP CHAPS Cif COPII COXIV CPAF CPM CR3 DAPI DC DMEM DMSO DNA EAF ECL EPEC EHEC ELISA ER Erk1/2 FAK FBS FcγR Gb3 GFP GM6001 h HAT HEK HRP Hsp HUS  mitochondrial membrane potential attaching and effacing acetyl-lysine antigen presenting cell ADP-ribosylation factor actin-related protein 2/3 american type culture collection adenosine trisphosphate bundle forming pilus bax inhibitor-1 bovine serum albumin cAMP response element-binding protein 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate cycle inhibitory factor coat protein-2 cytochorome c oxidase IV chlamydial protease-like activity factor counts per minute complement receptor 3 4',6-diamidino-2-phenylindole dendritic cell dulbecco’s modified eagle medium dimethyl sulfoxide deoxyribonucleic acids enteropathogenic Escherichia coli adherence factor enhanced chemiluminescence enteropathogenic Escherichia coli enterohaemorrhagic Escherichia coli enzyme-linked immunosorbent assay endoplasmic reticulum extracellular signal-regulated kinase 1/2 focal adhesion kinase fetal bovine serum fc gamma receptor globotriaosylceramide green fluorescent protein galardin, Ilomastat, N-[(2R)-2-(Hydroxamidocarbonylmethyl)-4methylpentanoyl]-L-tryptophan methylamide hours histone acetyltransferase human embryonic kindney horseradish peroxidase heat shock protein hemolytic uremic syndrome xvii  IκBα IC IEC IKKβ IL-1β IL-8 IMM IPTG JC-1 JNK kb kDa LC-MS/MS LDH LEE m/z Map MCS MDa MDCK min MIR m.o.i. MT mtHsp MTS N-WASP NAPS NEAA NF-κB NGS NHERF2 Nle NMR NOD1 NP40 nt NuR OMM PAGE PAI PAK PAM PAMPs PBST/BSA  inhibitor of kappaB apha isotype control intestinal epithelial cell inhibitor of kappaB apha kinase beta interleukin-1 beta interleukin-8 inner mitochondrial membrane isopropyl β-D-1-thiogalactopyranoside j-aggregate-forming lipophilic cation 5,5',6,6'-tetrachloro1,1',3,3'-tetraethylbenzimidazolcarbocyanine iodide c-Jun N-terminal kinase kilobase kilodalton liquid chromatography-tandem mass spectrometry lactate dehydrogenase locus of enterocyte effacement mass to charge ratio mitochondria associated protein multiple cloning site megadalton madin-darby canine kidney minute mitochondria isolation reagent multiplicity of infection microtubule mitochondrial heat shock protein mitochondrial targeting signal neuronal Wiskott-Aldrich Syndrome protein nucleic acid protein service non-essential amino acids nuclear factor-kappa B normal goat serum sodium hydrogen exchanger regulatory factor 2 non-LEE encoded nuclear magentic resonance nucleotide-binding oligomerization domain-containing protein 1 nonidet P40 non-targeting nuclear hormone receptor outer mitochondrial membrane polyacrylamide gel eletrophoresis pathogenicity island p21-activated kinase presequence translocase-associated motor pathogen associated molecular patterns phosphate-buffered saline/0.1% BSA/0.5% Tween 20  xviii  p.i. PI3K PRR PVDF r.c.f. RDEC RFU Rip2 RNA ROS RPS3 S. Typhimurium S.D. SDS siRNA SILAC STS Stx T3S T3SS TBS TBST TCA TER TIM Tir TLR TOM TNF UBC UI WT  post-infection phosphoinositide-3 kinase proline-rich repeat polyvinylidene fluoride relative centrifugal force rabbit diarrheagenic E. coli relative fluorescence units receptor interacting protein 2 ribonucleic acid reactive oxygen species ribosomal protein-s3 Salmonella enterica serovar Typhimurium standard deviation sodium dodecyl-sulfate small interfering ribonucleic acids stable isotope labeling of amino acids in cell culture staurosporine shiga toxin type-III secreted type-III secretion system tris-buffered saline tris-buffered saline/0.1% Tween 20 trichloroacetic acid transepithelial resistance translocase of the inner mitochondrial membrane translocated intimin receptor toll-like receptor translocase of the outer mitochondrial membrane tumor necrosis factor University of British Columbia uninfected wild-type  xix  Acknowledgements I would like to extend my utmost appreciation to the many people who made this work possible. The members of the Finlay Lab, past and present, have been an unrelenting source of insight and camaraderie.  I am honored to have spent time working and  socializing with you all and I feel I have made excellent friends in addition to colleagues. I would especially like to thank the EPEC group for our monthly meetings and all of your advice. I will dearly miss you all. Deng, thanks so much for all of your help (and it’s been a LOT) over my five-and-a-bit years in the lab and for proofreading this thesis in less than two days! Carmen, thank you so much for your help with MS Word. You’ve saved me countless headaches. I am indebted to my supervisor, Dr. Brett Finlay, for providing me with freedom to explore research projects that interested me, but ensuring that I remain focused. Your continuous support and advice has made me into the scientist that I am today. I would also like to thank my supervisory committee, Drs. Leonard Foster, Bruce Vallance, and Wayne Vogl for always challenging me and providing me with invaluable guidance throughout my graduate career. Thanks to my family, especially my parents, for always offering moral support and understanding that my pursuit of knowledge will take me far away from home. Finally, I would like to thank my husband, Tom, who was present for my journey through graduate school. Thank you for your love and support and for your unrelenting patience. Your continuous support encourages me to thrive as a scientist and accomplish all I can.  xx  Dedication This dissertation is dedicated to the memory of my dear friend and colleague Aaron Wyatt. You provided invaluable friendship and advice, always set an excellent example to live by, and were the most altruistic and self-less person I’ve ever known. I cherish our memories and think of you always. You didn’t get a chance to finish your own Ph.D., so this one is for you. Rest in peace, homeslice.  Aaron William James Wyatt (1983-2008)  xxi  Chapter 1. Introduction: virulence strategies of attaching and effacing pathogens 1.1 Manifestations of A/E pathogen infection A/E pathogens are a distinct class of diarrheagenic E. coli that are characterized by the formation of A/E lesions on the surface of intestinal epithelial cells (IECs). These pathogens bind to the apical surface of the intestinal epithelium, locally efface microvilli and form actin-rich protrusions on the surface of infected cells. Generation of A/E lesions is dependent on a 35- kilobase (kb) PAI called the locus of enterocyte effacement (LEE), which encodes a type-III secretion system (T3SS). The T3SS functions to inject bacterial ‘effector’ proteins directly into eukaryotic host cells where they subvert host cell signaling cascades.  The A/E pathogen family includes EPEC, EHEC, the murine  pathogen Citrobacter rodentium, rabbit diarrheagenic E. coli (RDEC) and EPEC strains that infect a wide range of warm-blooded animals. In this chapter, the discussion of A/E pathogens will focus on EPEC, EHEC, and C. rodentium, as they are the most studied. 1.1.1 EPEC EPEC is a human-restricted pathogen that causes significant infant mortality in developing countries (Croxen et al., 2010). Infection results in watery diarrhea and occasionally vomiting that are often self-limiting in adults. For EPEC to successfully infect healthy adults, a very high initial inoculum is required (Schroeder et al., 1968). EPEC is transmitted by the fecal-oral route followed by localization to the small bowel where it attaches to IECs and forms A/E lesions on their surface [reviewed in (Croxen et al., 2010)]. Diarrhea is thought to occur partially due to aberrant nutrient absorption 1  resulting from bacterial attachment to the epithelium (Rothbaum et al., 1982), which may result in osmotic loss of water in the large bowel. Attachment to the small bowel epithelium is essential for disease and it is therefore important to examine the details of EPEC infection. Bundle-forming pili (BFP) genes encoded on the EPEC adherence factor (EAF) plasmid and the E. coli common pilus are thought to mediate initial EPEC attachment to epithelial cells (Saldana et al., 2009, Croxen et al., 2010). The EAF plasmid is able to confer an attaching phenotype to the normally non-adherent E. coli HB101 (Baldini et al., 1983). BFP also facilitates bacteria-bacteria attachment and formation of EPEC microcolonies (Kaper et al., 2004, Knutton et al., 1999). The role of microcolony formation is poorly understood; however, it may mediate increased initial EPEC colonization leading to greater levels of bacterial colonization and shedding. Following initial pilus-mediated adhesion, EPEC uses its LEE-encoded T3SS to rapidly inject virulence proteins – termed effectors – into infected host IECs. The LEE is essential for EPEC-mediated disease and contains 41 open reading frames (orfs) (Deng et al., 2004).  The LEE encodes seven effector proteins, T3SS structural components,  chaperones, and translocator proteins (Schmidt, 2010). To date, EPEC has been shown to encode at least 21 effector proteins genome-wide, and more are still being identified (Iguchi et al., 2009). The initial event that leads to intimate attachment of EPEC to host IECs is the injection of the LEE-encoded translocated Intimin receptor (Tir) into the host cell. Tir integrates into the host apical plasma membrane where it then interacts with the bacterial outer membrane adhesin Intimin and this interaction leads to A/E lesion, or pedestal, formation under adherent bacteria (Kenny et al., 1997). 2  Unlike EHEC and C. rodentium, EPEC quickly and efficiently infects cultured cells, which enables identification of molecular disease mechanisms and facilitates investigation into the role of T3S effector proteins. 1.1.2 EHEC Although both EPEC and EHEC are human-restricted diarrheagenic E. coli, there are several major distinctions between these two pathogens.  EHEC infection occurs  primarily in the large bowel and worldwide and disease is not influenced by socioeconomic factors. EHEC was first distinguished from EPEC in 1983 based on the presence of bloody stools and cases of severe renal failure termed HUS (Riley et al., 1983, Karmali et al., 1983).  EHEC likely evolved from EPEC strains following  acquisition of lysogenic bacteriophages encoding Shiga toxin genes (stx) (O'Brien et al., 1984). No animal reservoir for EPEC has been identified; however, EHEC colonizes ruminants, which are a major source of human infection. Similarly to EPEC, EHEC is transmitted via the fecal-oral route and the majority of infections result from consumption of undercooked hamburger and water contaminated with cattle feces (Ferens et al., 2011). In addition to the Stx genes, EHEC also contains a LEE PAI and 60-MDa plasmid, which encodes enterohemolysin genes (Spears et al., 2006, Ferens et al., 2011). EHEC expresses different adhesion proteins from EPEC.  The bfp genes and EAF  plasmid are absent in EHEC; however, EHEC appear to express two operons encoding lpf and lpfA genes that are similar to the Salmonella long polar fimbriae (Spears et al., 2006). In addition, the EHEC eae gene, encoding Intimin, is thought to be responsible for persistence of the pathogen in the large bowel (Spears et al., 2006).  3  Since EPEC  colonizes the small bowel and EHEC colonizes the large bowel, it is not surprising that they express different adhesion proteins. The stx genes differentiate EHEC from other A/E pathogens.  There are two  immunologically distinct Stx proteins - Stx1 and Stx2 – with different EHEC strains encoding Stx1, Stx2, or both toxins. Stx1 and Stx2 are both AB toxins and the B subunit facilitates entry into host cells whereas the A subunit enzymatically cleaves 28S ribosomal RNA, thus halting cellular protein synthesis (Endo et al., 1988). Entrance of Stx into the bloodstream can result in HUS.  The Stx B subunit binds to  globotriaosylceramide (Gb3) to facilitate delivery of the A subunit and Gb3 is present in abundance on human renal cells (Obrig et al., 1993). Ruminants lack Gb3 and are therefore resistant to the effect of Stx (Mainil, 1999). HUS is the leading cause of renal failure in children in the United States (Fiorino et al., 2006). EHEC are A/E pathogens and therefore also require the LEE PAI to cause disease. The LEE PAI from EPEC can confer the A/E phenotype on E. coli K-12 (McDaniel et al., 1997); however, the EHEC LEE is not sufficient to allow E. coli K-12 to form pedestals (Elliott et al., 1999). The LEE is ~ 93% similar between EPEC and EHEC; however, the effector repertoire is very different. For pedestal formation, EHEC requires an Nle protein called EspFu/TccP to function with Tir (Campellone et al., 2004). In addition, EHEC is predicted to encode >40 effector proteins, with 14 of these being part of the NleG family (Tobe et al., 2006). Despite the difference in number of effectors, a core set of effectors are well conserved between EPEC and EHEC [reviewed in (Spears et al., 2006)].  4  1.1.3 Citrobacter rodentium C. rodentium is a natural murine pathogen that is often used to model EPEC and EHEC infection in vivo and causes transmissible colonic hyperplasia (Luperchio et al., 2001, Eckmann, 2006). C. rodentium encodes a LEE PAI, which enables formation of A/E lesions that are indistinguishable from those caused by EPEC and EHEC, and has been shown to secrete >30 effector proteins (Luperchio et al., 2001, Deng et al., 2010). Susceptibility to C. rodentium infection in laboratory mice is strain-specific. Vallance and colleagues used a panel of murine strains and demonstrated that C3H/HeJ, C3H/HeOuJ, and C3H/HeN mouse strains are the most susceptible to C. rodentium infection with 100% mortality observed after 10-14 days post-infection (p.i.) (Vallance et al., 2003).  Resistant murine strains tested, including C57BL/6, BALB/c and  129SI/SvImJ, did not demonstrate severe mortality (Vallance et al., 2003). No mortality was observed following infection of 129SI/SvImJ, whereas BALB/c, C57BL/6, and NIH Swiss mice experienced only 20% mortality; however, high bacterial burden is observed in the colon at 6 days p.i. in both resistant and susceptible murine hosts (Vallance et al., 2003). Despite the formation of A/E lesions, C. rodentium infection does not perfectly mirror A/E pathogen disease in humans. There is a milder diarrheal phenotype associated and the initial site of colonization is the caecum, an organ that less prominent in humans (Luperchio et al., 2001, Mundy et al., 2005). Although C. rodentium infection does not share all characteristics with either EPEC or EHEC, it is a valuable model organism to study A/E-mediated disease in vivo.  5  1.2 Type-III secreted effector proteins All A/E pathogens are dependent on T3S effector proteins for their virulence. The effector repertoire of EPEC, EHEC, and C. rodentium differs; however, there are similarities that provide insight into the function of many effector proteins. Most T3S effector protein functions have been investigated in EPEC, since this pathogen infects cultured cells more efficiently than EHEC or C. rodentium. Despite the plethora of studies focusing on the function of effector proteins, the function of many remains to be elucidated and it is likely that effectors of known function also have other roles within host cells during infection. 1.2.1 LEE-encoded T3S effector proteins The A/E pathogen LEE encodes seven T3S effector proteins including EspF, EspH, Map, Tir, EspB, EspG, and EspZ (Deng et al., 2004). Although EspB is a translocator that contributes to pore formation at the host plasma membrane, there is a host cell cytosolic pool of EspB that influences adherens junctions and the actin cytoskeleton (Iizumi et al., 2007). EspB interacts with α-catenin, which then inhibits interaction of α-catenin with actin and the E-cadherin/β-catenin protein complex. Inhibition of the E-cadherin/βcatenin/α-catenin protein complex at adherens junctions likely decreases cell-cell junction stability, which would enhance paracellular permeability between cells. EspB also interacts with several cellular myosins and inhibits their interaction with actin, which antagonizes phagocytosis and facilitates microvilli effacement (Iizumi et al., 2007). Based on its role as a translocator protein, EspB is secreted at early time points.  6  Recently, the secretion hierarchy of other LEE-encoded effector proteins has begun to be unveiled. Translocation of EspF, EspG, EspH, Map, Tir and EspZ by the T3SS is dependent on several factors including level of bacterial attachment, chaperone binding, and effector concentration (Mills et al., 2008). Mills and colleagues also suggested that Tir and EspZ are the first and most efficiently secreted effectors followed by EspF, EspH, EspG, and Map (Mills et al., 2008). Although this study has revealed a hierarchy for six effectors, the secretion levels, timing, and efficiency of the remaining effectors have yet to be elucidated.  Consequently, many effectors have been studied in isolation by use of  effector knockout mutants and expression of effector genes in eukaryotic cells. These techniques have provided much insight into the function of many effector proteins and the theme of multiple functions per effector is becoming more apparent. Tir is one of the best-studied effectors of the entire A/E pathogen effector repertoire. Tir is required to facilitate intimate attachment and pedestal formation under adherent bacteria and is essential for bacterial colonization in vivo (Deng et al., 2003). Intimin present on the surface of the bacteria is thought to cluster translocated Tir proteins present in the apical plasma membrane of IECs. Interestingly, Tir differs between EPEC and EHEC. Following clustering, TirEPEC is phosphorylated at Tyr474 and Tyr454 by eukaryotic tyrosine kinases c-Fyn, Abl-family kinases Abl and Arg, and the Tec-family kinase Etk (Campellone, 2010). Conversely, tyrosine phosphorylation of TirEHEC is not required for pedestal formation (DeVinney et al., 2001). Phosphorylated Tyr residues on TirEPEC subsequently recruit the adaptor protein Nck that then facilitates binding of neuronal Wiskott-Aldrich syndrome protein (N-WASP). The actin-related protein 2/3 7  (Arp2/3) then interacts with N-WASP and facilitates nucleation of actin and formation of the actin-rich pedestal (Gruenheid et al., 2001). Interestingly, the A/E pedestal shows similarity to focal adhesions in cultured cells and many eukaryotic proteins that are normally found in focal adhesions have been shown to be present in A/E pathogeninduced pedestals (Goosney et al., 2001). Recently, TirEPEC was shown to impact the function of EspG and the Nle effector EspG2, which will be discussed here, during infection in culture. Dean and colleagues observed that infection with a tir-deficient strain of EPEC (∆tir) results in loss of epithelial monolayer stability and cell death (Dean et al., 2010a). They further demonstrated that this phenotype could be antagonized by deletion of espG and espG2 or treatment with a calpain inhibitor, and that EspG and EspG2 cause over-activation of calpain, but only in the absence of Tir (Dean et al., 2010a). Prior to this work, EspG was shown to facilitate destruction of microtubules (MTs) and increase paracellular permeability in polarized cell monolayers (Matsuzawa et al., 2005, Shaw et al., 2005, Smollett et al., 2006). Very recent studies also demonstrate that EHEC EspG interacts with ADP-ribosylation factor (ARF) 6 GTPases and p21-activated kinase (PAK) (Selyunin et al., 2011, Germane et al., 2011).  Interaction of EspG with ARF6 functions to inhibit protein-mediated GTP  hydrolysis, which the authors suggest impairs Golgi-mediated trafficking (Selyunin et al., 2011). EspG also activates PAK, but the functional consequences of this activation during EPEC infection have yet to be explored (Clements et al., 2011, Germane et al., 2011). Moreover, a separate study demonstrated an interaction between EspG and the Golgi protein GM130, which resulted in dysregulation of Golgi structure and function (Clements et al., 2011). Thus, it appears that EspG functions to impair Golgi function  8  early in A/E pathogen infection, which may prevent secretion of immune mediators from host cells. Another very well studied LEE-encoded effector is EspF, which performs several functions during A/E pathogen infection.  EspF from all A/E pathogens examined  contains an N-terminal secretion signal and mitochondrial targeting sequence (MTS) in addition to several proline rich repeats. EspF contributes to apoptosis of EPEC-infected or espF-expressing host cells, which is dependent on its localization to mitochondria (Nougayrede et al., 2004, Nagai et al., 2005, Crane et al., 2001). The MTS of EspF is located in the first 101 amino acids of the protein and the translocase of outer mitochondrial membrane (TOM) 20 machinery is predicted to facilitate EspF import into the mitochondria (Nagai et al., 2005, Nougayrede et al., 2004). EspF-induced apoptotic cell death was shown to be influenced by the host protein Abcf2, which interacts with EspF in host cells (Nougayrede et al., 2007).  A C. rodentium ∆espF strain was  attenuated for virulence during infection of C3H/HeJ mice and complementation of this mutation with a WT copy of espF, but not a mitochondrial targeting mutant, espFL16E, could restore virulence (Nagai et al., 2005). EspF has also been shown to interact with cytokeratin 18 and sorting nexin 9 (Marches et al., 2006, Viswanathan et al., 2004). The influence of these interactions on A/E pathogen virulence has not been uncovered; however, regulation of cytoskeletal signaling and vesicle transport during infection has been implied (Alto et al., 2007).  Hodges and colleagues demonstrated that EspF  decreases sodium hydrogen exchanger 3 activity in host epithelial cells independent of other known phenotypes, which may contribute to diarrhea caused by A/E pathogens (Hodges et al., 2008). EspF also contributes to inhibition of macrophage phagocytosis  9  independently of mitochondrial targeting (Quitard et al., 2006) and recently, EspF was also shown to localize to the host nucleolus and impair protein production (Dean et al., 2010b).  EspF is an excellent example of a multi-functional effector protein that can  perform a plethora of roles during infection, although the spatiotemporal regulation of these functions during infection has not been explored in detail. The mitochondria associated protein (Map) from A/E pathogens was identified by interaction with host mitochondria. Map is also imported into the host mitochondria via an N-terminal MTS and utilizes TOM22 and TOM40 in conjunction with mitochondrial heat shock protein 70 (mtHsp70) to enter the mitochondrial matrix in yeast cells (Papatheodorou et al., 2006).  Although Map has not been directly associated with  increased apoptosis during infection, it does alter the structure of host mitochondria, impair Δψm, and facilitate increased perinuclear clustering of mitochondria during A/E pathogen infection (Ma et al., 2006, Papatheodorou et al., 2006). Map contains a WxxxE motif and activates the Rho GTPase Cdc42 to facilitate formation of filopodia on the surface of infected cultured host cells (Huang et al., 2009, Berger et al., 2009). Map has also been shown to work cooperatively with other effector proteins. Map and EspF act together to cause maximal loss of TER in polarized epithelial cells and EspF, Map, Tir, and Intimin function together to inhibit the sodium-D-glucose co-transporter, suggesting that Map promotes diarrhea in host organisms by acting together with other LEE-encoded effector proteins (Dean et al., 2004, Dean et al., 2006). Map also interacts with Na+/H+ exchanger regulatory factor 1 and induces its degradation, which may also contribute to A/E pathogen-induced diarrhea (Simpson et al., 2006). A C. rodentium map mutant (∆map) was able to colonize to the same extent as WT; however, the stools of C3H/HeJ  10  mice contained less water following infection with C. rodentium ∆map compared to WT infected mice (Simpson et al., 2006, Guttman et al., 2006). Of the seven LEE-encoded effectors, roles for EspH and EspZ have been the most recently unveiled. In 2005, EspZ was demonstrated to be a T3S effector protein by Kanack and colleagues and had an unknown function prior to the studies presented in this thesis (see Chapter 2 and Chapter 3). EspH was initially shown to influence filopodia and pedestal formation on the surface of EHEC-infected host cells (Tu et al., 2003). Mutation of espH influenced filopodia and pedestal formation while over-expression of espH caused an elongated pedestal phenotype (Tu et al., 2003). In yeast, EspH was also shown to disrupt MAP kinase signaling (Rodriguez-Escudero et al., 2005). However, a recent study by Dong and colleagues demonstrated that EspH targets and disrupts the host actin cytoskeleton, which likely occurs by EspH-mediated inhibition of Rho GTPase activity (Dong et al., 2010). This study also demonstrated that EspH contributes to inhibition of macrophage phagocytosis (Dong et al., 2010). An EHEC espH mutant could not colonize to WT levels in a rabbit in vivo infection model (Ritchie et al., 2005) and a C. rodentium ∆espH mutant was slightly attenuated for virulence following C3H/HeJ murine infection (Deng et al., 2004). LEE-encoded effector proteins appear to play important roles in modulation of the host cell cytoskeleton, cell death, and solute movement through the intestinal epithelium. A majority of the effectors function to subvert signaling by host GTPases, many of which regulate cytoskeletal dynamics. In addition, induction of cell death by effectors at late time points during infection may permit sloughing of infected epithelial cells and subsequent dissemination of the pathogen to a naïve host. 11  1.2.2 Non-LEE encoded T3S effectors Nle effector proteins are located on genomic islands throughout A/E pathogen genomes. Despite the high homology between the LEE PAIs of different A/E pathogens, the Nle effectors exhibit increased diversity. Here, the literature regarding T3S Nle effectors will be discussed. Nle proteins discussed below include NleA, NleB, NleC, NleD, NleE, NleF, NleG, NleH, NleL, EspJ, EspM, EspT, EspV, and cycle inhibitory factor (Cif). One of the most extensively studied effectors encoded outside the LEE is NleA (also called EspI), which is secreted at higher levels than many other Nle effectors and has the greatest influence on virulence in vivo of the Nle effectors examined to date (Gruenheid et al., 2004). NleA is highly conserved between EPEC, EHEC and C. rodentium and likely performs similar functions during infection (Gruenheid et al., 2004). Gruenheid and colleagues also found that EHEC infected cells had clustering of NleA under their nuclei that co-localized with the Golgi-specific mannosidase II (Gruenheid et al., 2004). NleA contains a PDZ-binding domain, which contributes to interactions with host proteins involved in vesicle transport, and is required for NleA host Golgi localization (Lee et al., 2008). An elegant study performed by Kim and colleagues subsequently revealed that NleA inhibits coat protein complex II (COPII) function to impair vesicle transport and protein secretion during A/E pathogen infection (Kim et al., 2007). These authors demonstrated that NleA interacts with COPII components and inhibits vesicle trafficking between the endoplasmic reticulum (ER) and the Golgi in addition to inhibition of protein secretion (Kim et al., 2007). NleA was also demonstrated to disrupt tight junctions during EPEC infection and this is related to disruption of COPII function (Thanabalasuriar et al., 2010). Identification of NleA demonstrated the presence of a 12  crucial A/E pathogen virulence factor encoded outside the LEE PAI and further investigation should focus on the contribution of NleA-mediated tight junction disruption and COPII subversion to virulence in vivo, which may contribute to decreased cytokine secretion and diarrhea. NleB was first identified as a T3S effector protein in a signature tagged mutagenesis screen for C. rodentium mutants attenuated for virulence in the C3H/HeJ murine infection model (Kelly et al., 2006). The authors found that C. rodentium ∆nleB could not colonize to the same extent as C. rodentium WT and that colonic hyperplasia, as indicated by colon weight, was decreased (Wickham et al., 2006, Kelly et al., 2006). Subsequently, Newton and colleagues demonstrated the involvement of NleB in dampening of innate immune signaling during A/E pathogen infection by inhibition of NF-κB translocation into host cell nuclei [discussed further in Section 1.4.1; (Newton et al., 2010)]. The mechanism of NleB-mediated inhibition of nuclear NF-κB translocation has yet to be elucidated, but was found to be different from the mechanism used by NleE, which also dampens innate immune signaling (Newton et al., 2010). In A/E pathogens, the nleE gene is present in the same genomic island as nleB and has been shown to play a similar role during infection. A C. rodentium ∆nleE mutant is attenuated for virulence, but to a lesser extent than the nleB mutant (Kelly et al., 2006). This effector is well conserved between EPEC, EHEC, and C. rodentium and in addition to attenuation of colonic hyperplasia and colonization, Wickham and colleagues demonstrated that NleE contributes to mortality in susceptible murine hosts and that inflammation and pathology were decreased in mice infected with C. rodentium ∆nleE (Wickham et al., 2007). Together with NleB and the Shigella effector OspZ, NleE was 13  shown to reduce translocation of the p65 subunit of NF-κB into host cell nuclei (Newton et al., 2010, Nadler et al., 2010). NleE inhibits phosphorylation and degradation of the inhibitor of kappaB alpha (IκBα) via decreasing of IκBα kinase beta (IKKβ) activity and an nleE mutant could not suppress pro-inflammatory signaling and IL-8 secretion as efficiently as WT strains of C. rodentium and EPEC (Newton et al., 2010, Nadler et al., 2010, Zurawski et al., 2008). NleE also inhibits p65 nuclear translocation in DCs and impaired pro-inflammatory responses in these cells [discussed above; (Vossenkamper et al., 2010)]. Although NleE appears to limit pro-inflammatory signaling, infection with C. rodentium ∆nleE resulted in reduced pathology. Surprisingly, production of proinflammatory cytokines in vivo by C. rodentium ∆nleE-infected mice has yet to be examined. The effectors NleC and NleD were previously investigated in parallel based on their close proximity on the EHEC EDL933 genome. Initial investigation of these effectors revealed that EHEC strains with mutations in nleC (∆nleC) or nleD (∆nleD) were not attenuated for persistence in lambs or for A/E lesion formation on human tissue (Marches et al., 2005). Recently, NleC and NleD were found to target specific signaling cascades via encoded HEXXH Zn-metalloprotease motifs (Baruch et al., 2011). NleD functions to cleave host c-Jun N-terminal kinase (JNK) during EPEC infection, which results in decreased JNK-mediated apoptotic signaling (Baruch et al., 2011).  These authors  investigated the role of NleD in reducing of innate immune signaling by EPEC and found that NleD-mediated JNK cleavage did not play a role in suppression of IL-8 secretion by EPEC (Baruch et al., 2011). When the role of NleC was examined, the authors observed increased IL-8 secretion from cells infected with EPEC ∆nleC compared to WT EPEC  14  (Baruch et al., 2011). NleC decreases IL-8 secretion via cleavage of p65 using its HEXXH Zn-metalloprotease motif (Baruch et al., 2011). Subsequently, three separate groups confirmed this phenotype and determined that NleC functions to modulate the host immune response during EPEC infection (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010). NleC also influenced p38 mitogen activated protein kinase (MAPK) signaling and reduced colitis in a murine infection model (Sham et al., 2011). NleC is very highly conserved between A/E pathogens and is homologous to the apoptosis inducing protein 56 kDa (AIP56) from Photobacterium damselae subspecies piscicida (do Vale et al., 2005). Despite homology to a pro-apoptotic toxin, the impact of NleC on apoptotic cell death during A/E pathogen infection has yet to be explored. The function of NleF during A/E pathogen infection remains unidentified. Echtenkamp and colleagues identified NleF as a substrate of the A/E pathogen T3SS and demonstrated that it does not play a significant role in bacterial attachment or A/E lesion formation. However, these authors demonstrated that the C. rodentium ∆nleF mutant was outcompeted by WT C. rodentium and that EHEC ∆nleF was attenuated for virulence in a gnotobiotic piglet model of infection (Echtenkamp et al., 2008). Since nleF is encoded on the same λ–like prophage as a functional nleH1 gene in EPEC and EHEC and nleH in C. rodentium (Garcia-Angulo et al., 2008, Iguchi et al., 2009), it may have a similar or complementary function during A/E pathogen infection. The nleG gene is abundant in genomes of pathogenic E. coli and 14 homologues of this gene are present in EHEC O157:H7 Sakai (Tobe et al., 2006). Very little was known about these proteins until a recent study identified them as a family of E3 ubiquitin ligases (Wu et al., 2010). A RING-finger/U-box motif was identified by solving the 15  structure of the C-terminal region of NleG by nuclear magnetic resonance (NMR) spectroscopy (Wu et al., 2010). The ability of NleG to act as an E3 ubiquitin ligase was confirmed in vitro; however, the contribution of this function to A/E pathogen virulence in vivo has not been determined. The C. rodentium and EHEC effector NleL/EspX7 also functions as an E3 ubiquitin ligase during infection. Using EPEC expressing EHEC nleL, Piscatelli and colleagues found that WT NleL, but not a catalytically inactive form, decreases pedestal formation during EPEC infection. In addition, NleL enhances colitis during C. rodentium infection (Piscatelli et al., 2011). Further contribution of NleL-decreased pedestal formation to A/E pathogen virulence has not been investigated. NleH1 and NleH2 have been shown to contribute to several virulence phenotypes exhibited by A/E pathogens. NleH1 was initially identified by proximity to the nleF gene and subsequently shown to have a homologue encoded elsewhere in the EPEC and EHEC genomes (Garcia-Angulo et al., 2008). The C. rodentium genome encodes a single nleH gene and a ∆nleH strain was attenuated for colonization at early time points in vivo (Garcia-Angulo et al., 2008). The A/E pathogen nleH genes are homologous to the T3S effector OspG from Shigella flexneri, which functions to dampen host NF-κB signaling (Kim et al., 2005).  An elegant in vivo study subsequently demonstrated that C.  rodentium NleH contributes to repression of NF-κB signaling and TNF-α gene expression (Hemrajani et al., 2008). These authors also demonstrated that WT EHEC outcompetes an EHEC ∆nleH1∆nleH2 double mutant for colonization in lambs (Hemrajani et al., 2008). Recently, many advances have been made in understanding the molecular mechanism of NleH action during A/E pathogen infection. The ability of 16  NleH to dampen innate immune signaling has been well demonstrated by two independent studies.  Gao and colleagues demonstrated that NleH1 and NleH2 are  Ser/Thr protein kinases that are auto-phosphorylated and interact with, but do not phosphorylate, ribosomal protein s3 (RPS3) (Gao et al., 2009). They found that NleH1, but not NleH2, dampens RPS3 nuclear localization, which functions to decrease NF-κB activity and IL-8 gene expression in response to tumor necrosis factor (TNF)-α stimulation (Gao et al., 2009). Furthermore, Wan and colleagues revealed that RPS3 phosphorylation by IKKβ is inhibited by NleH1, which causes decreased RPS3 nuclear translocation (Wan et al., 2011).  A related study revealed that NleH1 and NleH2  function similarly to S. flexneri OspG and inhibit ubiquitination and subsequent degradation of IκBα (Kim et al., 2005, Royan et al., 2010). NleH effectors of A/E pathogens function as innate immune regulators, which has been demonstrated in cell culture and in vivo models of infection (Gao et al., 2009, Royan et al., 2010). Several recent studies have also suggested an anti-apoptotic role for NleH1 and NleH2 during EPEC and C. rodentium infection. Hemrajani and colleagues observed increased nuclear condensation following infection of HeLa cells with EPEC strains lacking nleH1 and nleH2 genes and that complementation with either gene alone could restore WT levels of nuclear condensation (Hemrajani et al., 2010). They further demonstrated that procaspase-3 cleavage could be inhibited by NleH and that C. rodentium ∆nleH infection resulted in increased cleavage of procaspase-3.  This phenotype was attributed to  interaction with Bax-inhibitor 1 (BI-1) since depletion of host BI-1 caused increased intracellular Ca2+ levels in host cells during WT EPEC infection (Hemrajani et al., 2010). A similar study demonstrated that NleH1 could protect HeLa cells from Clostridium  17  difficile toxin B-induced apoptosis (Robinson et al., 2010). Furthermore, the ability of NleH1 to protect host cells against apoptotic cell death was influenced by interaction with Na+/H+ exchanger regulatory factor 2 (NHERF2) by an undetermined mechanism (Martinez et al., 2010). These authors also found that NHERF2 influenced trafficking of NleA and Map, and hypothesized that this protein may target effector proteins to specific cellular compartments (Martinez et al., 2010). EspJ is produced by all A/E pathogens and in the C. rodentium genome, espJ is encoded on the same genomic island as nleH (Garcia-Angulo et al., 2008).  However, the  functions found for EspJ to date do not show similarity to those demonstrated for NleH. EspJ does not contribute to A/E lesion formation by EPEC and an espJ mutant is not severely attenuated for virulence in several different infection models (Dahan et al., 2005). However, EspJ was shown to abrogate opsono-phagocytosis of either antibody- or complement-opsonized sheep red blood cells in trans (Marches et al., 2008).  A  subsequent study revealed that EspJ can localize to host mitochondria, despite the absence of a canonical MTS (Kurushima et al., 2010). These authors found that an espJover-expressing EPEC strain was more cytotoxic than WT EPEC in yeast and that cytochrome c was not involved in the enhanced cell death induced by EspJ (Kurushima et al., 2010). EspJ-mediated cytotoxicity towards host cells has yet to be investigated further. EspM and EspT are WxxxE effector proteins and have been demonstrated to subvert host cytoskeletal dynamics. The espT gene is present in C. rodentium but is absent from most EPEC and EHEC strains (Arbeloa et al., 2009). However, the function of EspT was investigated using an EPEC strain able to secrete EspT via the T3SS (Bulgin et al., 18  2009b). Bulgin and co-workers found that EspT can induce formation of membrane ruffles and lamellopodia on the surface of host cells via activation of Cdc42 and Rac-1 GTPases (Bulgin et al., 2009b). A subsequent study demonstrated that expression of espT could induce EPEC internalization into cultured cells (Bulgin et al., 2009a) and very recently, EspT was found to facilitate increased production of the pro-inflammatory cytokines IL-8, IL-1β, and TNF-α via nuclear factor-kappa B (NF-κB) and extracellular signal-regulated kinase 1/2 (Erk1/2)/JNK signaling (Raymond et al., 2011). In the strains used for these studies, espT is expressed from a plasmid in a pathogen that does not normally encode it and resultant phenotypes may result from aberrant regulation of other effector proteins or may be a by-product of effector over-expression. The authors suggest that strains encoding espT may use up-regulation of immune mediators as a pathogenic mechanism (Raymond et al., 2011); however, the temporal regulation of espT expression and function has not been examined. Unlike espT, the espM gene and its homologues are well distributed throughout the genomes of EPEC, EHEC, and C. rodentium although EPEC E2348/69 lacks both genes (Arbeloa et al., 2009, Arbeloa et al., 2008). Both O157 EHEC strains and C. rodentium ICC168 carry two homologous espM genes and EPEC B171 contains one espM gene (Arbeloa et al., 2008). Arbeloa and colleagues used EPEC E2348/69 transformed with plasmids encoding espM to examine the function of EspM effectors in culture. They found that EspM could activate RhoA/Rho-associated, coiled-coil containing protein kinase 1 signaling and facilitate actin stress-fiber formation (Arbeloa et al., 2008). This group subsequently demonstrated that EHEC EspM2 is a guanine nucleotide exchange factor and uses this function to activate RhoA (Arbeloa et al., 2010). A separate group  19  probed the function of espM gene products on EHEC infection using espM1 and espM2 deletion strains (Simovitch et al., 2009). An EHEC ∆espM1∆espM2 double mutant produced much larger pedestals on the surface of infected HeLa cells and infection with EPEC expressing espM1 and espM2 resulted in reduced pedestal formation as well as an enhanced presence of actin stress-fibers (Simovitch et al., 2009). These authors also demonstrated that EspM effectors perturb the organization of a polarized cell monolayer and cause mislocalization of tight junctions without disrupting barrier function (Simovitch et al., 2009). Examining the function of effectors that are not ubiquitously expressed in A/E pathogens may aid in understanding disease manifestations and pathogenic mechanisms associated with specific A/E pathogen infections. The identification of EspV occurred very recently and little is known about its function. The espV gene is present in C. rodentium and some clinical isolates of EPEC and EHEC, but was cloned into EPEC E2348/69 prior to phenotypic investigation (Arbeloa et al., 2011). C. rodentium ∆espV was not attenuated for colonization in C57BL/6 mice and WT C. rodentium did not have a competitive advantage over the ∆espV strain (Arbeloa et al., 2011). When translocated by the EPEC T3SS or expressed by HeLa cells, EspV induced cell rounding and formation of dendrite-like appendages (Arbeloa et al., 2011). The relevance of this phenotype is unknown, but the authors suggest that EspV may be altering the cell cycle. In addition, Arbeloa and colleagues found that EspV, when expressed in yeast, was cytotoxic. The ability of EspV to cause cell death in mammalian cells was not examined in the study. The Cif effector from EPEC and EHEC has been shown to perform two main functions during infection and has been crystallized (Hsu et al., 2008). Crystallization revealed the 20  presence of a papain-like fold and catalytic triad (Cys109-His165-Gln185) (Hsu et al., 2008). The effector initially gained its name by its ability to inhibit the G2/M transition of the cell cycle in eukaryotic cells (Marches et al., 2003). The mechanism of cell cycle inhibition by Cif involves inhibition of cyclin dependent kinase 1 activation independent of deoxyribonucleic acid (DNA) damage (Marches et al., 2003, Taieb et al., 2006). Further studies revealed that cell cycle arrest by Cif was mediated by accumulation of cyclin kinase inhibitors p21waf1 and p27kip1, which was dependent on the catalytic triad, and inhibition of their proteasome-dependent degradation (Samba-Louaka et al., 2008). Subsequently, Cif was shown to inhibit the Skp1-Cullin1-F-box-protein ubiquitin ligase, which facilitated accumulation of cycling inhibitory proteins (Morikawa et al., 2010, Cui et al., 2010). Inhibition of the cell cycle may result in late-stage apoptosis, which is induced by Cif and dependent on the catalytic triad (Samba-Louaka et al., 2009). Interestingly, cif is not encoded on the C. rodentium genome (Loukiadis et al., 2008) and its role in vivo has not been examined. Proteomic and bioinformatic studies have revealed a much larger than expected repertoire of effector proteins secreted by A/E pathogens (Deng et al., 2010, Tobe et al., 2006). Despite the similar pathogenesis exhibited by different A/E pathogens, several strainspecific effectors exist and it is very tempting to speculate that each individual strain has evolved its own repertoire to efficiently cause disease and avoid host responses. Further investigation will likely reveal an even larger set of effector proteins used by A/E pathogens to cause diarrheal disease.  21  1.3 Subversion of host cell death pathways Host cell death is a major consequence of A/E pathogen infection (Wong et al., 2011). Over a decade of investigation has revealed that A/E pathogens can intricately regulate host cell death during infection and this may be a major virulence mechanism employed by these pathogens.  Host cell death caused by A/E pathogens has been studied  extensively in cell culture models and has characteristics of both apoptosis and necrosis and involves several T3S effector proteins. 1.3.1 Apoptosis Apoptotic cell death is a well-defined hallmark of A/E pathogen infection. Apoptosis falls into two categories; intrinsic – initiated by intracellular stimuli, and extrinsic – facilitated by cell surface receptors [for review see (Fulda et al., 2006)].  Intrinsic  apoptosis is mediated by outer mitochondrial membrane (OMM) permeabilization followed by release of mitochondrial proteins and second messengers into the cell cytosol that bind cognate receptors and initiate apoptotic signaling; however, extrinsic apoptosis results from specific membrane receptor engagement – such as the TNF-α receptor - on the surface of cells (Fulda et al., 2006). A major consequence of both intrinsic and extrinsic apoptosis is activation of a family of cysteine proteases called caspases [for review see (Moffitt et al., 2010)]. Both forms of apoptosis result in activation of caspase3, which facilitates DNA damage and cell shrinkage by degradation of several substrates (Porter et al., 1999). Cell shrinkage results in membrane “blebbing” and the formation of small apoptotic bodies containing condensed chromatin, which are internalized by professional phagocytes (Aderem et al., 1999). In the absence of phagocytes, a process  22  called secondary necrosis occurs whereby the apoptotic bodies rupture (Festjens et al., 2006). A/E pathogens have been shown to instigate both intrinsic and extrinsic apoptosis by various mechanisms. Baldwin and colleagues first demonstrated loss of viability, via increased intracellular Ca2+ levels, in Hep-2 cells infected with EPEC (Baldwin et al., 1993). Subsequent studies in cell culture have revealed that EPEC infection results in characteristic phenotypes  associated  with  apoptosis,  such  as  outer  plasma  membrane  phosphatidylserine exposure, DNA fragmentation, nuclear condensation, membrane blebbing, release of cytochrome c, and activation of caspase-3 (Flynn et al., 2008, Barnett Foster et al., 2000, Crane et al., 1999).  Although effector proteins play a role in  apoptosis induction during A/E pathogen infection, there are several other virulence factors that cause apoptosis independently of the T3SS.  BFP and outer membrane  proteins of A/E pathogens also mediate apoptotic cell death (Abul-Milh et al., 2001, Shankar et al., 2009). Interestingly, EPEC outer membrane proteins can activate both the intrinsic and extrinsic apoptotic cascades by activation of JNK and by increasing expression of TNF-α, respectively (Shankar et al., 2009). No effector proteins have been shown to signal the extrinsic pathway, likely because they are translocated directly into host cells. Apoptotic cell death of IECs in vivo has been demonstrated following C. rodentium infection. Infection of susceptible mice with C. rodentium resulted in increased apoptosis in crypt IECs (Vallance et al., 2003). Crypt cell apoptosis has also been observed in biopsies of EHEC patients (Griffin et al., 1990). T3S proteins shown to cause apoptotic cell death play important roles in virulence in vivo. C. rodentium mutant strains lacking 23  either map or espF are attenuated for colonization and mortality in susceptible murine strains (Nagai et al., 2005, Deng et al., 2004). This suggests that apoptotic cell death is an important virulence strategy employed by A/E pathogens and several authors have speculated that apoptotic cell death may facilitate pathogen dissemination in the absence of overt inflammation. The presence of anti-apoptotic effectors suggests that apoptotic cell death is highly regulated by A/E pathogens. As discussed above, the T3S effectors NleH1, NleH2, and NleD antagonize apoptotic cell death by separate mechanisms (Baruch et al., 2011, Hemrajani et al., 2010). It is tempting to speculate that these effectors delay apoptotic cell death during A/E pathogen infection to facilitate greater pathogen colonization prior to dissemination. Unlike pro-apoptotic effectors, removal of these proteins from the T3S effector repertoire doesn’t result in strong virulence defects in vivo and it’s likely that they play redundant roles to avoid premature death of host cells during infection. Further understanding of the interplay between pro- and anti-apoptotic effectors is required to discern the role of apoptosis during A/E pathogen infection. 1.3.2 Necrosis In addition to the body of knowledge demonstrating that A/E pathogens hijack apoptotic cell death cascades, there is also evidence for subversion of necrotic cell death. Cell death by necrosis is generally described as an inflammatory and non-regulated form of cell death characterized by mitochondrial swelling, nuclear swelling and early plasma membrane rupture (Festjens et al., 2006, Golstein et al., 2007). Many mitochondrial alterations can result in necrotic cell death, such as the loss of mitochondrial inner  24  membrane potential (Δψm) (Lemasters et al., 1999). Sustained loss of Δψm results in release of mitochondrial components into the cellular cytosol, such as reactive oxygen species (ROS) (Lemasters et al., 1999, Festjens et al., 2006). ROS include superoxide radicals and hydrogen peroxide, which are highly unstable and can rapidly and irreversibly reduce cellular proteins and high levels of ROS resulting from mitochondrial dysfunction in the cytosol can result in necrotic cell death (Lemasters et al., 1999, Festjens et al., 2006). Identification of necrotic cell death relies on fewer characteristics than apoptosis and morphological analysis and cell membrane rupture are still widely used to characterize this form of cell death. Initially, Crane and colleagues demonstrated that EPEC infection resulted in plasma membrane permeability at very early time points p.i., indicative of necrosis (Crane et al., 1999). Although apoptotic cell death has been more frequently observed, A/E pathogens likely cause a combination of both forms of cell death in vivo. Based on their intricate regulation of the immune system (discussed below), A/E pathogens likely limit necrotic cell death prior to shedding from the intestinal epithelium. The work in Chapter 2 and Chapter 3 of this thesis details potential mechanisms for how A/E pathogens limit necrotic cell death during infection.  1.4 Subversion of innate immunity Diarrhea resulting from A/E pathogen infection is largely dependent on disruption of epithelial barriers in the host. Permeability in the intestinal epithelial layer results in lumenal products entering the underlying mucosa and stimulates recruitment of pro-  25  inflammatory cells, such as neutrophils. Interaction of bacterial pathogens with IECs also results in pro-inflammatory signaling and neutrophil recruitment.  Specifically,  expression of pathogen associated molecular patterns (PAMPs) by bacterial pathogens triggers innate immune responses from host cells. A/E pathogen flagellin is a particularly potent immune activator and facilitates immune signaling from Toll-like receptor (TLR) 5 (Khan et al., 2008). During infection, A/E pathogens actively dampen innate immune signaling, which would normally lead to increased inflammation and pathogen clearance. Several strategies for survival in the host intestinal environment have been described and A/E pathogen-mediated evasion of the host immune system at the level of epithelial and immune cells has been documented. The eventual response to A/E pathogens in resistant hosts is immune clearance of the bacteria; however, sufficient colonization and pathogen shedding has likely occurred thus increasing the potential for dissemination and transmission to naïve hosts (Sharma et al., 2006). 1.4.1 Epithelial cells Upon exposure to bacterial pathogens, host IECs detect PAMPs, which results in activation of signaling cascades leading to many downstream events. A major response of IECs is production of the pro-inflammatory chemokine IL-8. At steady state, IL-8 levels are below detection; however, the presence of bacterial PAMPs, IL-1, or TNF-α can stimulate its production (Hoffmann et al., 2002).  Three general mechanisms  contribute to transcription of the IL8 gene; (1) accessibility of the promoter region by opening of chromatin, (2) activation of gene transcription, and (3) MAPK-mediated stabilization of IL-8 mRNA (Hoffmann et al., 2002).  26  A/E pathogens have been  demonstrated to influence each of these mechanisms to dampen levels of secreted IL-8 and therefore subvert the innate immune response. Eukaryotic cells use several mechanisms to regulate gene expression including altering accessibility of gene promoter elements. Inert DNA is tightly wrapped around histone proteins, which have N-terminal “tails” that form electrostatic interactions with the negatively charged phosphate backbone of DNA. To increase accessibility to promoter regions, histone tails are modified by several mechanisms, which can enhance or repress their association with DNA.  Specifically, chromatin remodeling at the IL-8 gene  promoter involves acetylation of histone tails on positively charged Lys residues by the transcriptional co-activators cAMP response element binding (CREB)-binding protein (CBP) and p300, which are homologous histone acetyltransferases (HATs) that are recruited to the IL-8 gene promoter region by PAMPs, TNF-α, and/or IL-1 stimuli (Hoffmann et al., 2002, Roebuck, 1999). CBP/p300 hyper-acetylate histone tails, which enables transcription factors such as NF-κB dimers to facilitate transcription (Hoffmann et al., 2002). This thesis describes a mechanism by which the T3S effector NleC actively degrades the co-activator p300, which results in decreased IL-8 secretion (see Chapter 5). The data presented are the first to demonstrate that A/E pathogens alter the function of transcriptional co-activators. Although several transcription factors are involved in IL-8 gene transcription, only NFκB is essential (Kunsch et al., 1993, Hoffmann et al., 2002). Not surprisingly, A/E pathogens have evolved several mechanisms by which to repress NF-κB at early time points during infection. IL-8 gene expression is generally facilitated by the p50-p65 NFκB heterodimer, which interacts with a κB-enhancer element on exposed promoters. 27  Inactive NF-κB dimers are maintained in the host cell cytosol by the inhibitor of κB alpha (IκBα) and upon NF-κB activation by upstream signaling, IκBα is phosphorylated by IκB kinase (IKK), which leads to its ubiquitination and subsequent degradation by the 26S proteasome. NF-κB dimers are then phosphorylated and can translocate into host cell nuclei where they can interact with IL8 κB-enhancer elements and facilitate recruitment of RNA polymerase II (Roebuck, 1999, Hoffmann et al., 2002). NF-κB signaling is required for eventual A/E pathogen clearance in vivo. Mice deficient for the p50 subunit (p50-/-) cannot efficiently clear C. rodentium infection and experience increased colitis compared to p50+/+ mice (Dennis et al., 2008). Not surprisingly, A/E pathogens are proficient at disabling NF-κB-mediated gene expression in a T3SSdependent manner (Ruchaud-Sparagano et al., 2007, Hauf et al., 2003).  To date,  suppression of NF-κB activity has been attributed to several Nle effector proteins. NleE, NleB, NleH1/2, and NleC all contribute to suppression of NF-κB activity. Pearson and colleagues demonstrated that NleE and NleC function together to impair NF-κB (Pearson et al., 2011). NleE retains NF-κB dimers in the nucleus (described above) where NleC may then proteolyze p65. NleC will also degrade p65 that has entered the host cell nucleus resulting in further inhibition of NF-κB-mediated gene expression (Baruch et al., 2011). It is clear from this body of work that A/E pathogens utilize several redundant strategies to reduce NF-κB activity [reviewed in (Wong et al., 2011)]. This suggests that decreasing NF-κB activity is an important virulence mechanism for A/E pathogens. Instability of IL-8 mRNA is a regulatory mechanism to avoid unnecessary inflammation in host tissues.  The 3’ untranslated region of IL-8 mRNA, like that of other pro-  inflammatory cytokines, contains an AU-rich element that decreases the stability of 28  mRNA molecules (Shaw et al., 1986, Winzen et al., 1999). Winzen and colleagues determined that active p38 MAPK was necessary for IL-8 mRNA stability following stimulation with IL-1 and that downstream activation of the MAPK activated kinase 2 was necessary (Winzen et al., 1999). A/E pathogen flagellin mediates activation of p38 MAPK; however, p38 MAPK signaling is actively dampened by the T3S effector NleC [discussed below; (Sham et al., 2011)]. Although stabilization of IL-8 mRNA by p38 is not essential for IL-8 secretion by epithelial cells (Hoffmann et al., 2002), its suppression by A/E pathogens contributes to dampening the innate immune response during infection. 1.4.2 Immune cells Interactions between A/E pathogens and immune cells likely occurs via both sampling of the intestinal lumen by dendritic cells (DCs) and through bacterial invasion of the underlying mucosa following IEC barrier disruption. neutrophils, macrophages, and DCs.  Innate immune cells include  Neutrophils are short-lived effector cells that  function to cause increased inflammation at sites of infection. Infiltration of neutrophils generally results in tissue damage and decreased barrier integrity but also pathogen clearance (Lebeis et al., 2007). Although direct interaction between A/E pathogens and neutrophils has not been documented, recruitment of neutrophils during A/E pathogens infection has been observed.  Several in vivo studies using C. rodentium have  demonstrated that neutrophil recruitment is required for pathogen clearance (Khan et al., 2006, Lebeis et al., 2007, Spehlmann et al., 2009). Macrophages and DCs are professional phagocytes that function as antigen presenting cells (APCs). Following engagement of specific surface receptors that recognize either  29  PAMPs, complement component C3bi or Fc regions of antibodies, particles are internalized and degraded in phagosomes.  Peptides from the degraded proteins are  displayed on the surface of the APC and can subsequently facilitate an adaptive immune response by activating T cells. Recently, EPEC infection of human monocyte-derived DCs was documented. Vossenkämper and colleagues demonstrated that EPEC can infect and translocate effector proteins into DCs. EPEC infection dampened secretion of proinflammatory cytokines by decreasing translocation of NF-κB dimers into the cell nuclei (Vossenkamper et al., 2010). The authors found that EPEC ∆nleE was impaired for NFkB suppression, suggesting that NleE functions similarly in epithelial cells and DCs (Vossenkamper et al., 2010). In addition, DCs co-cultured with WT EPEC could not efficiently activate T cells in a mixed lymphocyte reaction (Vossenkamper et al., 2010). It is likely that WT EPEC also prevents its own uptake by DCs and therefore limits their ability to activate an adaptive immune response. Activation of host macrophages is dependent on the presence of pro-inflammatory cytokines and uptake of invading pathogens. Macrophages are also APCs that can present antigen to cells of the adaptive immune system and facilitate their activity. Phagocytosis by macrophages is essential for their effector function and uptake is dependent on the presence of receptor ligands. Macrophages possess receptors to bind PAMPs, complement-opsonized particles (complement receptor 3; CR3), and antibody opsonized particles (Fc gamma receptor; FcγR). Several studies have demonstrated that EPEC can inhibit phagocytic uptake by macrophages (Dong et al., 2010, Marches et al., 2008, Iizumi et al., 2007, Goosney et al., 1999, Celli et al., 2001). Celli and colleagues demonstrated that EPEC, in a T3SS-dependent manner, could inhibit phosphoinositide-3  30  kinase (PI3K)-mediated activation of AKT in macrophages, which resulted in decreased uptake of EPEC and Fc-coated beads (Celli et al., 2001). In addition, Marchès and colleagues observed EPEC-mediated decrease in macrophage uptake of C3bi- and Fcopsonized red blood cells and that EspJ was responsible for this phenotype (Marches et al., 2008). Since clearance of C. rodentium is dependent on B cells and the presence of FcγR (Simmons et al., 2003, Masuda et al., 2008, Maaser et al., 2004), inhibition of phagocytosis likely occurs early during infection and may provide time for increased A/E pathogen colonization prior to clearance. In addition to EspJ, EspF and EspB also function to dampen macrophage phagocytosis. EspF inhibits phagocytosis of EPEC via an undefined mechanism independent of AKT phosphorylation (Quitard et al., 2006). EspB also contributed to inhibition of macrophage phagocytosis independent of its translocator activity but by an unknown mechanism (Iizumi et al., 2007).  The  identification of three effectors that actively dampen phagocytosis suggests that this is an important pathogenic strategy during infection in vivo.  1.5 Summary of thesis Understanding the roles of effector proteins during A/E pathogen infection has provided insight into their virulence mechanisms and how they have evolved to efficiently cause disease.  Despite the expanding amount of literature focused on effector protein  functions, the roles of most effector proteins have yet to be elucidated. Identifying a role for effector proteins has classically been performed by mutagenesis of single effectors or gene clusters, protein sequence or structural analysis, or by identification of host binding  31  partners. The studies presented in this thesis were initiated by identification of host protein binding partners using stable isotope labeling of amino acids in cell culture (SILAC) and phenotypic analysis during infection of cultured cells. Many of the LEE-encoded effector proteins have been well studied; however, the function of EspZ had not been investigated. Since EspZ contributes to full virulence of C. rodentium in vivo (Deng et al., 2004), its role during EPEC infection in culture was investigated here. Chapter 2 details the function of EspZ as a pro-survival mediator during EPEC infection of cultured cells. Using SILAC and yeast two-hybrid (Y2H), EspZ was shown to interact with CD98 and this interaction contributed to pro-survival signaling via FAK activation.  The ability of EspZ to antagonize EPEC-mediated  cytotoxicity also relies on the host inner mitochondrial membrane translocase protein TIM17B, as described in Chapter 3. Subversion of innate immune responses during A/E pathogen infection relies on several T3S effector proteins encoded outside the LEE. NleC dampens innate immune signaling by limiting NF-κB activity and IL-8 secretion as shown in Chapter 4. As described above, NleC is a Zn-metalloprotease that degrades the p65 subunit of NF-κB, resulting in decreased IL-8 secretion (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011). Chapter 5 details a second mechanism by which NleC dampens innate immune signaling by degradation of the host acetyltransferase p300. EspZ and NleC both contribute to A/E pathogen-mediated disease and uncovering their functions during infection has shed light on mechanisms by which A/E pathogens can subvert host cell death and innate immune signaling. Determining the roles of T3S  32  effector protein will enable enhanced understanding of the virulence mechanisms of pathogenic bacteria and the host cell processes that are manipulated.  33  Chapter 2. The A/E pathogen effector EspZ interacts with host CD98 and facilitates host cell pro-survival signaling.  2.1 Introduction EspZ is a LEE-encoded effector protein that is present in a single copy in EPEC, EHEC, and C. rodentium, and has approximately 60% sequence identity among these pathogens (Kanack et al., 2005). Our laboratory previously demonstrated that a C. rodentium ∆espZ is severely attenuated for virulence and does not colonize efficiently in vivo (Deng et al., 2004). However, EPEC ∆espZ can adhere to WT levels during infection in tissue culture (Kanack et al., 2005). Despite its critical role in virulence, the host binding partner(s) and function of EspZ had not been elucidated. To gain insight into the function of EspZ, we aimed to identify host protein binding partners using Y2H and quantitative proteomics techniques.  SILAC has been used  extensively as a quantitative method to identify protein-protein interactions using differential labeling of cellular proteomes with stable isotopes of essential amino acids (Ong et al., 2003). Rogers and colleagues first used SILAC to identify binding partners to the S. Typhimurium T3S effector SopB/SigD (Rogers et al., 2008). Recently, an effector-wide screen of all S. Typhimurium effector proteins was undertaken using complementary SILAC methods (Auweter et al., 2011). Protein-protein interactions uncovered by this technique can provide valuable insight into the function of effector proteins.  34  The host protein CD98 (4F2hc, SLC3A2) was identified as a binding partner to EHEC EspZ in our study (Shames et al., 2010). CD98 is a type-II transmembrane glycoprotein that has several functions in eukaryotic cells including amino acid transport and regulation of β1-integrin signaling (Yan et al., 2008).  CD98 is responsible for  localization of L-type amino acid transporters LAT-2 and y+LAT1 to the basolateral surface of polarized intestinal epithelial cells via di-sulfide bonds between the proteins (Yan et al., 2008).  The CD98/LAT-2 heterodimer co-localizes with β1-integrin in  polarized epithelia, and has been shown to influence signaling downstream of β1containing integrin heterodimers (Kolesnikova et al., 2001, Rintoul et al., 2002, Cai et al., 2005, Feral et al., 2005); however, interaction with the LAT-2 light chain subunit is not necessary for integrin regulation (Yan et al., 2008). Regulation of β1-integrin by CD98 plays an important role in cell proliferation, survival, and apoptosis. Feral and colleagues demonstrated that CD98 activates PI3K and Rac and promotes cellular proliferation and increases tumor sizes in vivo (Feral et al., 2005). In addition, the β1-integrin-CD98 interaction promotes activation of pro-survival kinases AKT and FAK and clustering of CD98 exacerbates this phenotype (Rintoul et al., 2002, Cai et al., 2005). To gain insight into the function of EspZ, we examined its role during infection of cultured cells. The EPEC ∆espZ mutant caused increased cytotoxicity in HeLa and MDCK cells compared to WT EPEC, which was abrogated by expression of espZ in both cell lines. The ability of CD98 to influence host cell survival and resultant activation of signaling proteins during EPEC infection was also examined.  35  2.2 Experimental procedures 2.2.1 Tissue culture, bacterial strains, transfection, and infection conditions HeLa (ATCC), Madin-Darby Canine Kidney (MDCK) (ATCC), and HEK 293 cells (ATCC) were maintained in Dulbecco’s Modified Eagle Medium (DMEM) High Glucose (Thermo Scientific) supplemented with 10% fetal bovine serum (FBS) (Thermo Scientific),  1%  non-essential  amino  acids  (NEAA)  (Gibco),  0.1  Units/L  Penicillin/Streptomycin (Penn/Strep) (Gibco), and 1% GlutaMax (Gibco). Cells were used from passages 5-20. Bacterial strains used in this study are listed in Table 2.1, oligonucleotide primers used in this study are listed in Table 2.2, and plasmids used in this study are listed in Table 2.3. EHEC O157:H7 strain EDL933 was used for cloning espZ as described below. EPEC colonies from fresh Luria-Bertani (LB) agar plates supplemented with 50 µg/mL streptomycin were used to inoculate 3 mL LB broth and incubated overnight at 37°C without shaking. HeLa cells were washed in PBS and infected at a multiplicity of infection (m.o.i.) of ~100:1 with EPEC cultures in serum-free DMEM. Mock-infected cells were treated with pre-warmed, sterile LB broth. Infections were allowed to proceed for indicated times. HeLa cells were transiently transfected using FuGENE HD (Roche) with a ratio of 2:3 (µg DNA: µL reagent) in DMEM according to manufacturer’s instructions. Media were  36  changed 24 h post-transfection and cells were infected or assayed 48 h post-transfection. Lysates were subject to Western blot analysis to confirm transfection efficiency. HeLa-EspZ and MDCK-EspZ stable cells were made by transfection of pEHespZ-GFPN separately into HeLa and MDCK cells using Lipofectamine 2000™ (Invitrogen) according to manufacturer’s instructions. Stable clones were selected using DMEM + 600 µg/mL G418 + 10% FBS + 1% Penn/Strep for 2 weeks then maintained in DMEM containing 10% FBS and 300 µg/mL G418 until use. HEK 293 cells were transfected using calcium phosphate as described (Stone et al., 1991). Cells were assayed 48 h post-transfection. Table 2.1 Bacterial strains used in this study Name Source/Reference EPEC O127:H6 strain E2348/69 Wild-type (WT) (Levine et al., 1978) ∆espZ (Shames et al., 2010) ∆escN (Gauthier et al., 2003) ∆sepD (Deng et al., 2005) ∆sepD∆espZ (Thomas et al., 2007) ∆sepD∆espZ/pespZ This study EHEC O157:H7 strain EDL933 WT (Johnson et al., 1984) Escherichia coli SM10λpir (Miller et al., 1988) DH10B Invitrogen BL21 (DE3) Stratagene  2.2.2 Generation of CD98-FLAG fusions CD98 cDNA was obtained from clones from the BD Matchmaker™ pre-transformed HeLa cell cDNA library (BD Clontech) following Y2H experiments. The CD98 cDNA  37  lacking its stop codon was cloned into pCMV-TAG4A (pCMV4A). Primers CD98K-F and CD98-R were used to PCR amplify CD98 cDNA with a Kozak sequence on the 5’ end.  The purified PCR product was cloned as a BamHI/HindIII fragment into the  multiple cloning site (MCS) of pCMV4A in frame with the downstream FLAG-tag sequence.  2.2.3 Generation of pcDNA3::HA2 and pcDNA3::HA2espZ constructs A double HA tag downstream of a Kozak sequence was inserted into pcDNA3. Complementary oligonucleotides HA2-F and HA2-R were annealed followed by cleavage with HindIII. The HindIII fragment was cloned into HindIII-digested pcDNA3. Following ligation, a directional clone was identified by sequencing (NAPS Unit, UBC) to create the vector pcDNA3::HA2. The coding region of EHEC EDL933 espZ was PCR amplified using forward primer CdH37 and reverse primer CdH38 and cloned into pcDNA3::HA2.  Briefly,  pcDNA3::HA2 was cleaved with SrfI and NotI followed by ligation of a SrfI/NotI fragment of espZ. We achieved in-frame cloning of HA2espZ by adding one arginine (Arg) residue (CGG) between the second HA tag and espZ. For SILAC, this ensured the presence of a labeled Arg residue following tryptic digestion.  38  2.2.4 EPEC ∆espZ mutant generation The sacB gene-based allelic exchange method was used to generate an espZ in-frame deletion mutant in the streptomycin-resistant derivative (Smr) of EPEC O127:H6 strain E2348/69 using the suicide vector pRE112. PCR was used to generate two fragments (1.4 and 1.8 kb, respectively) using primer pairs EPespZ-1 and DEPespZ-R as well as DEPespZ-F and EPespZ-2. The PCR products were cloned into pCR2.1-TOPO and verified by DNA sequencing. After digestion with KpnI/NheI and NheI/SacI, respectively, the two fragments were gel-purified and cloned into pRE112 digested with KpnI/SacI in a 3-way ligation. The resulting plasmid pRE-∆EPespZ contained the espZ gene with an internal in-frame deletion from nucleotides 43 to 273 (about 79% of the coding region) and more than 1 kb of flanking regions on both sides of espZ. An NheI site was introduced into the deletion site. Plasmid pRE-∆EPespZ was transformed into E. coli SM10λpir by electroporation, and introduced into EPEC strain E2348/69 Smr by conjugation. After sucrose selection as previously described (Edwards et al., 1998), EPEC colonies resistant to sucrose and sensitive to chloramphenicol were screened for deletion of espZ by PCR using primers EPespZ-1 and EPespZ-2. The EPEC espZ mutant was further verified by PCR.  2.2.5 Green fluorescent protein (GFP) fusion to EHEC EspZ N-terminal GFP fusions to espZ of EHEC O157:H7 strain EDL933 were constructed using the vector pEGFP-N1. The coding region of EHEC espZ was amplified by PCR using primers EHespZ-NF and EHespZ-NR, cloned into pCR2.1-TOPO and verified by  39  DNA sequencing, and then sub-cloned as an XhoI/BamHI fragment into pEGFP-N1 to generate a fusion to the N-terminus of GFP (pEHespZ-GFPN). To truncate EHEC EspZ into its putative transmembrane segments lacking the N-terminal head, the N-terminal region (amino acid residues 1-42) and C-terminal membranespanning region (amino acid residues 43-99) were also cloned into pEGFP-N1 to generate C-terminal GFP fusions. Plasmid pEHespZNGFP was generated using EHespZNF and EHespZ-NR2, and primers EHespZ-NF2 and EHespZ-NR were used to generate pEHespZCGFP, respectively, in pEGFP-N1.  These constructs of EHEC EspZ-GFP  fusions were used to transfect HeLa or MDCK cells as described above.  2.2.6 Y2H assay The coding region of EHEC EDL933 espZ, including the stop codon, was amplified by PCR using primers EHespZ-F and EHespZ-R. The resultant PCR fragment was cloned into pCR2.1-TOPO and verified by DNA sequencing followed by sub-cloning into the NdeI/EcoRI sites of the yeast-two hybrid GAL4 DNA binding domain vector pGBKT7 to generate “bait” plasmid pGBK-EHespZ.  The bait plasmid was tested for protein  expression and subsequently used to screen a pre-transformed human HeLa cDNA library for proteins interacting with EspZ according to the manufacturer’s instructions in the BD MatchmakerTM Pre-transformed Libraries User Manual (BD Biosciences Clontech). Putative positive yeast clones containing library plasmids encoding human proteins interacting with EspZ were purified by re-streaking on the selective media, retested for their growth phenotypes and verified according to the User Manual. Crude genomic  40  DNA preparations for these yeast clones were generated by re-suspending the yeast cells in 50 µL of H2O, boiling for 10 min, incubating on ice for 10 min, and treating the samples with 1 µL of 10 mg/mL of RNase A for 15 min at room temperature. The supernatant of the yeast extract was used to analyze the cDNA inserts in the library plasmids by PCR amplification and primers AD/LD-F and AD/LD-R. The PCR products were cleaned up using the Qiagen PCR Purification Kit (Qiagen), and sequenced using the commercially available T7 sequencing primer. The cDNA sequences were used to search GenBank and the human genome using the NCBI BLAST server. Table 2.2 Oligonucleotide primers used in this study Name Sequence (5ʹ′3ʹ′)a,b CD98K-F (KpnI) attggatccgccaccatgagccaggacaccgaggtggatatgaagg CD98-R (HindIII) attaagcttggccgcgtaggggaagcggagcagcagccc HA2-F (HindIII) cccaagcttgccaccatgtacccatacgatgttccagattacgcttacccatacgatgttcca gattacgctgcccgggcaagcttccc HA2-R (HindIII) gggaagcttgcccgggcagcgtaatctggaacatcgtatgggtaagcgtaatctggaacat cgtatgggtacatggtggcaagcttggg CdH37 (SrfI) ggatggaagcagcaaatttaagtcc CdH38 (NotI) catgcggccgcttaggcatatttcatcgctaatgc EPespZ-1 (KpnI) gcggtacctgcttgtcgagcaacgaggcg DEPespZ-R (NheI) ccgctagcggattagcgatgaaatatgcc DEPespZ-F (NheI) gcgctagctggtaatactgcaccagaagg EPespZ-2 (SacI) ccgagctcgagtatctttgtatattgactc EHespZ-NF (XhoI) cctcgagatggaagcagcaaatttaagtcc EHespZ-NR (BamHI) cggatccgcatatttcatcgctaatgcacc EHespZ-NR2 (BamHI) cggatccgcatatttcatcgctaatgcacc EHespZ-NF2 (XhoI) cctcgagatgagagttatagccggattagcac EHespZ-F (NdeI) ggcccatatgatggaagcagcaaatttaagcc EHespZ-R (EcoRI) ggccgaattcttaggcatatttcatcgc AD/LD-F ctattcgatgatgaagataccccaccaaaccc AD/LD-R gtgaacttgcggggtttttcagtatctacgatt a – Restriction endonuclease cleavage sites are underlined. b – Kozak sequences are bolded.  41  2.2.7 SILAC HEK 293 cells were maintained and SILAC labeled as described previously (Rogers et al., 2008). Briefly, HEK 293 were split from normal growth media into SILAC labeling media. DMEM lacking arginine and lysine (Caisson Laboratories Inc.) was supplemented with 10% dialyzed FBS, 1% NEAA, 1% GlutaMax, 0.1 units/L Penicillin/Streptomycin and either 36.5 mg/L  13  C6-arginine and 2H4-lysine (Cambridge Isotope Laboratories,  Andover, MA) for heavy cells or normal isotopic abundance L-arginine and L-lysine for light cells (Sigma-Aldrich). Cells were maintained for five cell doublings to ensure complete labeling of the proteome and ten 10 cm cell culture dishes of each light and heavy labeled cells were transfected for 18-24 h each with 8 µg of pcDNA3::HA2espZ or pcDNA3::HA2 DNA, respectively, using calcium phosphate as described (Stone et al., 1991). Transfected cells were solubilized in 750 µL cold NP-40 lysis buffer [1% Nonidet P40 (NP40) (v/v), 20 mM Tris pH 7.5, 150 mM NaCl, 10 mM Na pyrophosphate, 50 mM NaF supplemented with 1 mM Na3VO4, and Protease Inhibitor Cocktail (Roche)] and centrifuged at 16,100 relative centrifugal force (r.c.f.) for 10 min at 4ºC.  Protein  concentrations of heavy and light lysates were measured using the Coomassie Plus Protein Assay (Pierce Scientific) according to manufacturer’s directions and then equal quantities of heavy and light lysates were mixed and pre-cleared on Protein G Sepharose™ 4 Fast Flow beads (GE Healthcare) with 150 µg mouse IgG (Jackson ImmunoResearch Laboratories) for 30 min at 4ºC with rocking. Pre-cleared lysates were spun and supernatant was transferred into a new tube. Beads bound to mouse α-HA antibody (8 µg/transfected plate) were added to the cold lysate and incubated at 4ºC for 2 h with rocking. Beads were washed three times with 1 mL of cold lysis buffer and after  42  the final wash, all remaining buffer was drawn from the beads using GELoader Tips (Eppendorf, Westbury, NY) until the beads were dry. At this point, 150 µL of elution buffer [6 M urea, 2 M thiourea, 25 mM Tris (pH 8.0)] were added and the bead slurry was agitated vigorously for 30 min at room temperature. Beads were centrifuged at 10,000 r.c.f. and the supernatant was transferred to a new tube. This elution procedure was repeated once and the supernatants from both elutions were combined and filtered through glass fiber paper to remove any remaining beads. The eluted proteins were then precipitated using ethanol/acetate and digested in-solution as previously described (Foster et al., 2003). Peptides were acidified and analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) on an LTQ-OrbitrapXL as previously described (Rogers et al., 2008).  Table 2.3 Plasmids used in this study Name pCMVTag-4A (pCMV4A) pCMV4A::CD98-FLAG pcDNA3 pcDNA3::HA2 pcDNA3::HA2espZ pRE112 pCR2.1-TOPO pRE-∆EPespZ pEGFP-N1 pEHespZ-GFPN pEHespZNGFP pEHespZCGFP pGBK-T7 pGBK-EHespZ  Reference/Source Stratagene (Shames et al., 2010) Invitrogen (Shames et al., 2010) (Shames et al., 2010) (Edwards et al., 1998) Invitrogen (Shames et al., 2010) BD Biosciences (Shames et al., 2010) (Shames et al., 2010) (Shames et al., 2010) BD Biosciences (Shames et al., 2010)  43  2.2.8 Lactate dehydrogenase (LDH) release assay For analysis of LDH presence in supernatants, the CytoTox96 Non-radioactive cytotoxicity assay (Promega Corporation, Madison, WI) was used according to manufacturer’s instructions. Briefly, 150 µL of cell supernatants were collected and transferred to a transparent 96 well plate. Plates were spun at 1,500 r.c.f. for 10 min and 50 µL of supernatant was transferred to fresh wells. Fifty µL of LDH substrate was added and the plate was incubated in the dark for 30 min at room temperature followed by addition of 50 µL of stop solution. Absorbance was read at 492 nm in a Tecan plate reader (TECAN). LDH release from bacterial strains alone was quantified and found to be negligible.  2.2.9 Cell lifting assay HeLa or HeLa-EspZ cells (1 x 104) were plated on glass coverslips in 24 well plates. After 24 h, the media were replaced with DMEM 10% FBS, 1% NEAA, 1% GlutaMax supplemented with 10 µCi methyl-[3H] thymidine (Amersham, U.K.). Cells were then grown for 2 days until ~90% confluent, washed 3x in PBS-/-, and infected in serum-free DMEM as described above. At the indicated time points, coverslips were washed 1x in PBS and transferred to 20 mL glass scintillation vials. Five mL of CytoScint scintillation cocktail (MP Laboratories) was added to each vial and counts per minute (CPM) were measured on a Beckman Coulter LS6000 Scintillation Counter (Beckman Coulter).  44  2.2.10 Caspase-3 activity assay HeLa cells transfected with either pcDNA3::HA2 or pcDNA3::HA2espZ were seeded into a black walled clear bottom 96 well plate (Costar) 24 h prior to treatment. Medium was replaced with DMEM and cells were treated with 10 µM staurosporine (STS) and incubated for 3 h at 37˚C/5% CO2. Caspase-3 activity was measured using the Apo-ONE Caspase-3/7 activity assay (Promega Corporation) according to manufacturer’s directions. Cells were incubated with Apo-ONE substrate for 2 h in the dark and relative fluorescence units (RFU) were obtained on a TECAN plate reader (excitation: 485 nm, emission: 535 nm).  2.2.11 siRNA siGENOME oligonucleotides targeting CD98 (SMARTpool SLC3A2) or non-targeting (nt) (SMARTpool Pool 1) (control) were purchased from Dharmacon (Thermo Scientific). Oligonucleotides were suspended in RNase-free water and stored at -80ºC until use. siRNA (100 nM) was transfected into HeLa or HeLa-EspZ cells at 30% confluence using Oligofectamine (Invitrogen) according to manufacturer’s directions and media were changed 24 h post-transfection. At 72 h post-transfection, cells were infected and assayed as described above. Lysates were collected for Western blot analysis to confirm knockdown.  45  2.2.12 Immunoblotting Lysates from HeLa cells were obtained by washing cells in ice cold PBS followed by addition of 75 µL of NP40 lysis buffer. Lysates were scraped into sterile tubes and spun at 16,000 r.c.f. for 10 min at 4°C. Cleared supernatant was transferred to a new tube containing 25 µL 3x Laemmli sample buffer. Lysates were either stored at -20°C for up to two days or run immediately on SDS-PAGE gels followed by transfer to nitrocellulose membranes (BioRad Laboratories)  using a semi-dry transfer apparatus (BioRad  Laboratories). Membranes were blocked in either 5% non-fat milk powder or 2% enhanced chemiluminescence (ECL) advance blocking powder (Amersham) in Tris-buffered saline (TBS) + 0.01% Tween 20 (TBS-T). Primary antibodies were purchased from New England BioLabs unless otherwise indicated and diluted as follows: 1:1000 a-CD98 (Santa Cruz Biotechnology); 1:1000 α-FLAG (Sigma); 1:1000 α-calnexin (Sigma); 1:1000 α-pAKT; 1:1000 α-pFAK; 1:2000 α-AKT; 1:2000 α-FAK; 1:2000 α-HA.11 (Sigma). Membranes were incubated with primary antibody diluted in blocking buffer for 3 h at room temperature or overnight at 4°C with rocking. Membranes were washed 3x 10 min with TBS-T followed by incubation with 1:5000 goat α-rabbit or goat αmouse horseradish peroxidase (HRP). Membranes were washed as above and ECL reagent (Amersham) was added followed by chemiluminescent developing on BioMax film (Kodak).  Anti-pAKT and α-pFAK blots were developed using ECL Advance  Reagent (Amersham) following manufacturer’s instructions. Membrane stripping was  46  performed at room temperature by 5 min wash in distilled (d) H2O, 5 min wash in 0.2 N NaOH, and 5 min wash in dH2O. Densitometric analysis was performed using Adobe Photoshop and ImageJ software on experiments performed at least twice.  2.2.13 Immunoprecipitation HEK 293 lysates were harvested 48 h post-transfection in NP40 lysis buffer and cleared by centrifugation as described above. Twenty µL of lysate was saved for analysis of transfection efficiency and the remainder was immunoprecipitated on either Protein G Sepharose™ 4 Fast Flow (GE Healthcare; for HA pull-down) or DynaL Protein-G conjugated beads (Invitrogen; for GFP pull-down) either with specific antibody, isotype control (mouse IgG whole molecule; Jackson ImmunoResearch Labs), or bead controls. Beads were re-suspended in 3X Laemmli sample buffer and subject to Western Blot analysis.  2.2.14 Confocal and wide field microscopy Coverslips were harvested and prepared by 3% paraformaldehyde fixation followed by 0.2% Triton X-100 permeabilization and blocking with 5% normal goat serum (NGS) and 50 mM NH4Cl in PBS-Tween 20/0.1% BSA (PBST/BSA). Mouse α-FLAG (Sigma) and Alexa 633-conjugated goat α-mouse antibody were used at 1:200 in blocking buffer containing 1% NGS in PBST/BSA.  Coverslips were blocked for 30 min at room  47  temperature, incubated in primary antibody overnight at 4°C and secondary antibody for 1.5 h at room temperature. Three 10 min wash steps with PBST/BSA were performed between antibody steps with shaking. Coverslips were mounted on glass slides using ProLong Gold with DAPI (Invitrogen) overnight. Images were obtained on a Zeiss Meta 510 laser scanning confocal microscope at the University of British Columbia (UBC) Bioimaging Facility.  Images were assembled using ImageJ and Adobe Photoshop  software. Pixel intensity was quantified using ImageJ software. Widefield images were taken on an Olympus CKX41 inverted light microscope mounted with an Infinity 1 camera (Lumenera).  2.2.15 Statistical analyses All statistics were performed with GraphPad software using an unpaired two-tailed t-test with a 95% confidence interval. In all experiments, error bars denote standard deviation (± SD) of samples in duplicates or triplicates.  2.3 EPEC ∆espZ causes enhanced detachment and death in HeLa cells and increased cell death in MDCK cells Since C. rodentium lacking the espZ gene (∆espZ) is severely attenuated for virulence in vivo (Deng et al., 2004), we attempted to identify a role for espZ during EPEC infection using tissue culture models. We infected HeLa cells with EPEC WT, ∆espZ, a T3SSdeficient mutant (∆escN), or mock infected. At 3 h p.i., EPEC ∆espZ caused increased  48  cell rounding and detachment over infections with WT EPEC, ∆escN, and mock-infected HeLa cells (Fig 2.1A). We then quantified the extent of host cell lifting during EPEC ∆espZ infection. HeLa cells were labeled by methyl-[3H]-thymidine incorporation into DNA and acquisition of [3H]-CPM was used to quantify cells remaining adherent at 2, 3, and 4 h p.i.. Increased HeLa cell lifting occurred following infection with EPEC ∆espZ compared to ∆escN or WT (Fig 2.1B). The most pronounced effect was found at 4 h p.i. (*, p < 0.05) (Fig 2.1B). To determine if HeLa cell lifting resulted from cytotoxicity, supernatants were collected from infected HeLa cells and the amount of LDH was quantified. LDH is a cytoplasmic metabolic enzyme that is released from cells lacking plasma membrane integrity and is commonly used to measure cytotoxicity (Kim et al., 2009a). Consistent with published results (Crane et al., 1999), WT EPEC caused significantly more cytotoxicity than ∆escN, at 4 h p.i. (*, p < 0.05) (Fig 2.1C). However, at 3 h and 4 h p.i., significantly more LDH was present in supernatants of cells infected with EPEC ∆espZ compared to WT and ∆escN strains (**, p < 0.001) (Fig 2.1C). To determine if this phenotype could be repeated in a junction-forming cell line, MDCK cells were infected with the aforementioned EPEC strains followed by quantification of cell adhesion and death. Unlike HeLa cells, MDCK cells did not detach from coverslips during EPEC ∆espZ infection over the time course analyzed (Fig 2.1D). However, at 4 h p.i., significantly more LDH was released following infection with EPEC ∆espZ than with WT EPEC (*, p < 0.05) (Fig 2.1E). This suggests that host cell death precedes lifting during EPEC ∆espZ infection in MDCK cells and that EPEC ∆espZ is more cytotoxic to host cells than WT EPEC. In addition, when MDCK cells were infected for 49  8 h with EPEC ∆espZ, we observed extensive damage to the cell monolayer compared to infection with EPEC WT and ∆escN (Fig 2.2). These results imply that EspZ dampens EPEC-mediated host cell cytotoxicity.  Fig 2.1 EPEC ∆espZ infection influences host cell lifting and death (A) Phase contrast images (20X) of HeLa cells infected with EPEC WT, ∆espZ, ∆escN, or mock infected for 3 h. (B) HeLa cell lifting during EPEC infection. HeLa cells were labeled with methyl- [3H] thymidine and infected with EPEC WT, ∆espZ, or ∆escN for 2, 3, or 4 h followed by scintillation counting to obtain radioactive CPM of cells that remained attached p.i.. (C) LDH in supernatants of infected HeLa cells was quantified as described in Experimental Procedures and plotted. (D) MDCK cell lifting during EPEC infection. MDCK cells were labeled and infected as described above and CPM following infection at 2, 3, and 4 h was plotted. (E) LDH in the supernatant of infected MDCK  50  cells was quantified and plotted. Stars denote statistical significance of p < 0.05 (*) and p < 0.001 (**).  Fig 2.2 MDCK cell adherence following 8 h infection MDCK cells were grown to confluence and were either mock infected or infected with EPEC WT, ∆espZ, or ∆escN. Following infection, cells were washed 1 x in PBS -/- and images were acquired by phase contrast microscopy as described in Experimental Procedures.  2.4 HeLa and MDCK cells expressing EspZ are protected from EPEC-mediated killing We attempted to complement the espZ mutation in EPEC by introducing the espZ gene on the pACYC184 vector (pespZ) to create a ∆espZ/pespZ strain. To determine if complementation of the ∆espZ mutation on a plasmid interfered with secretion of other effector proteins, we used EPEC strains lacking the regulator SepD (∆sepD), which hyper-secretes effector proteins (Deng et al., 2005). Removal of espZ from the EPEC genome did not alter the secretion of translocators or effectors; however, expression of espZ by the ∆espZ/pespZ strain disrupted the secretion of other effector proteins from EPEC (Fig 2.3, lane 4). This may be due to disruption of effector hierarchy, which is critical for T3S in A/E pathogens (Thomas et al., 2007, Deng et al., 2005), but the exact reasons remain to be elucidated.  As an alternative approach, espZ was ectopically  51  expressed in HeLa and MDCK cells by stable transfection with pEHespZ-GFPN (HeLaEspZ and MDCK-EspZ, respectively).  Fig 2.3 Coomassie Brilliant Blue staining of secreted proteins from various EPEC strains (A) Secretion profile of WT EPEC and ∆sepD mutants showing effector secretion. EspZ, Tir, and NleA are indicated by black arrows. (B) Translocator secretion profile of various EPEC strains. EspC, EspB/D, and EspA are indicated by black arrows.  We used the EspZ-expressing cell lines to ascertain whether ectopic expression of EspZ rescues host cells from increased lifting and killing by EPEC ∆espZ. Since infection with EPEC ∆espZ caused significantly greater lifting of HeLa cells, we quantified the extent of HeLa-EspZ cell lifting during infection with EPEC WT, ∆espZ, or ∆escN. The fusion to GFP was used to confirm expression of espZ (Fig 2.4A, C, inset). HeLa-EspZ cells exhibited no significant detachment during infection with EPEC WT, ∆espZ, or ∆escN (Fig 2.4A).  52  Death of HeLa-EspZ cells during EPEC infection was measured as previously described (see above). LDH release from HeLa-EspZ cells at 2, 3, and 4 h p.i. did not differ significantly between WT and ∆espZ strains of EPEC (Fig 2.4B). LDH levels, however, were still different between WT and ∆escN (Fig 2.4B). Upon infection of MDCK-EspZ cells, no significant differences between WT and ∆espZ EPEC strains were observed for cell detachment or LDH release (P > 0.05, Fig 2.4C, D). Together, these data strongly suggest that the ∆espZ mutation can be complemented by ectopic expression of EspZ in host cells.  Fig 2.4 Ectopic expression of EspZ protects against cytotoxicity (A) [3H]-labeled HeLa-EspZ cells were infected with EPEC WT, ∆espZ, or ∆escN for 2, 3, or 4 h followed by scintillation counting. EspZ-GFP expression was confirmed by immunofluorescence (inset). (B) LDH present in the supernatant of infected HeLa-EspZ cells was quantified and plotted as described above. (C) MDCK-EspZ cells were [3H]labeled and infected as described above followed by scintillation counting for adherent cells. EspZ expression was confirmed by immunofluorescence (inset). (D) LDH present  53  in the supernatant of infected MDCK-EspZ cells was quantified and plotted as described above.  2.5 EspZ does not antagonize STS-induced apoptotic cell death Since HeLa cells expressing EspZ are protected from EPEC-mediated cell death, its impact on chemically-induced cell death was investigated. STS is a well-characterized chemical inducer of apoptotic cell death and functions by inhibiting cellular kinases and causes caspase-3-dependent apoptosis (Jacobsen et al., 1996). To determine if EspZ antagonizes STS-induced cell death, HeLa cells transfected with pcDNA3::HA2 or pcDNA3::HA2espZ were treated with increasing concentrations of STS for 18 h followed by quantification of LDH present in cellular supernatants. EspZ was unable to protect HeLa cells against STS-induced cell lysis at the concentrations used in this study (Fig 2.5A).  Expression of HA2EspZ was also unable to inhibit Caspase-3/7 activation  following STS treatment (Fig 2.5B). Together, these data suggest that EspZ does not antagonize Caspase-3 dependent apoptosis caused by STS.  54  Fig 2.5 EspZ does not protect HeLa cells from Caspase-3 dependent apoptosis (A) HeLa cells were transfected with either pcDNA3::HA2 (vector) or pcDNA3::HA2espZ and treated with increasing concentrations of STS (0-500 nM) for 18 h. Cell death was quantified by LDH present in cell supernatants and values were set to 1.0 at 0 nM STS. Data are mean ± S.D. of samples in triplicates. (B) HeLa cells were transfected with either pcDNA3::HA2 or pcDNA3::HA2espZ and treated with 10 µM STS for 3 h followed by quantification of caspase-3 activity (relative fluorescence units; RFU) Data are mean ± S.D. of samples in triplicates.  2.6 EspZ interacts with host CD98 To screen for host protein targets of EspZ, SILAC and Y2H experiments were performed. SILAC is a quantitative proteomic technique that has been used for high confidence identification of protein-protein interactions including host protein targets of bacterial effector proteins [(Rogers et al., 2008), and reviewed in (Vermeulen et al., 2008)]. For SILAC screening, pcDNA3::HA2espZ and pcDNA3::HA2 were transfected into light (normal) or heavy (2H4-lysine and  13  C6-arginine) amino acid labeled HEK 293 cells,  respectively. Lysates harvested from heavy and light cell populations were mixed at a 1:1 ratio and immunoprecipitated using α-HA antibodies and tryptic digests of proteins in the immune complex were analyzed by LC-MS/MS (Fig 2.6A). Four unique peptides from CD98 (a.k.a. 4F2 cell-surface antigen heavy chain) all had very low heavy: light  55  ratios (Fig 2.6B, left panel), indicating specific binding to EspZ, while peptides from non-specifically binding proteins such as the 40S ribosomal protein S18 presented with ratios near 1.0 (Fig 2.6B, right panel). This SILAC experiment was performed three times with similar results. Proteins immunoprecipitated in this experiment with heavy: light isotope ratios < 0.33 are listed in Table 2.4. All proteins corresponding to peptides identified in this screen are listed in Appendix A. For Y2H, espZ was cloned into pGBKT7 and used as bait against a pre-transformed library of HeLa cell cDNA using the BD Matchmaker™ System. In the Y2H screen, CD98 was found 29 times by sequencing positive Y2H clones (Table 2.5). Other proteins identified and their putative functions are listed in Table 2.5.  CD98 was the only protein identified in both screens for  interaction with EspZ and therefore was pursued further.  56  Fig 2.6 SILAC screen for EspZ interacting proteins (A) Schematic representation of SILAC methodology. HEK 293 cells expressing either HA2EspZ or HA2 were differentially labeled and lysates were mixed in a 1:1 ratio and proteins immunoprecipitated with α-HA were trypsinized and analyzed by LC-MS/MS. (B) Representative mass spectra for peptides recovered from SILAC experiments. Right panel: Open and filled triangles indicate the expected mass: charge (m/z) of light and heavy forms, respectively, of a peptide (IPDWFLNR) from the non-specifically bound protein 40S ribosomal protein S18 from HA2espZ or HA2, respectively. Left panel: Open and filled triangles indicate the expected m/z of light and heavy forms, respectively, of a peptide (LLTSFLPAQLLR) from the specifically bound CD98 from pull-downs with HA2espZ or HA2, respectively. The light peptide of CD98 is present in a ratio of > 3.0 higher than the heavy peptide, indicating a specific interaction with EspZ.  57  Table 2.4 Summary of proteins identified by MS as specifically immunoprecipitated with HA2EspZ in SILAC experiments Protein Name Averagea S.D.b HA2EspZ 0.17 0.03 Ubiquitin and ribosomal protein 27a precursor 0.18 0.04 4F2 cell-surface antigen heavy chain (CD98) 0.25 0.19 Heat shock protein HSP 90-beta 0.29 0.05 Heat shock protein HSP90-alpha 2 0.29 0.04 a – Averaged SILAC ratio (heavy isotope: light isotope). b – S.D.: standard deviation as determined by quantification of several peptides from either 2 or 3 independent immunoprecipitation experiments.  Table 2.5 Proteins identified in Y2H screen for interaction with EspZ Interactor Number Protein Function Reference(s) of hits CD98 29 Activation of β1-integrin signaling, (Cai et al., 2005, Liu et al., 2003) amino acid transporter localization CD63 3 Tumor associated tetraspanin protein; (Metzelaar et al., associated with early stage melanoma 1991, Demetrick et al., 1992) Pmel17/SILV 2 Silver protein homolog, role in (Du et al., 2003) melanosome biogenesis EMP24/GPL25 2 Cargo receptor in vesicular biogenesis (Muniz et al., from ER 2000) FXYD5 1 Regulates E-cadherin expression; (Lubarski et al., regulates Na+/K+ ATPase 2005, Ino et al., 2002) MLF1-IP 1 Regulates Myeloid Leukemia Factor-1; (Hanissian et al., localizes to nuclei 2004, Suzuki et al., 2007) TIM17b 1 Translocase of inner mitochondrial (Bomer et al., membrane homolog b 1996) CD83 1 Surface antigen of mature dendritic (Zhou et al., 1995) cells CD99-1 1 Splice variant 1 of CD99; involved in (Byun et al., 2006) leukocyte trafficking, adhesion, and apoptosis PCBP1 1 Poly (rC)-binding protein 1; contains (Wang et al., 1999) KH domains involved in RNA binding CEACAM19 1 Carcinoembryonic antigen-related cell (Scorilas et al., adhesion molecule 19; CEAL1 2003)  58  To further probe CD98 as a binding partner of EspZ, we used immunofluorescence to observe the interaction in HeLa cells. HeLa-EspZ cells were transiently transfected with pCMV4A::CD98-FLAG or pCMV4A::FLAG. The FLAG-tag was imaged using mouse α-FLAG antibody followed by Alexa 633-conjugated goat α-mouse antibody (red) and EspZ-GFP imaged in green (Fig 2.7A). Red and green pixel values were quantified for the plasma membrane regions defined by the white line (Fig 2.7A). Pixel intensities in both channels were similar in HeLa-EspZ cells expressing CD98-FLAG (Fig 2.7B) but not FLAG-tag alone (Fig 2.7C), suggesting that EspZ and CD98 localize to membranes similarly in intact cells. Based on previous observations of EspZ at EPEC-induced pedestals (Kanack et al., 2005), we localized endogenous CD98 during EPEC infection. We found that CD98 accumulated under sites of bacterial attachment; however, its recruitment was independent of EspZ and the T3SS (Fig 2.8).  59  Fig 2.7 EspZ and CD98 both localize at the host cell plasma membrane (A) Immunofluorescence localization of EspZ and CD98. HeLa-EspZ cells were transiently transfected with pCMV4A::CD98-FLAG or pCMV4A::FLAG. Alexa fluor 633-conjugated secondary was used to image FLAG (red) by laser scanning confocal microscopy. Scale bar represents 10 µm. (B) Pixel intensities were measured for the red and green channels on the white line on the merged pCMV4A::CD98-FLAG transfected image and (C) on the white line in the merged pCMV4A::FLAG transfected image.  60  Fig 2.8. CD98 localizes at sites of bacterial attachment. HeLa cells were grown on glass coverslips and infected with EPEC WT, ∆espZ, or ∆escN or mock infected for 3 h. Cells were fixed and stained using DAPI to image nuclei (blue), Alexa-Fluor 633-conjugated phalloidin (actin, red), and goat α-CD98 followed by Alexa-Fluor 488-conjugated donkey α-goat antibodies (green). White arrowheads indicate regions enlarged in top right corners of merged image files.  61  Further evidence of the EspZ-CD98 interaction was obtained by co-immunoprecipitation. HEK  293  cells  were  co-transfected  with  pCMV4A::CD98-FLAG  and  pcDNA3::HA2espZ. Pull-down with α-FLAG immunoprecipitated HA2EspZ in cells coexpressing CD98-FLAG, but not in vector transfected cells (Fig 2.9A).  Bead-only  controls were employed to ensure lack of non-specific binding to Sepharose beads (Fig 2.9A). Cell lysates were imaged by Western blot to confirm protein expression (Fig 2.9B).  Fig 2.9. HA2EspZ co-immunoprecipitated with CD98-FLAG. (A) HA2espZ or HA2 were co-expressed with either FLAG or CD98-FLAG in HEK 293 cells as described in Experimental Procedures. Lysates were immunoprecipitated with αFLAG antibody. Western blot analysis for HA-tag and FLAG-tag was performed and CD98-FLAG, but not FLAG alone, could pull down HA2EspZ. (B) HA2EspZ and CD98FLAG input lysates detected by Western blot in lysates subject to coimmunoprecipitation. Proteins expressed in individual samples are indicated by +.  To determine which region of EspZ interacts with CD98, we divided EspZ into Nterminal (amino acids 1-42) and C-terminal (amino acids 43-99) regions (EspZN and EspZC, respectively) and fused them to GFP for imaging (Fig 2.10A). In the presence of CD98-FLAG, α-FLAG antibodies immunoprecipitated EspZGFP and EspZCGFP, but not  62  EspZNGFP or GFP alone (Fig 2.10B), implying that the C-terminal region of EspZ mediates the CD98-EspZ interaction. The amount of full length EspZGFP pulled down was less than that of EspZCGFP, suggesting a potential regulatory role of the N-terminus. Lysates were subject to Western blot analysis to confirm expression of transfected proteins (Fig 2.10C). Together, these data suggest that EspZ interacts with host CD98.  Fig 2.10 The first 42 amino acid residues of EspZ are not required for interaction with CD98 (A) Schematic illustration of full length and truncations of EspZ tagged to GFP. EspZN encodes amino acids 1-42 and EspZC encodes amino acids 43-99, which contain the putative transmembrane regions. White boxes depict putative transmembrane regions. (B) pEHespZ-GFP (ZG), pEHespZC-GFP (ZCG), pEHespZN-GFP (ZNG), and pEHGFPN1 were co-transfected with pCMV4A::CD98-FLAG, and pEH-GFPN1 (G) was cotransfected with pCMV4A::FLAG into HEK 293 cells. Co-immunoprecipitation of lysates with a-FLAG or IgG IC antibody was performed followed by Western blots for GFP and FLAG. (C) Input lysates from cells transfected with GFP-tagged constructs and CD98-FLAG used for α-FLAG co-immunoprecipitation.  63  2.7 CD98 contributes to EspZ-mediated protection from cytotoxicity To determine if CD98 impacts EspZ-mediated protection from EPEC-induced cytotoxicity, we transiently knocked-down CD98 in HeLa cells and HeLa-EspZ cells using siRNA and compared cytotoxicity to a nt siRNA control following EPEC WT, ∆espZ, ∆escN, or mock infection. LDH in supernatants of infected HeLa and HeLa-EspZ cells was quantified as described above and is presented as fold over the mock infected control (mock = 1) to enable comparison between samples in each cell line. There were slight fluctuations in LDH released from mock-infected cells treated with the two siRNAs; however, no significant differences were observed (Fig 2.11). When infected with EPEC ∆espZ, significantly more LDH was released from HeLa and HeLa-EspZ cells treated with CD98 siRNA compared to nt siRNA (Fig 2.12A). However, the difference in LDH released from cells treated with CD98 siRNA compared to nt siRNA was greater in HeLa-EspZ cells (p=0.0003) than HeLa cells (p=0.0176) (Fig 2.12A). Surprisingly, cytotoxicity caused by WT EPEC infection was not influenced by knockdown of CD98 in HeLa or HeLa-EspZ cells (Fig 2.12A) and potential reasons for this are discussed below. CD98 knockdown was confirmed by Western blot analysis with calnexin used as a control for total protein loaded (Fig 2.12B). These data suggest that EspZ is at least partially dependent on CD98 for promotion of host cell survival during infection.  64  Fig 2.11 LDH release of mock infected HeLa and HeLa-EspZ cells treated with nt siRNA or CD98 siRNA LDH release from siRNA treated HeLa and HeLaEspZ cells was quantified by measuring absorbance at 492 nm. Values are mean ± S.D. of samples in triplicates.  Fig 2.12 CD98 contributes to EspZ-mediated protection from EPEC-induced cytotoxicity (A) LDH release from HeLa and HeLa-EspZ cells treated with either CD98-specific or non-targeting (nt) siRNA followed by infection with EPEC WT, ∆espZ, or ∆escN, or mock infected. LDH release was plotted as fold over mock infection ± SD. (B) Reduction in CD98 protein levels was confirmed by Western blot analysis in HeLa (left panel) and HeLa-EspZ (right panel). The blots were stripped and re-probed for calnexin to visualize total protein. P-values indicated are from a two-tailed t-test with 95% confidence interval on samples in triplicates. 65  2.8 EspZ influences AKT and FAK phosphorylation during EPEC infection CD98 modulates signaling downstream of β1-integrin, including the enhancement of integrin survival signals via phosphorylation of AKT and FAK (Feral et al., 2005, Cai et al., 2005). Additionally, WT EPEC dampens FAK phosphorylation in epithelial cells (Shifrin et al., 2002) and AKT phosphorylation in macrophages (Celli et al., 2001). Based on our evidence that EspZ plays a protective role during EPEC infection, partially by interaction with CD98, we tested if EspZ could influence phosphorylation levels of AKT and FAK. HeLa cells were transiently transfected with pcDNA3::HA2 (vector) or pcDNA3::HA2espZ and expression of HA2espZ was confirmed by Western blot analysis (Fig 2.13A). Transfected cells were infected with EPEC WT, ∆espZ, ∆escN, or mock infected and lysates were harvested and immunoblotted for phospho-AKT (Ser473) (pAKT), phospho-FAK (Y576/Y577) (pFAK), and calnexin as a loading control. As reported previously (Shifrin et al., 2002), WT EPEC infection caused a decrease in pFAK levels compared to ∆escN (p = 0.104) (Fig 2.13A, B). In vector-transfected cells, infection with EPEC ∆espZ resulted in a slight decrease in FAK phosphorylation over WT (p = 0.419) (Fig 2.13A, B).  In HA2espZ-transfected cells, pFAK levels were  modestly increased following ∆espZ infection compared to WT infection (p = 0.194) (Fig 2.13A, B). Total FAK levels were slightly decreased following ∆espZ infection of vector control cells. Despite the loss of total FAK, the pFAK/FAK densitometric ratio was still modestly decreased in the absence of EspZ (Fig 2.13B), indicating that the loss of pFAK was not due to decrease in total protein.  66  In vector-transfected cells, pAKT levels were equivalent upon infection with EPEC WT and ∆espZ (Fig 2.13A, C). In HA2espZ-transfected cells, there was a modest increase in pAKT levels following WT infection, and a greater increase following ∆espZ infection when compared to vector-transfected cells (Fig 2.13A, C). The two bands observed are likely different isoforms of AKT.  Total AKT was slightly decreased upon ∆espZ  infection of vector transfected HeLa cells, but was maintained in all other samples (Fig 2.13A). These data suggest that EspZ influences AKT and FAK phosphorylation during EPEC infection.  Fig. 2.13 EspZ enhances phosphorylation of AKT and FAK during EPEC infection (A) HeLa cells transiently expressing either HA2 or HA2espZ were infected with WT, ∆espZ, ∆escN, or mock infected. Lysates were harvested 3.5 h post-infection and run on  67  SDS-PAGE followed by Western blot analysis for either pAKT (S473), pFAK (Y576/577), or HA. The pAKT and pFAK blots were stripped and re-probed for AKT and FAK, respectively, followed by stripping and re-blotting for calnexin to detect total protein. Densitometry was performed to quantify the relative amount of (B) pFAK (represented as pFAK/total FAK) and (C) pAKT (represented as pAKT/total AKT) in lysates.  2.9 Knock-down of CD98 decreases EspZ-mediated phosphorylation of FAK Since EspZ enhanced phosphorylation of FAK and AKT, we aimed to determine whether CD98 played a role. We treated HA2EspZ-expressing HeLa cells with either nt siRNA or CD98-specific siRNA followed by infection with EPEC WT, ∆espZ, ∆escN, or mock infected. Western blot analysis of infected lysates for CD98 and calnexin confirmed siRNA knock-down and total protein loaded, respectively (Fig 2.14A). Western blot analysis of lysates of HeLa cells infected with ∆espZ revealed a decrease in FAK phosphorylation following treatment with CD98 siRNA compared to nt siRNA (Fig 2.14A). Total FAK levels decreased following infection with ∆espZ and treatment with CD98 siRNA. When pFAK/FAK was quantified by densitometric analysis, a significant decrease in FAK phosphorylation was observed when CD98 was knocked-down (**, p < 0.001) (Fig 2.14B). AKT phosphorylation levels were not affected by CD98 siRNA (Fig 2.14C). Together, these data suggest that CD98 is involved in EspZ-mediated induction of FAK phosphorylation during EPEC infection.  68  Fig 2.14 Influence of CD98 knock-down on EspZ-mediated signaling (A) Western blot analysis of HA2espZ-transfected HeLa cells treated with either nontargeting (nt) or CD98 siRNA and infected with EPEC WT, ∆espZ, ∆escN, or mock infected. Lysates were immunoblotted for pAKT, pFAK, CD98, and HA as described above. pAKT, pFAK and CD98 blots were stripped and re-probed for AKT, FAK and calnexin, respectively. (B) Densitometric analysis of phosphorylated FAK (pFAK/FAK) following infection and treatment with either nt or CD98 siRNA. Stars denote statistical significance of p < 0.001 (**). (C) Densitometric analysis of phosphorylated AKT (pAKT/AKT) following infection and treatment with either nt or CD98 siRNA.  69  2.10 Discussion A/E pathogens inject a variety of T3S effectors into host cells during infection and their functions within host cells have begun to be elucidated.  Previously, Deng et al.  demonstrated that the A/E T3S effector EspZ is critical for C. rodentium virulence in a susceptible mouse model (Deng et al., 2004). Subsequently, others have found that EspZ is translocated into HeLa cells and does not impair pedestal formation in EPEC (Kanack et al., 2005). Due to the severity of the espZ mutation (∆espZ) in impairing virulence, we attempted to define a molecular mechanism for EspZ function in cultured cells. We initially observed enhanced cell lifting and death in HeLa cells infected with EPEC ∆espZ. EPEC causes both necrotic and apoptotic cell death (Abul-Milh et al., 2001, Barnett Foster et al., 2000) and thus we quantified LDH in cell supernatants, which measures depletion of plasma membrane integrity and would include all death caused by EPEC (Kim et al., 2009a). We found that EPEC ∆espZ caused increased release of LDH from infected HeLa cells, suggesting that EspZ may be protective to host cells. This was confirmed upon espZ expression in HeLa cells and resultant reduction in cytotoxicity during EPEC ∆espZ and WT infection. Although ectopic expression results in a greater quantity of EspZ in cells, our observations that ∆espZ infection of EspZ-expressing cells results in similar phenotype to WT infection suggests that the overexpression is likely not producing artifacts. The absence of cell lifting and delay in cell death following EPEC infection of MDCK cells provides interesting implications for EPEC infection of the polarized epithelia of the gut. In the first study to examine EPEC infection of MDCK cells, Canil and colleagues did not observe perturbation of the epithelial monolayer until 10 h p.i. (Canil et al., 1993). This demonstrates that polarized epithelial monolayers can 70  withstand EPEC infection longer than HeLa cells and suggests that EPEC can maintain attachment and colonization in the gut for prolonged periods prior to sloughing of infected cells. We believe that EspZ plays a role in prolonging host cell survival in the polarized epithelia of the gut during EPEC infection. Bacterial T3S effectors interact with proteins in the host cell to influence host cell signaling pathways and we identified CD98 as a host protein target of EspZ. CD98 is a type-II transmembrane glycoprotein that regulates amino acid transport in epithelial cells and interacts with β1-integrin to facilitate enhanced cell survival, cell adhesion, cell spreading, and cellular transformation (Rintoul et al., 2002, Henderson et al., 2004, Feral et al., 2005). Several other proteins were identified to interact with EspZ; however, CD98 was the only protein identified by both Y2H and SILAC screens. To date, no other bacterial effector protein has been shown to interact with CD98. Immunofluorescent analysis revealed that CD98 and EspZ localize to peripheral membranes in HeLa cells.  Kanack and colleagues previously observed EspZ  accumulation at some sites of EPEC-induced pedestals at late time points post infection (Kanack et al., 2005). To determine if CD98 localizes to EPEC-induced pedestals, we stained HeLa cells with α-CD98 and phalloidin to observe pedestals. Surprisingly, we found CD98 clustered at sites of bacterial attachment following infection with WT, ∆espZ, and ∆escN strains, suggesting that bacterial attachment is sufficient for CD98 recruitment. Importantly, following EPEC WT and ∆espZ infection, CD98 was at the sites of pedestal formation, but did not overlay with EPEC-induced actin tails. In the study by Kanack and colleagues, EspZ was not observed at all pedestals and was only found under large microcolonies of EPEC at 5 h p.i., with no EspZ observed at 3 h p.i. 71  (Kanack et al., 2005). This suggests that EspZ accumulates under pedestals at late time points p.i.; however, molecular details of T3S EspZ localization at early time points are not yet clear. We found that CD98 contributes to EspZ-mediated protection from cytotoxicity influenced by EPEC ∆espZ. Interestingly, cytotoxicity of EPEC WT did not increase following treatment with CD98-specific siRNA. There are several possibilities for this observation: (1) EspZ translocated into the HeLa cells via the T3SS is able to utilize residual CD98 present in cells treated with CD98 siRNA to delay cell death; (2) there may be pleiotropic effects between EspZ and other effector proteins that only occur via injection with the T3SS; or (3) EspZ also interacts with other host proteins to influence cytotoxicity (see Chapter 3). Previously, Kenny and Warawa observed that EPEC Tir behaves differently when transfected into host cells when compared to Tir secreted via the T3SS (Kenny et al., 2001) and this may also be the case for EspZ. Despite this, our data does suggest that CD98 contributes to EspZ-mediated protection from cytotoxicity, and other mechanisms by which EspZ functions during EPEC infection are being investigated. CD98 facilitates phosphorylation of AKT and FAK via interaction with β1-integrin (Feral et al., 2005) and clustering of CD98 propagates this effect (Rintoul et al., 2002). AKT is a well known pro-survival mediator, which is activated by the concerted action of PI3K and 3-phosphoinositide-dependent protein kinase [reviewed in (Kim et al., 2002)]. We demonstrate that ectopically expressed EspZ influences AKT phosphorylation during infection with EPEC ∆espZ, which appears to be independent of CD98. In HeLa cells, EPEC WT and ∆espZ dampen AKT phosphorylation, which has previously been reported 72  for WT infection of macrophages (Celli et al., 2001). The influence of EspZ on AKT phosphorylation remains inconclusive and more work is required to tease out the impact of EPEC effectors on AKT signaling and host cell survival. FAK is a tyrosine kinase that performs a plethora of functions in eukaryotic cells. FAK is regulated by CD98-mediated signaling events (Rintoul et al., 2002) and activates host cell survival signaling pathways (Zouq et al., 2009, Huang et al., 2007). Thus, the observation that EspZ influenced FAK signaling supports the idea that EspZ acts through CD98 to enhance pro-survival signaling during EPEC infection. Decrease in total FAK levels during infection with EPEC ∆espZ could be due to activation of caspase enzymes, since FAK is a known target of these proteases (Grossmann et al., 2001); however, we found that EspZ had only a subtle effect on STS-induced cell death, indicating that inhibition of apoptotic cell death may be only a minor role played by EspZ during infection. Based on this study and previous reports, we propose a model by which EspZ interacts with CD98 in host cell membranes to promote host cell survival. Activation of β1integrin and resultant activation of FAK may occur via the EspZ-CD98 interaction (Fig 2.15), since antibody-mediated cross-linking of CD98 has produced this effect (Rintoul et al., 2002). Although EspZ influences AKT phosphorylation, this was not affected by CD98 knock-down and it is possible that EspZ uses other mechanisms to modulate AKT signaling. During EPEC infection, survival signaling from FAK likely antagonizes cytotoxic signals from other EPEC effectors to prevent premature host cell death (Fig 2.15) (Crane et al.,  73  2001, Nagai et al., 2005). The observation that cytotoxicity is enhanced in the presence of tyrosine kinase inhibitors (Crane et al., 1999) supports our hypothesis that EspZ acts through FAK to delay host cell death. Premature death of cells in vivo may explain the severe attenuation in virulence observed following C. rodentium ∆espZ infection (Deng et al., 2004) and this possibility is currently being investigated. Delaying cell death via effector proteins such as EspZ would therefore provide the pathogen with valuable time to colonize efficiently prior to dissemination. The regulation of host cell cytotoxicity as a pathogenicity mechanism by EPEC has been previously suggested (Crane et al., 1999) and is further supported by this study. Ultimately, understanding the role and interplay between T3S effector proteins will provide valuable knowledge by which to combat A/E pathogen infection.  74  Fig 2.15 Schematic model for EspZ function during EPEC infection EspZ binds CD98 in its terminal 43-99 amino acids. EspZ (red) binding to CD98 (green) may enhance signaling from β1-integrin (blue), leading to activation of the FAK survival signals and possibly activation of PI3K/AKT. Once activated, FAK initiates survival and adhesive signaling via PI3K-dependent (Bouchard et al., 2007) and -independent (Huang et al., 2007) mechanisms. EspZ influences AKT signaling, but likely through mechanisms independent of CD98. Signaling from EspZ likely antagonizes pro-death signals from other EPEC effectors. Yellow appendages protruding from CD98 represent its heavy glycosylation.  75  Chapter 3: EspZ localizes to host mitochondria and interacts with host translocase of inner mitochondrial membrane (TIM) 17B  3.1 Introduction Host cell death during A/E pathogen infection is intricately regulated by several T3S effector proteins. The effectors EspF, Map, and Cif cause apoptosis in host cells (see Sections 1.2.1, and 1.2.2). However, A/E pathogens also inject effectors that enhance host cell survival such as NleH1, NleH2, and NleD, which down-regulate pro-apoptotic signaling (see Section 1.2.2). We have also shown that host cell death is strongly antagonized by the effector EspZ during EPEC infection [see Chapter 2; (Shames et al., 2010)]. In this study, we further investigated mechanisms of cell death antagonized by EspZ and other host protein targets. In our previous study, we identified TIM17B as a putative EspZ-interacting protein by Y2H (Shames et al., 2010). TIM17 is a component of the TIM23 complex, which is responsible for transport of proteins into the inner mitochondrial membrane and the mitochondrial matrix (Frazier et al., 2003, Chacinska et al., 2009). This complex is composed of three inner membrane proteins (TIM50, TIM23, and TIM17) and is dependent on energy derived from the mitochondrial matrix motor (PAM), an intact Δψm, and ATP (Chacinska et al., 2009). Specifically, TIM17 is responsible for recruiting the PAM and sorting pre-proteins in the inner mitochondrial membrane (IMM) (Chacinska et al., 2005).  76  In this study, we identified a second mechanism by which EspZ is able to antagonize host cell death since the EspZ-CD98 interaction does not fully account for the pro-survival effects of EspZ (Shames et al., 2010). To further characterize the mechanism of host cell survival promoted by EspZ, we investigated whether EspZ functions at mitochondria, since this organelle plays a major role in host cell death and survival pathways. We determined that a significant decrease in ∆ψm accompanies cytotoxicity mediated by EPEC ∆espZ infection.  We further demonstrated that EspZ localizes to host  mitochondria and confirmed its interaction with host TIM17B. The ability of EspZ to protect cells against death during EPEC infection was dampened following siRNA knock-down of TIM17B. We have thus uncovered another mechanism by which EspZ protects host cells against rapid cell death during EPEC infection.  3.2 Experimental procedures 3.2.1 Tissue culture, bacterial strains, primers, transfection and infection conditions HeLa and HEK 293T cells were purchased from the ATCC and maintained in DMEM High Glucose (Thermo Scientific) supplemented with 10% FBS (Thermo Scientific), 1% NEAA (Gibco), and 1% GlutaMAX (Gibco). Cells were used from passages 5-20. Bacterial strains used in this study are listed in Table 3.1. Oligonucleotide primers used in this study are listed in Table 3.2 and plasmids used in this study are listed in Table 3.3. HeLa cells were transfected using FuGENE HD (Roche) as previously described (Shames et al., 2010) and were assayed 24-36 h post-transfection.  Lysates were  subjected to Western blot analysis to determine efficiency of transfection. HEK 293T 77  cells were transfected using calcium phosphate as described previously (Shames et al., 2010). Infections were performed as previously described [see Chapter 2; (Shames et al., 2010)]. For the JC-1 assay, a 1:600 dilution of overnight standing bacterial cultures were prepared in DMEM and used to infect HeLa cells in a 96-well plate.  3.2.2 Generation of espZ and espZHA complemented strains Complementation of espZ was achieved by chromosomal insertion with Tn7 into EPEC ∆espZ or ∆espZ∆sepD as previously described (Shames et al., 2011a). Briefly, espZ was PCR amplified from EPEC genomic DNA using oligonucleotides espZ-f and espZ-r. An HA-tagged version of espZ was also created by PCR using oligonucleotides espZ-f and espZHA-r. These PCR products were sequenced at NAPS Unit (UBC, Vancouver, British Columbia) and cloned into the BamHI/EcoRI site of pMAC5 prior to conjugation and site specific integration into the chromosome of EPEC ∆espZ or ∆espZ∆sepD. The complementing  strains  were  designated  MCE007  (∆espZ/espZ),  MCE008  (∆espZ/espZHA), MCE010 (∆espZ∆sepD/espZ), and MCE011 (∆espZ∆sepD/espZHA).  3.2.3 Generation of TIM17B-FLAG fusion TIM17B cDNA was obtained from clones from the BD Matchmaker™ pre-transformed HeLa cell cDNA library as previously described (Shames et al., 2010). The TIM17B cDNA lacking its stop codon was cloned into pCMV-TAG4A (pCMV4A). Primers  78  TIM17-F and TIM17-R were used to PCR amplify TIM17B cDNA with a Kozak sequence on the 5’ end.  The purified PCR product was cloned as a NotI/HindIII  fragment into the multiple cloning site (MCS) of pCMV4A in frame with the downstream FLAG-tag sequence. TIM17B gene sequence was confirmed by DNA sequencing. Table 3.1 Bacterial strains used in this study Strain EPEC O127:H6 Strain E2346/89 WT ∆espZ ∆espZ/espZ (MCE007) ∆espZ/espZHA (MCE008) ∆sepD ∆sepD∆espZ ∆sepD∆espZ/espZ (MCE010) ∆sepD∆espZ/espZHA (MCE011) Escherichia coli SM10λpir DH5αλpir EC100Dpir+ ω7249  Source or Reference (Levine et al., 1978) (Shames et al., 2010) This study This study (Deng et al., 2005) (Thomas et al., 2007) This study This study (Miller et al., 1988) (Miller et al., 1988) Epicentre Biotechnologies (Babic et al., 2008)  Table 3.2 Oligonucleotide primers used in this study Name Sequence (5'3')a,b TIM17-F aattgcggccgcggcaccatggaggagtacgctcgggagccc TIM17-R aataagcttgtggtactgatagctgggg espZ-f cgacggatccttagttatactctaaagcaaacgtaac espZ-r tagaattcttaggcatatttcatcgctaatc espZHA-r tagaattcttaagcgtaatctggaacatcgtatgggtaggcatatttcatcgctaatccg a. Restriction endonuclease cleavage sites are in bold. b. Kozak sequences are underlined.  Table 3.3 Plasmids used in this study Name pCMV-TAG4A pCMV4A::TIM17B pcDNA3::HA2EspZ pcDNA3::HA2 pEHespZ-GFPN  Source or Reference Stratagene This study (Shames et al., 2010) (Shames et al., 2010) (Shames et al., 2010) 79  Name pEGFP-N1 pMAC5  Source or Reference BD Biosciences Clontech (Sham et al., 2011)  3.2.4 JC-1 ∆ψm assay HeLa cells were plated at 5 x 104 cells/well in a 96-well black clear bottom tissue culturetreated plate (Costar) 24 h prior to infection. Cells were washed in PBS+/+ and a 1:600 dilution of EPEC strains or un-inoculated LB broth in DMEM was added to each well. Cells were infected for 2, 3, or 4 h, as indicated. The JC-1 Mitochondrial Membrane Potential Assay Kit (Cayman Chemical Company) was used according to manufacturer’s instructions. Briefly, 15 min prior to terminating the infection, 10 µL of prepared JC-1 reagent was added to each well and mixed by gently swirling plate. Plate was centrifuged for 5 min at 400 r.c.f. in a Beckman table-top centrifuge and supernatant was aspirated. Two hundred microlitres of assay buffer was added to each well and plate was centrifuged as above. Wash steps were repeated followed by addition of 100 µL assay buffer and analyses using a Tecan M200 microplate reader (TECAN). J-aggregates and J-monomers were quantified using excitation/emission wavelengths of 560/595 nm and 485/535 nm, respectively. Data were plotted as J-aggregates/J-monomers of samples in triplicates.  3.2.5 LDH release assay For analysis of LDH presence in supernatants, CytoTox96 Non-radioactive cytotoxicity assay (Promega Corporation, Madison, WI) was used according to manufacturer’s  80  instructions and as previously described (see Chapter 2).  Percent cytotoxicity was  measured by normalizing 492 nm values to a 100% cytotoxicity control and uninfected controls according to manufacturer’s instructions. LDH release from bacterial strains alone was quantified and found to be negligible.  3.2.6 Confocal microscopy HeLa cells were plated on sterile glass coverslips 24 h. prior to transfection. Cells were washed in PBS+/+ and fixed in 3.7% paraformaldehyde (Sigma) for 20 min at room temperature. Coverslips were washed 3X 10 min in PBS-/- and permeabilized in 0.2% Triton-X 100 for 5 min at room temperature.  Coverslips were inverted on 50 µL  blocking buffer [5% NGS in PBST/BSA, 50 mM NH4Cl] for 30 min humidified at room temperature. Coverslips were incubated in (1:200 dilution) mouse α-HA.11 (Sigma), rabbit α-COX IV (Cell Signaling Technologies), mouse α-cytochrome c (BD Biosciences)  or  rabbit  α-TIM17B  (ProteinTech)  primary  antibodies  in  1%  NGS/PBST/BSA overnight at 4˚C. Coverslips were washed 3 X 10 min in PBST/BSA followed by inverting on secondary antibody solution (Alexa 488-conjugated goat αmouse or Alexa 633-conjugated goat α-rabbit antibodies + 1% NGS in PBST/BSA) in the dark, humidified, at room temperature for 90 min. Coverslips were washed in the dark 3 X 10 min in PBST/BSA and mounted on clean glass slides in ProLong Gold with DAPI (Invitrogen) mounting medium.  Coverslips were sealed with nail polish and  imaged on an Olympus Fluoview 10i laser scanning confocal microscope at the UBC  81  Bioimaging Facility (Vancouver, British Columbia).  Images were assembled using  ImageJ and Adobe Photoshop software.  3.2.7 Immunoprecipitation HEK 293T cell lysates were generated using NP40 lysis buffer and cleared by centrifugation as described previously (Shames et al., 2010).  Protein G-conjugated  DynaL beads (Invitrogen) were bound to mouse α-FLAG M2 (Sigma) or normal mouse IgG isotype control (Jackson ImmunoResearch Labs).  Immunoprecipitations were  performed according to manufacturer’s instructions. Twenty microlitres of 3X Laemmli sample buffer was used to resuspend beads prior to Western blot analysis.  3.2.8 Immunoblotting Proteins separated by denaturing SDS-PAGE were transferred to Pure Nitrocellulose or methanol-activated polyvinylidene fluoride (PVDF) membranes (BioRad) using a wet transfer cell (BioRad). Membranes were blocked for 30 min in blocking buffer [5% nonfat milk in TBST] with rocking at room temperature. Primary antibodies were diluted in blocking buffer as described below and membranes were incubated in this solution overnight at 4˚C with rocking. Wash steps were performed in TBST for 3 X 10 min followed by incubation in a 1:5000 of goat α-mouse or goat α-rabbit HRP-conjugated antibodies for 60 min at room temperature with rocking. Membranes were washed as described followed by addition of ECL reagent (Amersham) and chemiluminescent  82  imaging on Kodak BioMax film. Primary antibodies were diluted as follows: 1:1000 αHA.11 (Sigma); 1:1000 α-TIM17B (ProteinTech); 1:1000 α-GFP (Abcam); 1:2500 αCalnexin (Enzo Pharmaceuticals); 1:1000 α-β tubulin (Sigma); 1:1000 α-COX IV (Cell Signaling Technologies); and 1:1000 α-FLAG M2 (Sigma).  3.2.9 Isolation of cellular mitochondria Mitochondria-enriched fractions were isolated from HeLa cells either infected with EPEC strains or transfected with pcDNA3::HA2espZ or pcDNA3::HA2 using the Mitochondria Isolation Kit for Cultured Cells (Pierce; Thermo Scientific). Briefly, 2 x 107 cells were pelleted and resuspended in Mitochondrial Isolation Reagent (MIR) A and vortexed at medium speed for 5 seconds followed by incubation on ice for 2 min. MIR B was added and tubes were incubated on ice for 5 min with 5 second vortex mixing every minute. MIR C was added and tube was inverted to mix followed by centrifugation at 700 r.c.f. for 10 min at 4˚C. Supernatants were transferred to a new tube and spun at 12,000 r.c.f. for 15 min at 4˚C. Supernatants (cytosolic fraction) were transferred to a new tube and stored at -20˚C until use. Pellet (mitochondrial-enriched fraction) was washed in MIR C and centrifuged at 12,000 r.c.f. for 5 min. Mitochondria were solubilized in 100 µL 2% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate  (CHAPS)  in  TBS.  Cytosolic and mitochondrial protein concentrations were quantified using the Coomassie Plus Protein Assay (Pierce; Thermo Scientific) and resuspended in 3X Laemmli sample buffer prior to protein separation by SDS-PAGE and Western blot analysis.  83  3.2.10 siRNA siGENOME oligonucleotides targeting TIM17B (SMARTpool TIMM17B) or nt (SMARTpool Pool 1) (control) were purchased from Dharmacon (Thermo Scientific). Oligonucleotides were suspended in 1X siRNA buffer (Dharmacon) and stored at -80ºC until use. siRNA (100 nM) was transfected into HeLa cells at 30% confluence using Oligofectamine (Invitrogen) according to manufacturer’s directions and media were changed 24 h post-transfection.  At 48 h post-transfection, cells were infected and  assayed as described above. Lysates were collected for Western blot analysis to confirm knockdown.  3.2.11 T3S assay Proteins secreted by the EPEC T3SS were assayed as previously described (Deng et al., 2005). Briefly, bacteria were subcultured from overnight shaking cultures into prewarmed DMEM and incubated in a 37˚C/5% CO2 incubator for 6 h. Bacterial pellets were saved and supernatants were filtered followed by precipitation with trichloroacetic acid (TCA) overnight at 4˚C. Secreted proteins were pelleted by centrifugation and resuspended in 3X Laemmli sample buffer. Saturated Tris solution was added to adjust the pH. Proteins (secreted and pellet) were separated by SDS-PAGE followed by either staining with Coomassie Brilliant Blue G250 or Western blot analysis.  84  3.2.12 Statistical analysis All statistics were performed using an unpaired two-tailed t-test with a 95% confidence interval. In all experiments, error bars denote S.D. of samples in duplicates or triplicates.  3.3 Chromosomal insertion of espZ can complement the ∆espZ knock-out and preserve type-III secretion of other effector proteins EspZ is an essential virulence factor in attaching and effacing bacterial pathogens and we have previously determined that it acts to protect host cells from rapid cell death using an espZ knockout strain (∆espZ) (Deng et al., 2004, Shames et al., 2010). In our previous study (see Chapter 2), we used cell lines stably expressing EspZ-GFP to complement the ∆espZ mutation since over-expression of espZ on a plasmid interfered with secretion of other effector proteins [Chapter 2; (Shames et al., 2010)]. For this study, we generated EPEC ∆espZ strains containing a single chromosomal copy of espZ, either epitope tagged with HA or left un-tagged, outside the LEE. We then utilized an EPEC ∆espZ∆sepD double-knockout to visualize secretion of effector proteins, as performed previously [see Chapter 2;(Shames et al., 2010, Deng et al., 2005)]. Type-III secretion assays were then used to visualize proteins secreted by our various EPEC strains. We found that the EPEC ∆espZ/espZ and ∆espZ/espZHA strains exhibited a normal translocator secretion profile since EspB/D, and EspA could all be visualized (Fig 3.1A). The ∆espZ∆sepD strains all had normal effector secretion profiles indicated by the presence of Tir and NleA (Fig 3.1A) (Deng et al., 2005). EspZ could only be visualized in the supernatants of EPEC ∆sepD; however, the chromosomally inserted espZHA was visualized by Western blot,  85  demonstrating that EspZ expressed by the chromosomal copies of espZ in the ∆espZ background is indeed secreted but likely at a lower level (Fig 3.1B). We then utilized the newly constructed ∆espZ/espZ strain as a tool to further understand the mechanism(s) by which EspZ protects infected host cells from rapid cell death.  Fig 3.1 Chromosomal insertion of espZ preserves type-III secretion of effectors and translocators from EPEC (A) Secreted proteins from EPEC strains were separated by SDS-PAGE and visualized by Coomassie G250 staining. Translocator (black text) and effector (red text) secretion was analyzed using ∆espZ, ∆espZ/espZ and ∆espZ/espZHA in a WT or ∆sepD background, respectively. EPEC ∆escN was used as a control for no type-III secretion, since only EspC, a non-type III secreted protein, is observed. (B) Secreted proteins were subjected to Western blot analysis with α-HA antibodies. A ~11 kDa HA-reactive band was observed in the secreted protein fraction of the ∆sepD∆espZ/espZHA strain.  86  3.4 EspZ protects EPEC-infected cells from severe loss of inner mitochondrial membrane potential (Δψ m) We previously demonstrated that EspZ protects EPEC-infected cells from premature cell death by interaction with CD98 (Shames et al., 2010).  However, the EspZ-CD98  interaction does not account for all of the protection elicited by EspZ, suggesting that other mechanisms exist by which EspZ is protecting host cells [Chapter 2; (Shames et al., 2010)]. Since cell death caused by EPEC ∆espZ was characterized by rapid cell lysis, indicative of necrotic cell death, we examined ∆ψm in HeLa cells infected with EPEC strains. To observe changes in ∆ψm over time during the course of EPEC infection, we utilized  the  J-aggregate-forming  lipophilic  cation  5,5',6,6'-tetrachloro-1,1',3,3'-  tetraethylbenzimidazolcarbocyanine iodide (JC-1) (Smiley et al., 1991). JC-1 localizes preferentially to mitochondria and is used to measure ∆ψm (Smiley et al., 1991, Cossarizza et al., 1993). In healthy cells with high ∆ψm, JC-1 forms aggregates (Jaggregates), which emit a red fluorescence; however, in unhealthy cells when ∆ψm is low, JC-1 is present in a monomeric form (J-monomers), which emits a green fluorescence. In this assay, ∆ψm can be quantified by plotting the ratio of J-aggregates: J-monomers. HeLa cells were infected with either EPEC WT, ∆espZ, ∆espZ/espZ, or left uninfected for 2, 3, or 4 hours followed by staining with JC-1. Fluorescence intensity was measured at excitation/emission of 560/595 nm and 485/535 nm for monomers and aggregates, respectively. Aggregate/monomer ratios were normalized to 1.0 in uninfected cells (Fig 3.2A). Infection with WT EPEC resulted in lower ∆ψm compared to uninfected cells (Fig 3.2A), as previously demonstrated (Ma et al., 2006). However, EPEC ∆espZ caused a significant decrease in the aggregate/monomer ratio compared to WT and ∆espZ/espZ  87  infection at 3 and 4 h p.i. (*, p < 0.05), indicating that ∆ψm is greatly decreased (Fig 3.2A). Supernatants were taken from infected cells and LDH was quantified. Untreated cells were lysed using Triton-X 100 and LDH present in the supernatant of these cells was set at 100% cytotoxicity. As observed previously, significantly greater LDH levels were present in the supernatants of HeLa cells infected with EPEC ∆espZ when compared to WT at 3 h and 4 h p.i. (**p < 0.01) (Fig 3.2B). Infection with EPEC ∆espZ/espZ resulted in similar LDH release to that caused by WT EPEC infection at 3 h p.i. but significantly greater LDH release was observed following ∆espZ/espZ infection at 4 h p.i. (*p < 0.05) (Fig 3.2B). These data demonstrate that EPEC ∆espZ infection causes a severe loss in ∆ψm that occurs concurrently with host cell lysis.  Fig 3.2 Infection with EPEC ∆espZ causes severe loss of ∆ψm (A) HeLa cells were infected with either EPEC WT, ∆espZ, ∆espZ/espZ, or left UI. JC-1 reagent was added to cells 15 minutes prior to terminating infection and cells were washed with Assay Buffer three times. Fluorescence intensity was measured at excitation emission wavelengths of 560/595 nm and 485/535 nm for J-monomers (low ∆ψm) and J-aggregates (high ∆ψm), respectively. ∆ψm was plotted as the ratio of Jaggregates/ J-monomers. Asterisks indicate statistical significance (p < 0.05) of samples in triplicates by Student’s t-test. (B) Supernatants of HeLa cells infected for 2, 3, or 4 hours with either EPEC WT, ∆espZ, ∆espZ/espZ, left UI, or treated with Triton-X 100 (100% cytotoxicity; not shown) were assayed for the presence of LDH. Values were normalized against Triton-X 100-treated cells (100% cytotoxicity) and UI cells (1% cytotoxicity). Asterisks (**) indicate statistical significance (p < 0.05) of samples in triplicates by Student’s t-test. These data are representative of three independent experiments.  88  3.5 EspZ interacts with TIM17B In our previous study, we identified TIM17B as a putative host protein binding partner to EspZ by Y2H (Shames et al., 2010). TIM17B is a mitochondrial inner membrane protein that is responsible for voltage gating of the TIM23 protein-import complex and relies on an intact ∆ψm to function (Bomer et al., 1996, Martinez-Caballero et al., 2007). We thus aimed to confirm our previous Y2H data that EspZ interacts with host TIM17B. Coimmunoprecipitation experiments were performed to determine if an EspZ-GFP fusion protein could co-immunoprecipitate with TIM17B-FLAG.  We used an α-FLAG  antibody to immunoprecipitate proteins interacting with TIM17B-FLAG and normal mouse IgG as an IC. We were able to specifically immunoprecipitate EspZ-eGFP but not eGFP alone with TIM17B-FLAG (Fig 3.3A). Input lysates were analyzed by Western blot to confirm expression of TIM17B-FLAG, EspZ-eGFP, and eGFP (Fig 3.3B). As suggested by our previous Y2H data (Shames et al., 2010), EspZ does indeed interact with TIM17B.  89  Fig 3.3 TIM17B interacts with EspZ (A) HEK 293T cells were co-transfected with pCMV4A::TIM-17, pCMV-Tag4A, pEHespZ-GFP, or pEGFP as indicated. Cells were lysed and subjected to immunoprecipitation with α-FLAG antibodies or mouse IgG IC as described. Precipitated proteins were subjected to Western blot analysis. EspZ-GFP and TIM17FLAG are indicated with arrows and non-specific antibody bands are indicated with asterisks (*). (B) Lysates from HEK 293T cells were subjected to Western blot analysis to confirm transfection efficiency. EspZ-GFP, GFP, and TIM17-FLAG are indicated with arrows. 3.6 EspZ localizes to mitochondria We have previously demonstrated that EspZ localizes to host cell membranes (Shames et al., 2010) and based on the interaction between EspZ and TIM17B, we hypothesized that EspZ likely also localizes to host mitochondria. To test this hypothesis, we transfected HeLa cells with pcDNA3::HA2espZ and stained these cells for cytochrome c oxidase (COX) IV, which is a mitochondrial inner membrane protein (Kurushima et al., 2010).  90  We used Alexa 633-conjugated goat α-rabbit antibodies to image COX IV (red) and mouse α-HA.11 followed by Alexa 488-conjugated goat α-mouse antibodies to image EspZ (green). We observed similar staining patterns in the α-COX IV and α-HA images suggesting that EspZ localized similarly to COX IV (Fig 3.4A). To further demonstrate that EspZ localizes to mitochondria in addition to other cellular membranes, we isolated mitochondria from HeLa cells expressing HA2EspZ and performed Western blot analysis on the cytosolic and mitochondrial fractions. COX IV and β-tubulin were used as markers for mitochondrial and cytosolic fractions, respectively (Fig 3.4B).  We observed HA2EspZ in mitochondria-enriched but not  cytosolic fractions (Fig 3.4B), which provides further evidence that EspZ can localize to host cell mitochondria in addition to other cellular membranes.  91  Fig 3.4 EspZ localizes to host cell mitochondria (A) HeLa cells were plated on glass coverslips and transfected with pcDNA3::HA2espZ. Cells were stained with rabbit α-COX IV (red) and mouse α-HA.11 (green) followed by Alexa 633-conjugated goat α-rabbit and Alexa 488-conjugated goat α-mouse antibodies, respectively. Coverslips were mounted in ProLong Gold with DAPI to image cellular nuclei (blue). (B) HeLa cells were transfected with either pcDNA3::HA2espZ or pcDNA3::HA2 and mitochondrial fractions were isolated as described. Antibodies that recognize cytochrome c, and β tubulin were used as controls for mitochondrial (Mito) and cytoplasmic (Cyto) fractions, respectively. Anti-HA antibodies were used to visualize HA2EspZ.  92  3.7 EspZ mitochondrial localization is independent of TIM17B TIM17B is a component of the TIM23 protein import complex that functions to transport proteins into the inner mitochondrial membrane or mitochondrial matrix (Bomer et al., 1996). We therefore hypothesized that TIM17B may be responsible for mitochondrial localization of EspZ. To test this hypothesis, we utilized siRNA specific to TIM17B and imaged cells expressing EspZ-GFP or GFP alone using confocal microscopy. Cells were transfected with either TIM17B-specific siRNA (TIM17B) or nt (control) siRNA for 48 h followed by transfection with pEHespZ-GFP or pEGFP. Transfected cells were treated with rabbit α-TIM17B and mouse α-cytochrome c antibodies, to image mitochondria. Alexa 405-conjugated goat α-rabbit antibodies were used to image TIM17B (blue) and Alexa 633-conjugated goat α-mouse antibodies were used to image cytochrome c (red). Cytochrome c staining was similar in the presence or absence of TIM17B (Fig 3.5, arrowheads). In addition, EspZ-cytochrome c co-staining was unaltered in the absence of TIM17B (Fig 3.5; arrowheads). Based on these observations, TIM17B does not play a major role in localizing EspZ to host cell mitochondria.  93  Fig 3.5 The EspZ-TIM17B interaction is not required for mitochondrial localization of EspZ HeLa cells grown on glass coverslips were transfected with either non-targeting (control) siRNA or TIM17B-specific siRNA followed by transfection with pEGFP::espZ or pEGFP alone. Cells were stained with rabbit α-TIM17B and mouse α-cytochrome c antibodies followed by Alexa 633-conjugated goat α-mouse and Alexa 405-conjugated goat α-rabbit secondary antibodies. TIM17B (Blue) staining was used to determine knock-down efficiency and cytochrome c (Red) staining was used to visualize mitochondria. Knock-down was confirmed by decreased TIM17B staining in cells treated with TIM17B-specific siRNA. “n” indicates presence of cell nuclei as seen in phase contrast images.  3.8 siRNA knock-down of TIM17B dampens EspZ-mediated survival during EPEC infection EspZ protects EPEC-infected host cells from premature cell death and is essential for successful colonization of C. rodentium in vivo (Shames et al., 2010, Deng et al., 2004).  94  To determine if TIM17B contributes to EspZ-mediated protection from cell death during EPEC infection, we performed LDH release assays using HeLa cells treated with TIM17B-specific (TIM17B) or nt (control) siRNAs.  HeLa cells transfected with  TIM17B and control siRNA were infected with EPEC WT or EPEC ∆espZ for 2, 3, or 4 h. TIM17B knock-down resulted in significantly greater HeLa cell cytotoxicity during WT EPEC infection compared to the non-targeting siRNA control at all time points examined (*p < 0.05; Fig 3.6A). However, knock-down of TIM17B did not influence cytotoxicity levels during infection with EPEC ∆espZ (Fig 3.6B). Uninfected and detergent treated cells were used as controls for spontaneous and total lysis, respectively. Lysates from infected cells treated with TIM17B and control siRNA were collected and subjected to Western blot analysis to confirm TIM17B knock-down. Indeed, TIM17Bspecific siRNA resulted in a reduction of TIM17B protein (Fig 3.6C). Calnexin was used as a loading control (Fig 3.6C). Together, these data suggest that in the presence of EspZ, TIM17B contributes to host cell survival during EPEC infection.  95  Fig 3.6 siRNA knock-down of TIM17B dampens the ability of EspZ to protect HeLa cells from EPEC-mediated cell lysis HeLa cells treated with either TIM17B-specific or non-targeting (control) siRNA were infected with EPEC WT (A) or ∆espZ (B) for 2, 3, or 4 h. LDH present in cellular supernatants was quantified and plotted as percent cytotoxicity. Asterisks (*) indicate statistical significance of samples in triplicates (p < 0.05). Data are representative of two independent experiments. (C) Lysates were generated from EPEC-infected and uninfected cells and subjected to Western blot analysis using α-TIM17B and α-Calnexin (loading control) antibodies to demonstrate siRNA knock-down efficiency.  96  3.9 Discussion Diarrheagenic Escherichia coli rely on a plethora of virulence factors secreted by a T3SS to efficiently hijack host cell signaling pathways. Specifically, several effector proteins have been shown to intricately regulate cytotoxicity during EPEC infection by promoting antagonistic pro-survival and pro-death pathways. We had previously demonstrated that the T3S effector EspZ protects EPEC-infected cells from premature cytotoxicity partially due to interaction with host CD98 (Shames et al., 2010). Since CD98 accounted for only partial protection against cell death elicited by EspZ, we aimed to further investigate the mechanisms of EspZ-mediated cell survival. Host cell death resulting from EPEC ∆espZ infection had characteristics of necrotic cell death. Cell lysis, as characterized by the presence of LDH in cell supernatants, occurs rapidly following necrotic cell death and was displayed by EPEC ∆espZ-infected cells. Rapid cell lysis resulting from necrosis generally occurs after mitochondrial swelling, ATP depletion, and decreased ∆ψm (Golstein et al., 2007). We initially examined ∆ψm as a measure of necrotic cell death during EPEC infection and found that ∆ψm decreased significantly during EPEC ∆espZ infection concurrently with LDH release. Infection with a chromosomally complemented ∆espZ/espZ strain resulted in similar ∆ψm to WT infection, which suggests that in the absence of EspZ, EPEC causes necrotic cell death. We then aimed to further investigate secondary mechanisms by which EspZ could be protecting infected host cells from rapid cell death. In our previous study, we identified several putative host binding partners to EspZ via SILAC and Y2H screening. TIM17B was found to interact with EspZ by Y2H but not  97  SILAC (Shames et al., 2010).  Since TIM17B contributes to voltage gating in  mitochondria, and requires intact ∆ψm to function, we tested whether EspZ does indeed interact with TIM17B. We found initially that EspZ bearing an N-terminal epitope tag does not interact with TIM17B (data not shown), which would explain why TIM17B did not immunoprecipitate with HA2EspZ in our SILAC experiments. However, EspZ with a C-terminal GFP epitope tag does interact with TIM17B by co-immunoprecipitation. We were thus able to validate the interaction between TIM17B and EspZ. TIM17B is an IMM protein and we therefore hypothesized that EspZ likely localizes to host mitochondria. To test this hypothesis, we transiently expressed HA2EspZ in HeLa cells. We observed that HA2EspZ localized similarly to COXIV and attempted to further confirm EspZ localization using organellar fractionation. We observed EspZ localization in mitochondrial enriched fractions of HeLa cells expressing HA2EspZ, which further suggests that ectopically expressed EspZ localizes to mitochondria. Interestingly, EspZ does not encode a canonical MTS.  However, the effector EspJ also localizes to  mitochondria in the absence of a known MTS (Kurushima et al., 2010). We subsequently aimed to determine if EspZ requires TIM17B to gain access to host mitochondria. Interestingly, siRNA knock-down of TIM17B did not impact localization of EspZ to host mitochondria, which implies that EspZ either utilizes another host protein for import or encodes a non-canonical MTS, similar to EspJ (Kurushima et al., 2010). Interaction with TIM17B likely occurs following EspZ localization to mitochondria and may regulate mitochondrial localization of Map and EspF by interfering with their import. Further investigation is required to determine if the EspZ-TIM17B interaction is required for import of other effectors into host mitochondria. 98  To determine if the EspZ-TIM17B interaction contributes to EspZ-mediated protection against host cell death, we utilized siRNA knock-down of TIM17B and LDH release assay. We observed increased cytotoxicity during infection with EPEC WT but not EPEC ∆espZ following siRNA knock-down of TIM17B compared to nt siRNA controls, suggesting that TIM17B plays a role in protection against premature cytotoxicity in the presence of EspZ. Since siRNA knock-down of CD98 did not influence cytotoxicity during WT EPEC infection in our previous study, EspZ is likely using complementary mechanisms to achieve this function. TIM17B has not been implicated in cell death; however, its function is strongly correlated with ∆ψm (Martinez-Caballero et al., 2007). Further studies are required to reveal the precise mechanisms by which EspZ is utilizing TIM17B to protect host cells against rapid cell death. It has become increasingly apparent that T3S effector proteins carry out several functions during A/E pathogen infection, as observed for the effector EspF (Holmes et al., 2010). By further investigating roles of effector proteins, we are uncovering mechanisms by which these pathogens are able to efficiently cause disease and incapacitate their hosts. Regulation of host cell death is a critical function of several A/E pathogen effector proteins and understanding how they manipulate cell death pathways will aid in understanding their mechanisms of infection.  99  Chapter 4. The non-LEE encoded effector NleC dampens host NF-κB signaling and IL-8 secretion during EPEC infection 4.1 Introduction NleC is one of the most conserved Nle effector proteins with >95% similarity in NleC proteins from EPEC, EHEC, and C. rodentium (Deng et al., 2004).  NleC was  characterized as a substrate of the T3SS by Marchés and colleagues in EPEC and EHEC (Marches et al., 2005). The function of NleC remained elusive until publication of four recent studies implicating NleC in suppression of NF-κB activity during EPEC infection in cell culture (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011). Prior to publication of these studies, the function of NleC was investigated here. All known A/E pathogen effector proteins that influence NF-κB signaling are encoded outside the LEE and include NleB, NleC, NleE, NleH1, and NleH2.  NleB and NleE  function to stabilize IκBα, thereby retaining NF-κB dimers in the host cell cytosol (Newton et al., 2010, Nadler et al., 2010). NleH1 interacts with RPS3 and decreases its nuclear abundance whereas NleH2 reduces NF-κB signaling by an as yet undetermined mechanism (Gao et al., 2009, Royan et al., 2010) (see Section 1.4.2). In this study, the ability of the effector NleC to impair NF-κB signaling during EPEC infection was examined. We found that NF-κB p65 was present in higher abundance in the nuclear fractions of Caco-2 cells following infection with an nleC deficient EPEC strain (∆nleC). In addition, infection of Caco-2 cells with EPEC ∆nleC resulted in increased NF-κB activity and IL-8 secretion.  100  4.2 Experimental procedures 4.2.1 Tissue culture, bacterial strains and infection conditions Caco-2 cells were purchased from the ATCC and maintained in DMEM High Glucose (HyClone) supplemented with 10% FBS (HyClone), 1% NEAA (Gibco), and 1% GlutaMAX (Gibco). Cells were used from passage 5-25. Bacterial strains used in this study are listed in Table 4.1. Oligonucleotides used in this study are listed in Table 4.2. Plasmids used in this study are listed in Table 4.3. Caco-2 cells were infected with an m.o.i of ~100:1 in serum-free DMEM for the times indicated. Bacteria were cultured for infection as described (Shames et al., 2011a).  4.2.2 Generation of EPEC ∆nleC An nleC in-frame deletion mutant was generated in the streptomycin-resistant derivative (Smr) of EPEC O127:H6 strain E2348/69 (Table 4.1) using the sacB gene-based allelic exchange method and the suicide vector pRE112 (Edwards et al., 1998). PCR was used to generate two fragments (1.0 and 2.18 kb, respectively) using primer pairs EPnleCD-1 and EPnleC-DR as well as EPnleC-DF and EPnleCD-2 (Table 4.2). The PCR products were cloned into pCR2.1-TOPO (Table 4.3) and verified by DNA sequencing. After digestion with KpnI/NheI and NheI/SacI, respectively, the two fragments were gelpurified and cloned into pRE112 digested with KpnI/SacI in a 3-way ligation. The resulting plasmid pRE-∆EPnleC contained the nleC gene with an internal in-frame deletion from codons # 27 to #332 (about 90% of the coding region). An NheI site was  101  introduced into the deletion site. Plasmid pRE-∆EPnleC was transformed into E. coli SM10λpir by electroporation, and introduced into EPEC strain E2348/69 Smr by conjugation. After sucrose selection, EPEC colonies resistant to sucrose and sensitive to chloramphenicol were screened for deletion of nleC by PCR using primers EPnleB-DF and EPnleD-DR (Table 4.2). T3S of effectors and translocators by the EPEC ∆nleC mutant was confirmed as previously described (Deng et al., 2005).  4.2.3 Generation of ∆nleC/nleC and ∆nleC/nleCHA complemented strains Complementation of nleC was achieved by chromosomal insertion with Tn7 into EPEC ∆nleC. A chloramphenicol marked Tn7 delivery vector was created by subcloning a SacI frt-cat-frt, Klenow end-filled, fragment from pFCM1 into the EcoRV site of pUC18R6KT-mini-Tn7T (Choi et al., 2005), yielding pMAC5, and transformed into DH5αλpir. The orientation of the cat cassette was confirmed with EcoRI/NcoI restriction digests. The nleC complementation construct was made by fusing the nleBCD operon promoter to nleC that contained 19 nt upstream of the predicted start site. A 348 bp region containing the nleBCD promoter was PCR amplified with oligonucleotides nleBCD-f and nleBCDp-r (Table 4.2). A 1033 bp PCR product that contains the nleC gene, with 19 bp upstream of the ATG was amplified with oligonucleotides nleC19-f and nleC-r (Table 4.2). The promoter and nleC gene were fused by PCR SOEing, using 1 ng of each PCR product as template in a single PCR reaction as previously described (Croxen et al., 2006), using oligonucleotides nleBCD-f and nleC-r (Table 4.2). An HAtagged nleC was also amplified using the same templates and PCR SOEing protocol, except using oligonucleotide nleCHA-r (Table 4.2) instead of nleC-r. Both PCR products  102  were sequenced (NAPS Unit, UBC) and subcloned as an EcoRI/BamHI fragment into a similarly cut pMAC5 yielding pMAC5/19nleC and pMAC5/19nleCHA, and transformed into EC100Dpir+. The complementing constructs were mobilized into EPEC ∆nleC by mating with ω7249 harboring the Tn7 helper plasmid, pTNS2, and either pMAC5/19nleC or  pMAC5/19nleCHA.  Transconjugants  were  selected  on  LB  containing  chloramphenicol, and the proper Tn7 insertion was checked as previously described (Choi et al., 2005). The resulting complementing strains were designated MCE003 (nleC) and MCE004 (nleC-HA). Table 4.1 Bacterial strains used in this study Name SM10λpir DH5αλpir EC100Dpir+ ω7249 EPEC E2348/69 WT EPEC E2348/69 ∆nleC EPEC E2348/69 ∆nleC/nleC EPEC E2348/69 ∆nleC/nleCHA EPEC E2348/69 ∆escN  Source/Reference (Miller et al., 1988) (Miller et al., 1988) Epicentre Biotechnologies (Babic et al., 2008) (Levine et al., 1978) (Sham et al., 2011) (Sham et al., 2011) (Sham et al., 2011) (Gauthier et al., 2003)  Table 4.2 Oligonucleotides used in this study Name Sequence (5´  3´)a EPnleCD-1 gggtaccgtttgaacctaatcctgaacg EPnleC-DR cgctagcacgattaggagcaatgggagc EPnleC-DF cgctagcgtggacaaacacaatcagcga EPnleCD-2 ggagctccatcaacagcacgttcagtgg EPnleB-DF ggctagccagtatacatgcagttcatgg EPnleD-DR ggctagcctgatgtaataccaagttgag nleBCD-f tcagaattcccaagctatatgttaactgc nleBCDp-r gtttatccatattttcttcacaac nleC19-f gttgtgaagaaaatatggataaaccagggtattagatataaacatg nleC-r cgacggatcctcatcgctgattgtgtttgtc nleCHA-r cgacggatcctaagcgtaatctggaacatcgtatgggattcgctgattgtgtttgtccac a – Restriction endonuclease cleavage sites are underlined  103  4.2.4 NF-κB activity assay Caco-2 cells were seeded into 24 well plates at 2.5 x 105 cells/well in 1 mL of medium and incubated overnight in a 37˚C 5% CO2 incubator. One hour prior to transfection, cells were washed with sterile PBS-/- and 0.5 mL of pre-warmed low serum media (0.5% FBS, 1% NEAA, 1% GlutaMax) was added to each well. Per well, 25 ng pNF-κB Luc vector (Clontech Labs) was mixed with 75 ng phRL-TK (Renilla) vector (Promega Corporation), and topped up to 500 ng of total DNA with pCMVTag-4A vector. DNA was diluted in 50 µL DMEM and 3 µL of GenJet for Caco-2 cells (SignaGen Laboratories) was diluted in 50 µL DMEM in a separate tube. Diluted GenJet was added directly to diluted DNA and mixed.  Transfection mixture was incubated at room  temperature for 15 min prior to addition of 100 µL to each well of Caco-2 cells. Twentyfour hours post-transfection, media were aspirated from Caco-2 cells and replaced with 0.5 mL pre-warmed serum-free DMEM. Cells were infected with 1 µL of overnight standing EPEC cultures as indicated. Infections were performed in triplicates for each strain used. For UI cells, 1 µL of un-inoculated LB media was added to Caco-2 cells. At 3 h p.i., media were aspirated and replaced with fresh DMEM supplemented with 100 µg/mL gentamicin and 5 ng of recombinant human IL-1β per well for 3 h. The DualLuciferase Reporter Assay (Promega Corporation) was used according to manufacturer’s instructions. Briefly, infected Caco-2 cells were washed with 500 µL of sterile PBS-/- and 100 µL of passive lysis buffer was added followed by 15 min incubation at room temperature with rocking. Twenty microlitres of cell lysate was used for luciferase quantification. Data were plotted as Firefly/Renilla to enumerate NF-κB activity relative  104  to a constitutive CMV promoter and NF-κB activity resulting from all infections was plotted relative to that of UI cells, which was set at 100% activity.  Table 4.3 Plasmids used in this study Name pCR2.1-TOPO pMAC5 pTNS2 pCMVTAG-4A pNF-κB phRL-TK  Source/Reference Invitrogen (Shames et al., 2011a) (Choi et al., 2005) Stratagene Clontech Promega Corporation  4.2.5 IL-8 enzyme-linked immunosorbent assay (ELISA) Prior to infection of Caco2 cells, media were aspirated and cells were washed in PBS-/-. Cells were infected in serum-free DMEM per well for 3 h at 37˚C/5% CO2. Media were aspirated and cells were incubated in medium containing 100 µg/mL gentamicin (Sigma) and 5 ng/mL IL-1β (RnD Systems). Total secreted IL-8 was quantified as described using the BD OptEIA™ Human IL-8 ELISA kit (BD Biosciences) according to manufacturer’s directions and as described (Shames et al., 2011a).  4.2.6 Nuclear isolation and western blot analysis Isolation of host cell nuclei was performed using the NE-PER Nuclear Isolation Kit (Pierce Scientific) according to manufacturer’s instructions and as described (Shames et al., 2011a). Nuclear and cytosolic proteins were quantified using the Coomassie Plus Assay Reagent (Pierce Scientific). Equal amounts of protein (15 µg) were separated by SDS-PAGE followed by Western blot analysis as previously described (Shames et al., 105  2011a). Primary antibodies were diluted as follows; 1:500 rabbit α-p300 (Santa Cruz Biotechnology); 1:1000 rabbit α-p65 (Cell Signaling Technology); 1:2500 rabbit αcalnexin (Enzo Life Sciences); 1:1000 rat α-HA (Roche). HRP-conjugated goat α-rabbit (Sigma) and goat α-rat (Sigma) antibodies were used a 1:5000.  4.2.7 Statistical analyses Statistics were performed using an unpaired two-tailed t-test with a 95% confidence interval. In all experiments, error bars denote S.D. of samples in duplicates or triplicates. GraphPad software was used to determine P-values.  4.3 NleC reduces IL-8 secretion during EPEC infection EPEC actively dampens IL-8 secretion during infection and several effectors have recently been shown to influence this phenotype [see Sections 1.3.1 and 1.4.2] (Royan et al., 2010, Gao et al., 2009, Newton et al., 2010, Nadler et al., 2010, Ruchaud-Sparagano et al., 2007). A panel of EPEC strains mutated for effector genes revealed that the nleC gene is required for full suppression of host cell IL-8 secretion by EPEC (Sham et al., 2011). To demonstrate that NleC contributes to suppression of IL-8 secretion, EPEC ∆nleC was chromosomally complemented with either nleC or tagged nleCHA, resulting in ∆nleC/nleC and ∆nleC/nleCHA EPEC strains. Caco-2 cells were infected for 3 h with EPEC WT, ∆nleC, ∆nleC/nleC, ∆nleC/nleCHA, ∆escN, or left uninfected followed by 3 h incubation with gentamicin to inhibit further infection and IL-1β to stimulate IL-8 gene expression. Supernatants were collected and secreted IL-8 was quantified by sandwich 106  ELISA. Fold IL-8 of samples in triplicates was plotted for all infections relative to UI cells (set at 1.0) (Fig 4.1).  Infection with EPEC ∆nleC and ∆escN resulted in  significantly greater IL-8 secretion compared to WT EPEC (*p < 0.05; **p < 0.01) (Fig 4.3). EPEC ∆nleC/nleC and ∆nleC/nleCHA infection resulted in significantly less IL-8 secretion compared to ∆nleC infection, thus demonstrating that NleC contributes to dampening IL-8 secretion from EPEC infected cells (Fig 4.1).  Fig 4.1 NleC contributes to repression of IL-8 secretion during EPEC infection Caco2 cells were infected with EPEC WT, ∆nleC, ∆nleC/nleC, ∆nleC/nleCHA, ∆escN or left UI. At 3 h p.i., cells were treated with 100 µg/mL gentamicin and IL-1β and incubated for 3 h. Supernatants were collected and used in an IL-8 sandwich ELISA. Values presented are mean ± S.D. fold IL-8 secretion compared to UI cells following IL1β treatment. Asterisks (*p < 0.05; **p < 0.001) denote statistical significance comparetd to WT infection of samples in triplicates. 4.4 EPEC ∆nleC infection results in nuclear p65 accumulation Expression of IL-8 is dependent on activation and nuclear translocation of host NF-κB dimers. To determine if NleC influences translocation of NF-κB to host cell nuclei, we infected Caco-2 cells for 3 h with EPEC WT, ∆nleC, ∆nleC/nleC, ∆nleC/nleCHA, and  107  ∆escN followed by isolation of host cell nuclear and cytosolic fractions. Host p300 and calnexin were used as controls for nuclear isolation and equal amounts of protein were loaded into each well for Western blot analysis. Infection with EPEC ∆nleC and ∆escN resulted in increased p65 protein levels in the nuclear fractions of Caco-2 cells in comparison to WT infection (Fig 4.2). Following Caco-2 infection with the ∆nleC/nleC and ∆nleC/nleCHA strains, we were able to observe that NleC localizes to host cell nuclear and cytosolic fractions and that NleC contributes to inhibition of p65 nuclear tranlocation (Fig 4.2).  Fig 4.2 p65 translocates to nuclear fractions in EPEC ∆nleC-infected Caco2 cells Caco2 cells were infected with either EPEC WT, ∆nleC, ∆nleC/nleC, ∆nleC/nleCHA, ∆escN, or UI. At 3 h p.i., cells were lysed and nuclei were extracted and subjected to Western blot analysis using α-Calnexin, α-p65, α-p300, and α-HA antibodies. Equal amounts of protein were loaded in each lane with Calnexin and p300 used as controls for cytosolic and nuclear proteins, respectively.  108  4.5 NleC contributes to repression of NF-κB signaling To further determine whether NleC could inhibit NF-κB-mediated gene activation, quantification of gene expression was performed using a Dual-Luciferase reporter assay. Caco-2 cells were transfected with reporter plasmids with Firefly and Renilla luciferase genes under the control of the κB-enhancer and CMV promoter, respectively, followed by infection with EPEC WT, ∆nleC, ∆nleC/nleC, ∆nleC/nleCHA, ∆escN, or UI and treatment with or without IL-1β.  We confirmed that the ∆nleC strain was unable to  repress NF-κB activity to WT levels following IL-1β stimulation and that the ∆nleC/nleC and ∆nleC/nleCHA strains could complement the nleC knock-out by repressing NF-κB activity to the same level as WT EPEC (Fig 4.3). In comparison to infection with EPEC WT, ∆nleC/nleC, and ∆nleC/nleCHA, infection with EPEC ∆nleC and ∆escN resulted in significantly greater NF-κB activity (*, p < 0.05 and **, p < 0.001, respectively) (Fig 4.3). Thus, our complemented ∆nleC/nleC and ∆nleC/nleCHA strains repress NF-κB activity similarly to WT EPEC.  109  Fig 4.3 Infection of Caco-2 cells with EPEC ∆nleC/nleC results in similar NF-κB activity levels to WT EPEC Caco-2 cells transfected with Dual-Luciferase reporter plasmids were infected with EPEC WT, ∆nleC, ∆nleC/nleC, ∆nleC/nleCHA, ∆escN, or left uninfected. At 3 h post-infection, cells were incubated with 100 µg/mL gentamicin +/- IL-1β. NF-κB activity was obtained by reading luminescence following addition of substrates specific for Firefly (κBspecific) and Renilla (constitutive) Luciferase proteins. Data are expressed as mean ± standard deviation of relative luminescence units (RLU) of Firefly/Renilla Luciferase activity of samples in triplicates and are representative of three independent experiments. Asterisks (*p < 0.05; **p < 0.001) indicate statistical significance compared to NF-κB activity following WT EPEC infection.  4.6 Discussion Regulation of innate immune signaling is an important virulence strategy of A/E pathogens. The list of T3S effector proteins contributing to this phenotype continues to grow and they have been shown to utilize complementary mechanisms by which to dampen pro-inflammatory signaling from infected IECs. Here, we demonstrate that the effector NleC can dampen NF-κB activity and secretion of the pro-inflammatory cytokine IL-8. During the course of our work, four other groups demonstrated that NleC 110  could dampen NF-κB activity by proteolytic degradation of p65, which contributed to suppression of IL-8 secretion (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011). We were able to confirm previous reports that EPEC infection dampens IL-8 secretion and that NleC contributes to this phenotype (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011, Ruchaud-Sparagano et al., 2007). Subsequently, nuclear isolation of EPEC infected cells was used to determine the localization of p65 during EPEC infection. Following infection of Caco2 cells, we found that p65 localizes to nuclear fractions in the absence of NleC, suggesting that NF-κB dimers that facilitate IL-8 gene expression are present in cellular nuclei. We also demonstrated that T3S NleC localizes to host cell nuclei which had previously been observed only for ectopically expressed NleC (Baruch et al., 2011).  We chose Caco2 cells since Baruch and  colleagues could only observe p65 degradation in this cell line following infection with an EPEC strain overexpressing nleC on a plasmid (Baruch et al., 2011). NleC also inhibits the p38 MAPK (Sham et al., 2011), which can enhance NF-κB activity by phosphorylation of p65 (Vanden Berghe et al., 1998). We subsequently used quantitative techniques to confirm repression of NF-κB and IL-8 secretion during infection with EPEC strains encoding nleC. Several other bacterial pathogens also utilize effector proteins to modulate NF-κB activity to subvert pro-inflammatory signaling [for review, see (Rahman et al., 2011)]. Yersinia pseudotuberculosis, Y. pestis, and Y. enterocolitica are the causative agents of gastroenteritis (Y. pseudotuberculosis and Y. enterocolitica) and bubonic plague (Y. pestis) and dampen pro-inflammatory signaling in host macrophages. 111  Yersinia spp.  utilize the effector YopP/J to impair NF-κB activity by post-translational modification of the IKKβ and inhibition of IκBα degradation. Mukherjee and colleagues found that YopJ is an acetyl-transferase that transfers acetyl groups to IKKβ (Mukherjee et al., 2006). This specific acetylation inhibits phosphorylation and subsequent activation of IKKβ (Mukherjee et al., 2006). YopJ also functions as a de-ubiquitinase that prevents degradation of IκBα and consequently NF-κB translocation to the host cell nuclei (Zhou et al., 2005). The effector SseL from S. Typhimurium is also a de-ubiquitintase that prevents degradation of IκBα and subsequent activation of NF-κB (Le Negrate et al., 2008). Yersinia spp. and Salmonella spp. infect host macrophages and have evolved effector proteins that function to maintain NF-κB dimers in the host cell cytosol. Inhibition of NF-κB activity is thus a common theme in bacterial pathogenesis. Further investigation into the function of A/E effector proteins may reveal more levels of redundancy regarding modulation of NF-κB, and resultant pro-inflammatory, signaling cascades.  112  Chapter 5. NleC interacts with and degrades the host acetyltransferase p300  5.1 Introduction EPEC utilizes several complementary mechanisms to dampen innate immune signaling during infection.  As a strategy to dampen NF-κB activity during EPEC infection,  degradation of p65 is mediated by a zinc metalloprotease domain (HEIIH) encoded within nleC (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011). Degradation of p65 occurred in the nucleus and cytosol of cells either ectopically expressing nleC or infected with EPEC strains encoding nleC (Yen et al., 2010, Baruch et al., 2011). NleC was also shown to act synergistically with NleE and NleB to dampen IL-8 release from host cells (Pearson et al., 2011, Baruch et al., 2011, Yen et al., 2010). No other targets of the NleC protease were identified in these studies. To gain further insight into the function of NleC in host cells, we aimed to identify other host protein targets. Using SILAC, we identified the acetyltransferase p300 (see Section 1.3.1) as a target of NleC and subsequently determined that the TAZ1 domain of p300 is required for this interaction. We also determined that nuclear p300 levels are decreased in the presence of NleC and that the recently characterized metalloprotease activity of NleC is responsible for p300 degradation. Furthermore, overexpression of p300 in Caco2 cells dampens EPEC-mediated repression of IL-8 secretion during infection and siRNA knock-down of p300 results in decreased IL-8 secretion by EPEC ∆nleC-infected cells.  113  We therefore provide the first example of a bacterial virulence factor targeting host p300, and a complementary mechanism used by EPEC to influence IL-8 levels.  5.2 Experimental procedures 5.2.1 Tissue culture, bacterial strains, transfection and infection condition HeLa (ATCC), Caco-2 (ATCC), HEK 293 (ATCC), and HEK 293T cells (ATCC) were maintained in DMEM High Glucose (Thermo Scientific) supplemented with 10% FBS (Thermo Scientific), 1% NEAA (Gibco), and 1% GlutaMax (Gibco). Cells were used from passages 5-20. Bacterial strains used in this study are listed in Table 5.1, oligonucleotide primers used in this study are listed in Table 5.2, and plasmids used in this study are listed in Table 5.3. HeLa cells were transfected using FuGENE HD (Roche) as previously described (Shames et al., 2010). Cells were infected or assayed 24-36 h post-transfection and lysates were subjected to Western blot analysis to confirm transfection efficiency. Caco2 cells were transiently transfected using GenJet for Caco-2 transfection reagent (SignaGen Laboratories) according to manufacturer’s instructions. Media were changed 24 h post-transfection and cells were either infected or assayed at 24 h or 48 h posttransfection as specified.  Western blot analysis was used to confirm transfection  efficiency. GM6001 (Millipore) was used at a concentration of 20 µM and added to cells 1 hour post-transfection. Volume equivalent of the vehicle control dimethylsulfoxide (DMSO) was added simultaneously to separate wells.  114  Infection of HeLa and Caco-2 cells was performed as previously described (Shames et al., 2010).  5.2.2 p300 truncations The plasmid pCMVβ::p300HA was used for generation of all truncations used in this study. All primers used for PCR amplification are listed in Table 5.2. A NotI/HindIII digestion of pCMV-Tag4A was used to clone in all truncations. p300NM was amplified using primers N-F and M-R; p300 N was amplified using N-F and N-R; p300 C was amplified using C-F and C-R; p300 NuR-TAZ was amplified using N-F and TAZ-R; p300 NuR-KIX was amplified using N-F/NurKIX-R and NuRKIX-F/KIX-R followed by digestion with KpnI. All constructs were digested with NotI/HindIII and ligated into NotI/HindIII digested pCMV-Tag4A.  The NuR-KIX construct was generated as a  NotI/KpnI NuR product and a KpnI/HindIII KIX product and both digested products were used for ligation into the NotI/HindIII digested pCMV-Tag4A vector.  p300  truncation sequences were confirmed by DNA sequencing at the NAPS Unit (UBC, Vancouver, British Columbia).  5.2.3 GFP and HA2 epitope tag fusions to EHEC nleC C-terminal GFP fusions to the nleC gene of EHEC O157:H7 strain EDL933 were constructed using the vector pEGFP-N1. The coding region of EHEC nleC was amplified by PCR using primers EHnleC-NF and EHnleC-NR and cloned into pCR2.1-TOPO and  115  verified by DNA sequencing, and then sub-cloned as an XhoI/BamHI fragment into pEGFP-N1 to generate a fusion to the N-terminus of GFP (the resulting plasmid designated pEHnleC-GFPN). pcDNA3::HA2nleC was generated as described previously using primers CdH65 and CdH66 (Shames et al., 2010).  5.2.4 Generation of NleCE184A-GFPN and NleCE184A-FLAG pEHnleC-GFPN and pCMV-Tag4A::nleC were used as templates for QuikChange mutagenesis (Stratagene) using primers NleCE184A-F and NleCE184A-R according to manufacturer’s instructions. Resultant plasmids were designated pEHnleCE184A-GFP and pCMV4A::nleCE184A. Mutation of Glu 184 was confirmed by DNA sequencing using primers pEHnleC-NF and pEHnleC-NR or commercially available T7-F and T3-R primers (NAPS Unit, UBC).  Table 5.1 Bacterial strains used in this study Strain EPEC O127:H6 E2348/69 WT E2348/69 ∆escN E2348/69 ∆nleC E2348/69 ∆nleC/nleC (MCE003) EHEC O157:H7 EDL 933 WT  Reference (Levine et al., 1978) (Gauthier et al., 2003) (Shames et al., 2011a) (Shames et al., 2011a) (Johnson et al., 1984)  Table 5.2 Oligonucleotides used in this study Name Sequence (5'3')a CdH65 ggatgaaaattccctcattacagtcc CdH66 catgcggccgctcattgctgattgtgtttgtcc  116  Name Sequence (5'3')a SRS5 attctcgagatgaaaattccctcattacagtcc EHnleC-NF cctcgagatgaaaattccctcattacagtcc EHnleC-NR cggatcctgctgattgtgtttgtccacatcc NleCE184A-F gcaggaaggactgattcacgcgattattcatcatgttactg NleCE184A-R cagtaacatgatgaataatcgcgtgaatcagtccttcctgc N-F aattgcggccgcggcaccatggccgagaatgtggtgg N-R attaagcttattcatggaaactggaacc M-R attaagcttggaagggtcatccccc TAZ1-F atttgcggccgcggcaccatgccagccccgcaggtccagc TAZ1-R attaagctttgtcggcatctgatttac C-F aattgcggccgcggcaccatgcagcctcaaactacaataaataaag C-R aataagctttccgagggaggcgtagtc NuRKIX-F attggtacccaaccccaggtgcaagc NurKIX-R attggtaccctgttgacccatgttgggc a. Restriction enzyme cleavage sites are in bold. Table 5.3 Plasmids used in this study Name pCMVTAG-4A (pCMV4A) pCMVβ::p300HA pEGFP-N1 pCR2.1-TOPO pcDNA3::HA2 pcDNA3::HA2nleC pCMV4A::nleC pEHnleC-GFPN pEHnleCE184A-GFP pCMV4A::nleCE184A  Source/Reference Stratagene Addgene #10718 BD Biosciences Invitrogen (Shames et al., 2010) (Shames et al., 2011a) (Shames et al., 2011a) (Shames et al., 2011a) (Shames et al., 2011a) (Shames et al., 2011a)  5.2.5 Recombinant protein purification EHEC nleC was cloned into pGEX-6P-3 (GE Lifesciences) with an upstream triple HA tag (HA3) (lab vector; PGEX-6P-3::HA3) and transformed into E. coli BL21 (DE3). An overnight shaking culture in LB supplemented with 100 µg/mL ampicillin (LB Amp 100) was used to make a 1% subculture in 750 mL LB Amp 100. Cells were grown to log phase (~3 h) and then induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) for 4 h at 37˚C with shaking. Pellet was harvested by centrifugation at 5,000 r.c.f., 117  washed with PBS-/-, and stored at -20˚C overnight.  Cells were thawed on ice and  resuspended in 10 mL lysis buffer (50 mM Tris pH 8.0, 100 mM NaCl, 1 mM EDTA) supplemented with fresh lysozyme (200 µg/mL), complete protease inhibitors (Roche), 2 µM dithiothreitol (DTT), and 10 µg/mL DNase I. Cells were incubated on ice for 30 min followed by 5 sonication steps (Fisher Dismembranator) at maximum amplitude with 1-2 minute ice incubations between steps. Sonicates were centrifuged at 12,000 r.c.f. for 30 min at 4˚C. Supernatants were transferred to new tubes and spun at 48,000 r.c.f. for 25 min at 4˚C. Supernatants were added to freshly swelled and equilibrated glutathioneagarose beads (Sigma). Lysates were batch bound to beads for 1 hour at 4˚C with rotating. Lysate + bead mixtures were transferred to a 10 mL BioRad Polyprep column and flowthrough discarded. Beads were washed consecutively with 20 mL PBS + 0.05% Triton X 100 (Tx100), 20 mL PBS + 0.05% Tx100 + 0.5M NaCl, and 20 mL PBS. Proteins were eluted with 10 mM glutathione in 50 mM Tris (pH 9.5) followed by dialysis into un-supplemented lysis buffer using SlideAlyzer dialysis cassettes with 10 kDa molecular weight cutoff (Pierce Scientific). Dialyzed proteins were stored at 4˚C until use and analyzed by SDS-PAGE to confirm purity. GST-HA3 and GST-HA3-NleC were quantified using a Bradford Protein Assay (BioRad Laboratories). Glutathione-agarose was swelled overnight at 4˚C in cold, sterile milliQ water. Fifty microlitres of beads was washed 5 x with 1 mL cold PBS-/- and equilibrated into 50 µL lysis buffer. Wash steps were performed with 800 r.c.f. spins at 4˚C for 5 min between buffer changes. Twenty microlitres of beads were transferred into a fresh tube and incubated with 20 µL of 20 pmol GST-HA3 or GST-HA3-NleC at 4˚C for 60 minutes  118  with rotation. Beads were spun at 2,100 r.c.f for 1 min and a thin layer of supernatant was left on the beads. Tubes were kept on ice until use.  5.2.6 SILAC HEK 293 cells were maintained and SILAC labeled as described previously (Rogers et al., 2008). Briefly, HEK 293 cells were split from normal growth media into SILAC labeling media. DMEM medium lacking arginine and lysine (Caisson Laboratories Inc) was supplemented with 10% dialyzed FBS, 1% NEAA, 1% GlutaMax, 0.1 units/L Penicillin/Streptomycin and either 36.5 mg/L  13  C6-arginine and 2H4-lysine (Cambridge  Isotope Laboratories, Andover, MA) for heavy cells or normal isotopic abundance Larginine and L-lysine for light cells (Sigma-Aldrich). Cells were maintained for five cell doublings to ensure complete labeling of the proteome and ten 10 cm cell culture dishes of each light and heavy labeled cells were solubilized in 750 µL cold NP-40 lysis buffer [1% Nonidet P40 (v/v), 20 mM Tris pH 7.5, 150 mM NaCl, 10 mM Na pyrophosphate, 50 mM NaF supplemented with 1 mM Na3VO4, and Protease Inhibitor Cocktail (Roche)] and centrifuged at 16,100 r.c.f. for 10 min at 4ºC. Protein concentrations of heavy and light lysates were measured using the Coomassie Plus Protein Assay (Pierce Scientific) according to manufacturer’s directions. Ten milligrams of lysate was transferred into fresh tubes and beads containing 20 pmol of recombinant protein were added. Tubes were incubated at 4˚C for 60 min with rotation and then spun in a Beckman Table Top centrifuge for 5 min at 2,100 r.c.f. at 4˚C. Beads were washed with 1 mL of cold NP40 lysis buffer followed by cold PBS. One hundred microlitres of 10 mM glutathione in 50  119  mM Tris (pH 9.5) was added to each tube and incubated overnight with rotation at 4˚C. Tubes were spun at 2,100 r.c.f. for 5 minutes at 4˚C and all remaining buffer was drawn from the beads using GELoader Tips (Eppendorf, Westbury, NY) until the beads were dry and all supernatants were pooled into one tube. The eluted proteins were then precipitated using ethanol/acetate and digested in-solution as previously described (Foster et al., 2003). Peptides were acidified and analyzed by liquid chromatography-tandem mass spectrometry on an LTQ-OrbitrapXL exactly as previously described (Rogers et al., 2008).  5.2.7 Immunoprecipitation HEK 293T cell lysates were generated in NP40 lysis buffer as described previously (Shames et al., 2010). Protein G-conjugated DynaL beads (Invitrogen) were bound to either rabbit α-p300 (Santa Cruz Biotechnology) and IC (normal rabbit IgG; Santa Cruz Biotechnology) or mouse α-FLAG (M2) (Sigma-Aldrich) and IC (normal mouse IgG; Jackson ImmunoResearch Labs). Immunoprecipitations were performed according to manufacturer’s instructions. Beads were resuspended in 20 µL 3X Laemmli sample buffer and boiled for 10 min followed by Western blot analysis.  5.2.8 Confocal microscopy Preparation of slides for laser scanning confocal microscopy was performed as described previously (Shames et al., 2010). For EPEC infected cells, coverslips were incubated in  120  1:200 rabbit α-p300 (Santa Cruz Biotechnology) in blocking buffer [PBST/BSA, 1% NGS] overnight at 4˚C. Following 3 x 10 min washes, a 1:200 dilution of Alexa 488conjugated goat α-rabbit antibody (Invitrogen) and 1:500 dilution of Alexa 633conjugated phalloidin (Invitrogen) in blocking buffer was added to the coverslips. Wash steps were repeated followed by mounting the coverslips on slides with ProLong Gold with DAPI (Invitrogen). HeLa cells transfected with eGFP constructs were stained with α-p300 and mounted in ProLong Gold with DAPI as described.  Quantification of  immunofluorescent images was performed by imaging greater than 50 cells per infection and counting the number of infected cells retaining p300 staining. Images were acquired on an Olympus Fluoview 10i laser scanning confocal microscope at the UBC Bioimaging Facility and processed using ImageJ and Adobe Photoshop.  5.2.9 Isolation of cellular nuclei HeLa or Caco-2 cells grown in 6 well plates that had been transfected, infected, or untreated were washed in PBS-/- and treated with 0.05% or 0.25% Trypsin-EDTA, respectively. Lifted cells were resuspended in DMEM and spun at 500 r.c.f. at 4˚C for 5 minutes. Whole cells were washed in PBS-/-. The NE-PER Nuclear Isolation Kit (Pierce Scientific) was used according to manufacturer’s instructions. Nuclear and cytosolic extracts were stored at -80˚C until use and quantified using the Coomassie Plus Protein Assay to quantify fractionated proteins. Equal amounts of protein (15 µg) were loaded on SDS-PAGE gels for Western blot analysis.  121  5.2.10 In vitro cleavage assay Recombinant GST-HA3-NleC and GST-HA3 were produced as described. Recombinant p300 protein was purchased (Active Motif, USA) and stored at -80˚C until use. p300 (50 nM) was mixed with either GST-HA3-NleC (0.3 nM) or GST-HA3 (0.3 nM) in Protease Cleavage Buffer (10 mM Tris-HCl pH 7.5, 150 mM NaCl, 2.5 mM CaCl2, 0.5 mM DTT, 0.5 mM MgCl2, 1 µM ZnCl2) with or without 10 mM EDTA. Reactions were carried out at 25˚C for 8 h, as described (Yen et al., 2010).  Proteins were separated and  immunoblotted as described.  5.2.11 Immunoblotting Proteins separated by denaturing SDS-PAGE were transferred to Pure Nitrocellulose (BioRad) using a wet transfer cell. Membranes were blocked in blocking buffer [5% non-fat milk in TBST] for 30 min at room temperature. Anti-HA (rat; Roche), α-Ac-K (rabbit; Cell Signaling Technology), α-Lamin B1 (rabbit; Abcam), α-calnexin (rabbit; Enzo Life Sciences), and α-p65 (rabbit; Cell Signaling Technology) were diluted in blocking buffer at 1:1000 and rabbit α-p300 (Santa Cruz Biotechnologies) was diluted 1:500. Membranes were incubated in primary antibody overnight at 4˚C with rocking. Wash steps were performed for 3 x 10 min in TBST. HRP-conjugated goat α-rat and goat α-rabbit antibodies were diluted 1:5000 in blocking buffer and incubated with membranes for 1 h. Membranes were washed as described above followed by addition of ECL reagent (Amersham) and chemiluminescent developing on Kodak BioMax film.  122  Membrane stripping was performed at room temperature by 5 min wash in distilled (d) H2O, 5 min wash in 0.2 N NaOH, and 5 min wash in dH2O.  5.2.12 siRNA siGENOME oligonucleotides targeting p300 (SMARTpool EP300) or nt (SMARTpool Pool  1)  (control)  were  purchased  from  Dharmacon  (Thermo  Scientific).  Oligonucleotides were suspended in 1X siRNA buffer (Dharmacon) and stored at -80ºC until use. siRNA (100 nM) was transfected into HeLa cells at 30% confluence using Oligofectamine (Invitrogen) according to manufacturer’s directions and media were changed 24 h post-transfection. assayed as described.  At 48 h post-transfection, cells were infected and  Lysates were collected for Western blot analysis to confirm  knockdown.  5.2.13 IL-8 ELISA Caco-2 or HeLa cells were maintained in culture medium as described above. Prior to infection, media were aspirated and cells were washed in PBS-/-. Cells were infected in 0.5 mL of serum-free DMEM per well for 3 h at 37˚C/5% CO2 in a 24 well dish (BD Falcon). Media were aspirated and cells were incubated in 0.5 mL of DMEM containing 5 ng/mL IL-1β (RnD Systems) and 100 µg/mL gentamicin (Sigma) for 3 h. Supernatants were harvested and stored at -80˚C until use. IL-8 sandwich ELISA was performed on  123  cell supernatants using the BD OptEIA™ Human IL-8 ELISA kit (BD Biosciences) according to manufacturer’s directions.  5.2.14 Statistical analysis Statistics were performed using an unpaired two-tailed t-test with a 95% confidence interval. In all experiments, error bars denote S.D. of samples in duplicates or triplicates. GraphPad software was used to determine p-values.  5.3 NleC interacts with host p300 by SILAC Identification of host proteins targeted by bacterial virulence factors can provide insight into their function during infection. We used the A/E type-III secreted effector NleC in a SILAC screen to identify putative host protein targets. Two constructs, NleC fused to GST and a triple HA tag (GST-HA3-NleC) or GST-HA3 alone, were either expressed directly in HEK293 cells as previously described (Shames et al., 2010, Rogers et al., 2008) or were produced in E. coli BL21 (DE3) and purified on glutathione-agarose resin. HEK 293T cells were grown in media lacking L-lysine and L-arginine and supplemented with either light (normal) or heavy (2H4-lysine and  13  C6-arginine) amino acids.  Exogenously-expressed constructs were immunoprecipitated from cell lysates and, in parallel, equal amounts of light and heavy 293T cell lysates were incubated with glutathione-agarose beads bound to GST-HA3-NleC and GST-HA3, respectively. Nonspecific proteins were removed via sequential washes and proteins remaining on the  124  column were eluted using 10 mM glutathione and then eluates were mixed prior to tryptic digestion. Peptides were then analyzed by LC-MS/MS. In this scheme, non-specifically interacting proteins should present a ratio near 1.0 and, indeed, peptides from peroxiredoxin 2 had a heavy: light ratio of ~1.1 (Fig 5.1A). However, 22 unique peptides of the E1A-associated protein p300 (herein called p300; Appendix B) all had low heavy: light ratios, indicating specific binding to NleC (Fig 5.1B). All proteins identified in this SILAC experiment with heavy: light isotope ratios <0.66 are listed in Table 5.4. As we have previously observed, heat shock proteins and ubiquitin are often observed as interactors by this technique (Rogers et al., 2008, Shames et al., 2010). CBP was also identified in this screen as a specific interactor to NleC (Table 5.4). Based on high homology between p300 and CBP and the lower average ratio and standard deviation for p300 peptides, we continued our investigation with p300 as a putative binding partner to NleC.  This SILAC experiment was performed in triplicates with similar results  suggesting that NleC interacts specifically with host p300.  Fig 5.1 Representative mass spectra for peptides recovered from SILAC experiments (A) Open and filled triangles indicate the expected mass: charge (m/z) of light and heavy forms, respectively, of a peptide (GLFIIDDK) from the non-specifically bound Peroxiredoxin 2 from GST-HA3-NleC or GST-HA3, respectively. (B) Open and filled triangles indicate the expected m/z of light and heavy forms, respectively, of a peptide  125  (QWHEDITQDLR) from the specifically bound p300 from glutathione agarose beads bound to GST-HA3-NleC or GST-HA3, respectively. The light peptide of p300 is present in a ratio of > 1.5 higher than the heavy peptide, indicating a specific interaction with NleC.  Table 5.4 Summary of proteins identified by MS as specifically interacting with GST-HA3-NleC in SILAC experiments Protein Averagea S.D.b NleC 0.02 0.01 Heat shock 70 kDa protein 1 0.10 0.06 Heat shock protein HSP90a2 0.24 0.11 Heat shock protein b 0.14 0.01 Heat shock protein HSP90b 0.24 0.11 CREB-binding protein 0.66 0.08 E1A-associated protein p300 0.65 0.03 Ubiquitin and S27a 0.25 0.10 a. Averaged SILAC ratio (heavy isotope: light isotope) b. S.D.: standard deviation as determined by quantification of several peptides from either two or three independent SILAC experiments 5.4 NleC co-immunoprecipitates with endogenous p300 To further confirm the interaction between NleC and p300, we performed coimmunoprecipitations  to  determine  if  α-p300  antibodies  could  specifically  immunoprecipitate NleC. We transfected pcDNA3::HA2nleC or pcDNA3::HA2 (vector) into HEK 293T cells and used an α-p300 antibody to pull-down endogenous p300 followed by Western blot analysis on the precipitated immune complexes.  We  determined that HA2NleC was specifically immunoprecipitated with α-p300 but not with an IC antibody (Fig 5.2). Based on these observations, we concluded that NleC interacts with host p300.  126  Fig 5.2 NleC co-immunoprecipitates with endogenous p300 HA2NleC was expressed in HEK 293T cells and subjected to co-immunoprecipitation with an α-p300 antibody. Normal rabbit IgG was used as an isotype control. Western blot analysis using α-p300 and α-HA was used to detect HA2NleC immunoprecipitated with endogenous p300 protein.  5.5 The NleC-p300 interaction is dependent on the TAZ1 domain of p300 The acetyltransferase p300 is a 300 kDa protein composed of several specific domains, including a HAT domain [for a review, see (Goodman et al., 2000)]. To determine which domain(s) of p300 are necessary for interaction with NleC, we generated several truncations of p300 with C-terminal FLAG-tag fusions (Fig 5.3A). Expression of p300 truncations was variable, with the N, C, and NuR-TAZ1 domains expressing at the highest levels in the presence of NleC (Fig 5.3A, B). In the absence of NleC, expression levels of the NuR-KIX truncation were increased (Fig 5.3B). Anti-calnexin was used as a loading control (Fig 5.3B).  Using α-FLAG antibodies, HA2NleC was specifically  immunoprecipitated with the N-terminal region of p300 containing nuclear hormone receptor (NuR), TAZ1, and KIX domains (p300N-FLAG) (Fig 5.3C). We observed a  127  faint band of the approximate molecular weight of HA2NleC in co-immunoprecipitated samples containing p300C; however, this band appeared to be non-specific (Fig 5.3C; Fig 5.4) and therefore the C-terminal region of p300 containing the HAT domain does not interact with NleC (Fig 5.3C; Fig 5.4). We further truncated the p300N region into its component domains consisting of NuR, TAZ1, and KIX domains.  FLAG fusion  constructs containing only NuR, TAZ1, KIX, or TAZ-KIX could not be detected in lysates of transfected HEK 293T cells; however, we were able to specifically immunoprecipitate HA2NleC with the NuR-TAZ1-FLAG fusion protein using an αFLAG antibody (Fig 5.3C). This result suggested that the KIX domain was not necessary for interaction between NleC and p300. To determine if the TAZ1 domain contributed to interaction with NleC, we generated a NuR-KIX-FLAG fusion construct. HA2NleC was not immunoprecipitated by NuR-KIX-FLAG (Fig 5.3C). These data suggest that the TAZ1 domain of p300 is required for interaction with NleC.  128  Fig 5.3 The TAZ1 domain of p300 is required for interaction with NleC. (A) Schematic diagram depicting truncations of p300 cloned into pCMV-Tag4A upstream of a FLAG epitope tag. Expression levels in HEK 293T cells are shown in the right panel. (B) Western blot analysis of cell lysates from HEK 293T cells co-expressing p300-FLAG truncations and either HA2 or HA2NleC constructs used as input for coimmunoprecipitation experiments. Asterisks (*) indicate non-specific bands. (C) Coimmunoprecipitation with α-FLAG antibody. Inputs from (B) were subjected to immunoprecipitation with either mouse α-FLAG or mouse IgG IC. Arrows indicate HA2NleC and asterisks (*) indicate antibody heavy chain bands.  129  Fig 5.4 NleC does not co-immunoprecipitate with the C-terminus of p300 (A) Lysates from HEK 293 cells expressing p300C-FLAG and either HA2 or HA2NleC were subjected to immunoprecipitation using mouse α-FLAG antibodies or mouse IgG isotype control followed by Western blot analysis using α-FLAG or α-HA. (B) Western blot analysis of input lysates from HEK 293 cells co-expressing p300C-FLAG and either HA2 or HA2NleC.  5.6 Global acetylation levels in Caco-2 cells are increased during infection with EPEC ∆nleC To determine if NleC interferes with the function of p300 during EPEC infection, we examined global acetyl-lysine (Ac-K) levels. Caco-2 cells were infected for 4 h with EPEC WT, ∆nleC, ∆nleC/nleC, ∆escN or mock infected followed by cell lysis and nuclear extraction. Cytosolic and nuclear proteins were separated by SDS-PAGE and immunoblotted using antibodies specific to Ac-K or calnexin (loading control). Infection with EPEC ∆nleC resulted in increased band intensity at approximately 75 kDa, 80 kDa, and 30 kDa compared to infection with WT, ∆nleC/nleC, or ∆escN strains (Fig 5.5A). Increased Ac-K appeared to occur exclusively in the cytosolic fraction of EPEC ∆nleC-  130  infected Caco-2 cells (Fig 5.5A). Anti-Calnexin was used to confirm equal protein loading and separation of cytosolic from nuclear proteins (Fig 5.5B). These data suggest that NleC dampens the acetyltransferase activity of p300 during EPEC infection.  Fig 5.5 Global acetylation levels in Caco-2 cell cytosol is increased during infection of Caco-2 cells with EPEC ∆nleC Caco-2 cells were infected with EPEC WT, ∆nleC, ∆nleC/nleC, ∆nleC/nleCHA, ∆escN, or left uninfected. Cells were lysed and nuclei were isolated as described. (A) Western blot analysis was used to observe global acetylation in the cytosol and nuclei of infected cells using an α-Ac-K antibody. (B) Membranes were stripped and re-probed using αcalnexin as a loading control.  5.7 NleC facilitates decreased nuclear p300 levels in HeLa cells Since NleC is a Zn-metalloprotease, we aimed to determine if NleC decreases p300 abundance in the context of EPEC infection. HeLa cells were infected and p300 was  131  imaged by immunofluorescence. HeLa cells grown on glass coverslips were infected with either WT, ∆nleC, ∆nleC/nleC, ∆escN, or not infected (Mock). Following infection, coverslips were harvested, fixed and stained with rabbit α-p300 followed by Alexa 488conjugated goat α-rabbit antibody (green). Alexa 633-conjugated phalloidin (red) was used to visualize actin and nuclei were imaged with DAPI. Concentrated phalloidin staining indicates EPEC-induced pedestals (Fig 5.6A, arrows). We observed HeLa cells devoid of p300 following infection with WT and ∆nleC/nleC EPEC strains (Fig 5.6A, arrowheads). Infected cells positive for p300 staining were quantified and plotted as percent cells retaining p300. We found significantly more cells retaining nuclear p300 during infection with EPEC ∆nleC compared to infection with EPEC WT and ∆nleC/nleC (*, p < 0.05) (Fig 5.6B).  These data suggest that NleC can facilitate  decreased nuclear p300 protein levels.  132  Fig 5.6 NleC facilitates decreased nuclear p300 levels (A) Immunofluorescent analysis of HeLa cells infected with either EPEC WT, ∆nleC, ∆nleC/nleC, ∆escN or left uninfected. Cells were infected for 3 hours on glass coverslips followed by fixation and permeabilization. Cells were stained with α-p300 and 133  subsequently Alexa 488-conjugated goat α-rabbit antibody and Alexa 633-conjugated phalloidin to stain cellular actin. Coverslips were mounted in medium containing DAPI to stain bacteria and cellular nuclei. Images were acquired on an Olympus Fluoview 10i laser scanning confocal microscope. White arrows show cells infected with EPEC and white arrowheads show cells exhibiting decreased staining of nuclear p300. Scale bar represents 15 µm. (B) Quantification of infected HeLa cells retaining nuclear p300 staining. Cells positive for EPEC pedestals and p300 staining were quantified and plotted as percentage of infected cells containing nuclear p300. Asterisks (*) denote statistical significance (p < 0.05) of samples in triplicate. Data are representative of two independent experiments.  To further determine if NleC facilitates degradation of p300, we overexpressed NleC ectopically in HeLa cells and examined p300 protein levels by Western blot. We transfected pcDNA3::HA2nleC (HA2NleC) or pcDNA3::HA2 (Vector) into HeLa cells followed by isolation of cellular nuclei. For Western blot analysis, equal amounts of protein (15 µg) were loaded in each lane followed by immunoblot with α-HA, α-p300, and α-Lamin B1 as a nuclear loading control. p300 was not observed in cytosolic extracts and HA2NleC appeared to preferentially localize to nuclei (Fig 5.7A, B). In the nuclei of HeLa cells expressing HA2NleC, p300 levels were decreased (Fig 5.7B). Densitometric analysis revealed that p300 protein levels were significantly decreased in HeLa cells expressing HA2NleC compared to HeLa cells transfected with vector alone (*, p < 0.05) (Fig 5.7C).  p300 levels were also decreased with increasing levels of  ectopically expressed HA2NleC in Caco-2 cells (Fig 5.8). These data suggest that NleC facilitates a decrease in nuclear p300 levels.  134  Fig 5.7 Ectopic expression of NleC decreases nuclear p300 protein levels in HeLa cells (A) Western blot analysis of cytosolic proteins from cells transfected with 2 µg of pcDNA3::HA2nleC or pcDNA3::HA2. Western blot analysis was performed with α-p300, α-p65, α-HA, and α-calnexin. (B) Western blot analysis of nuclear proteins following transfection with 2 µg pcDNA3::HA2nleC or pcDNA3::HA2. Western blot analysis was performed with α-p300, α-HA, and α-Lamin B1 as a nuclear loading control. (C) Densitometric analysis of nuclear p300 band intensity. Pixel intensities of bands corresponding to p300 were quantified using ImageJ. Relative amounts of p300 are shown as a function of the loading control, Lamin B1. Asterisks (*) indicate statistical significance (p < 0.05) of samples from two independent experiments.  Fig 5.8 Increasing concentration of HA2NleC causes decreased p300 protein levels in Caco-2 cells Caco-2 cells were transfected with increasing concentrations of pcDNA3::HA2nleC and lysates were generated. Western blot analysis was performed with α-p300, α-HA, and αCalnexin (loading control). Amounts of pcDNA3::HA2nleC are 0.1, 0.3, 0.5, 0.7, and 0.9 ng plasmid DNA topped up to 1.0 µg with pcDNA3::HA2.  135  5.8 The metalloprotease domain of NleC contributes to decreased nuclear p300 NleC contains an HEXXH metalloprotease motif, which is responsible for p65 degradation (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011). We therefore hypothesized that the metalloprotease activity of NleC may also be required for p300 degradation. Baruch and colleagues identified amino acid residue 184 as the catalytic glutamate in the HEIIH motif of NleC and showed that an E184A mutation abolished its activity (Baruch et al., 2011). We generated the same mutation in nleC cloned upstream of GFP (pEHnleCE184A-GFP) or a FLAG epitope tag (pCMV4A::nleCE184A). Initially, we determined that NleCE184A is still able to interact with endogenous p300 (Fig 5.9). We then used pEHnleCE184A-GFP in addition to the wild type nleC gene (pEHnleC-GFP) and GFP-only vector (pEGFP) to transfect HeLa cells. HeLa cells expressing eGFP, NleC-eGFP, or NleCE184A-eGFP were stained with rabbit α-p300 followed by Alexa 633-conjugated goat α- rabbit antibody (red). Coverslips were mounted in ProLong Gold with DAPI (blue). HeLa cells expressing NleC-eGFP showed decreased p300 staining (Fig 5.10A). However, cells expressing NleCE184A-eGFP had similar p300 staining to HeLa cells expressing only GFP (Fig 5.10A). We also used the metalloprotease inhibitor GM6001, which was previously shown to abrogate NleC-mediated p65 cleavage (Muehlen et al., 2011), to treat cells transfected with NleC-eGFP. When compared to vehicle (DMSO) treated cells, GM6001 treatment inhibited p300 degradation by NleC (Fig 5.10A).  136  Fig 5.9 NleCE184A interacts with endogenous p300 HEK 293T cells were transfected with pCMV4A::nleC or pCMV4A::nleCE184A and lysates were subjected to immunoprecipitation using rabbit α-p300 or IgG isotype control. Input lysates were immunoblotted for endogenous p300 or FLAG-tagged nleC constructs. Immunoprecipitated lysates were subjected to Western blot analysis with αp300 and α-FLAG antibodies. NleC-FLAG and NleCE184A-FLAG were immunoprecipitated with α-p300 antibodies. We further confirmed involvement of the NleC metalloprotease domain in p300 degradation by Western blot analysis using HeLa cells expressing HA2NleC.  We  transfected HeLa cells with either pcDNA3::HA2nleC or empty vector and treated cells with either GM6001 or vehicle control (DMSO).  As an additional control, we  immunoblotted with α-p65 and observed decreased levels of p65 in cells expressing HA2NleC and treated with DMSO (Fig 5.10B, C). Decreased nuclear p300 was observed in cells expressing HA2NleC and treated with DMSO but not with GM6001 (Fig 5.10B). Expression of HA2NleC had no impact on p65 or p300 levels in cells treated with GM6001; however, with DMSO treatment, cells expressing HA2NleC had significantly  137  less p300 than those transfected with empty vector (*, p < 0.05) (Fig 5.10C). These data suggest that the metalloprotease activity of NleC is required for p300 degradation.  138  Fig 5.10 The metalloprotease domain of NleC contributes to decreased nuclear p300 (A) Immunofluorescent analysis of HeLa cells transfected with either pEGFP, pEHnleCGFPN, or pEHnleCE184A-GFPN. Cells expressing either eGFP, NleC-eGFP, or NleCE184A-eGFP (green) were either left untreated or treated with GM6001 or volume equivalent of DMSO (vehicle control) where indicated. Cells were immunostained with α-p300 followed by Alexa 633-conjugated goat α-rabbit antibody (red) and coverslips were mounted in medium containing DAPI (blue) to image cellular nuclei. White arrows indicate cells expressing GFP-containing proteins. Images were acquired on an Olympus Fluoview 10i laser scanning confocal microscope. Scale bar represents 20 µm. (B) Western blot analysis of HeLa cell nuclei expressing either HA2NleC or HA2 and treated with either GM6001 (+) or DMSO (-). Cellular nuclei were isolated and immunoblotting with α-p300, α-p65, α-HA, and α-Lamin B1 (loading control) was performed. (C) Densitometric analysis of p300 (right panel) and p65 (left panel) protein levels in cells expressing HA2NleC or HA2 and treated with either GM6001 or DMSO. Asterisks (*) indicate statistical significance compared to cells treated with GM6001 (p < 0.05) of samples from two independent experiments.  5.9 NleC facilitates p300 degradation in vitro Next, the ability of recombinant NleC to cleave recombinant p300 protein in vitro was investigated. To determine if NleC degrades p300 in vitro, we performed a protease cleavage assay using equal amounts of recombinant p300 in the presence of either GSTHA3-NleC or GST-HA3. Degradation of p300 was analyzed by Western blot. We found that addition of GST-HA3-NleC, but not GST-HA3, caused a reduction in p300 protein levels compared to input levels (Fig 5.11). In the presence of the divalent ion chelator EDTA, recombinant NleC was unable to facilitate a reduction in p300 protein (Fig 5.11). Together, these data indicate that NleC is sufficient to mediate cleavage of p300 and that the Zn-metalloprotease activity of NleC is required.  139  Fig 5.11 NleC cleaves p300 in vitro Purified p300 protein was incubated with either GST-HA3-NleC or GST-HA3 in the presence or absence of 10 mM EDTA for 8 h at 25˚C. Reactions were quenched as described and p300 protein levels were determined by Western blot analysis using αp300 antibodies.  5.10 Overexpression of p300 enhances IL-8 secretion during WT EPEC infection NleC contributes to repression of IL-8 secretion during EPEC infection by degradation of NF-κB p65 (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011).  p300 is necessary for expression of IL-8 in cells stimulated with pro-  inflammatory stimuli (Hoffmann et al., 2002) and we therefore hypothesized that NleCmediated degradation of p300 contributes to suppression of IL-8 secretion in stimulated cells. We infected Caco-2 cells that had been transfected with an empty vector or pCMVβ::p300HA with WT, ∆nleC, or ∆nleC/nleC EPEC strains followed by stimulation with IL-1β. IL-8 present in cell supernatants was quantified using an IL-8 ELISA. As previously observed in HeLa cells (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011), Caco-2 cells infected with ∆nleC secreted significantly more  140  IL-8 than cells infected with WT or the ∆nleC/nleC complemented strain (p < 0.05) (Fig 5.12A). We found that Caco-2 cells overexpressing p300 secreted significantly more IL8 during infection than cells harboring empty vector following infection with WT EPEC (p < 0.05) (Fig 5.12A).  To confirm overexpression of p300 in those samples, we  collected lysates from infected cells and performed Western blot analysis with α-p300, α-HA, and α-calnexin as a loading control. We determined that p300HA was indeed expressed in Caco-2 cells transfected with pCMVβ::p300HA and that total p300 levels were increased in those cells (Fig 5.12B). These data indicate that overexpression of p300 facilitates increased IL-8 secretion during infection with WT EPEC.  Fig 5.12 Over-expression of p300 causes enhanced IL-8 secretion during EPEC infection (A) Caco-2 cells were transfected with either pCMVβ::p300HA (p300HA) or pCMV4A (empty vector; EV) followed by infection with EPEC WT, ∆nleC, or ∆nleC/nleC, as indicated. ELISA for IL-8 present in cellular supernatant was performed following infection and stimulation with IL-1β. Concentrations of IL-8 (pg/mL) from cellular  141  supernatants are shown. Asterisks (*) indicate statistical significance (p < 0.05). Data are representative of three independent experiments. (B) Western blot analysis of Caco-2 cell lysates following infection with EPEC strains. Lysates were immunoblotted with αHA, α-p300, and α-calnexin (loading control) to confirm overexpression of p300HA. Data are representative of three independent experiments.  5.11 Knock-down of p300 dampens IL-8 secretion from cells infected with EPEC ∆nleC To determine if NleC-mediated decrease in host p300 levels contributes to decreased IL-8 secretion during EPEC infection, we utilized p300-specific siRNA. HeLa cells were transfected with nt (control) siRNA or p300-specific siRNA prior to infection with EPEC WT, ∆nleC, or ∆nleC/nleC strains and stimulation with IL-1β.  IL-8 secretion was  quantified by sandwich ELISA. Following transfection with nt siRNA, IL-8 secretion was suppressed in cells infected with EPEC WT and ∆nleC/nleC compared to cells infected with EPEC ∆nleC (b, p < 0.05) (Fig 5.13A). However, we found that siRNA knock-down of p300 resulted in significantly less IL-8 secretion during infection with EPEC ∆nleC (b, p < 0.05) (Fig 5.13A). To confirm p300 knock-down, lysates were collected from infected HeLa cells treated with either nt (control) or p300-specific siRNAs. Western blot analysis revealed that p300-specific siRNAs facilitated a decrease in p300 protein levels (Fig 5.13B). These data suggest that decreasing total p300 protein levels contributes to suppression of IL-8 secretion during EPEC infection.  142  Fig 5.13 siRNA knock-down of p300 dampens IL-8 secretion during EPEC ∆nleC infection (A) HeLa cells were transfected with either nt or p300-specific siRNA followed by infection with EPEC WT, ∆nleC, or ∆nleC/nleC, as indicated. ELISA for IL-8 present in cellular supernatant was performed following infection and stimulation with IL-1β. Concentrations of IL-8 (pg/mL) from cellular supernatants are shown. b indicates statistical significance compared to a (p < 0.05). Data are representative of two independent experiments. (B) Western blot analysis of HeLa cell lysates following siRNA treatment and EPEC infection. Lysates were immunoblotted with α-p300 and αcalnexin (loading control) to confirm knock-down of p300. Data are representative of two independent experiments.  5.12 Discussion The ability of A/E pathogens to decrease pro-inflammatory signaling has been well established and recently, T3S effector proteins contributing to this phenotype have begun to be identified. Of all A/E effectors identified that impact on NF-κB signaling, and resultantly IL-8 secretion, only NleC was shown to be a protease that specifically targets a member of the NF-κB protein family, p65 (Shames et al., 2011c). Cleavage of p65 by  143  NleC was shown to result in enhanced repression of IL-8 secretion and gene expression following treatment with pro-inflammatory stimuli.  In this study, we attempted to  identify additional targets of NleC within host cells and determine if they contribute to repression of IL-8 secretion by EPEC. As an approach to identify host protein targets of NleC, we first employed SILAC. We rationalized that targets of NleC would be bound specifically by the enzyme prior to their degradation. Our screen identified several mammalian proteins as specifically interacting with NleC (Table 5.4). Since heat shock proteins and ubiquitin are very common in these assays (Shames et al., 2010, Rogers et al., 2008), we focused our attention on host p300 based on its importance in IL-8 gene expression and its high homology to the other putative interactor CBP. p300 is a well-studied acetyltransferase that functions as a coactivator in the transcription of many genes within the host cells. Acetylation of histone tails by p300 causes them to lose electrostatic interaction with DNA in chromatin and allows subsequent transcription of target genes. In addition to histone tails, p300 has been demonstrated to acetylate p65 and the tumor suppressor p53 (Chen et al., 2002, Chen et al., 2001, Itahana et al., 2009). p300 is targeted by a plethora of viral proteins and was identified by its interaction with the E1A protein of adenovirus (Whyte et al., 1989, Hottiger et al., 2000). Other viral proteins targeting p300/CBP include E6 and E7 from the human papilloma virus (HPV) (Patel et al., 1999), Tat from human immunodeficiency virus (HIV), Zta and EBVNA-2 from Epstein-Barr virus, and viral interferon responsible factor (vIRF) from Karposi’ssarcoma virus (Hottiger et al., 2000). Recently, the ORV002 protein from Orf virus (ORV) was shown to inhibit nuclear NF-κB by reducing its interaction with p300 (Diel et 144  al., 2011). Viruses have evolved to hijack p300 and CBP based on their essential role in cellular transcription, cell cycle progression, and regulation of the tumor suppressor p53 (Hottiger et al., 2000). Interestingly, only the E6 and E7 proteins of HPV have been shown to subvert p300 for the purposes of dampening IL-8 secretion (Huang et al., 2002). The Gram-negative bacterial pathogen Legionella pneumophila relies on histone acetylation to mediate pro-inflammatory signaling; however, no L. pneumophila virulence factors have been attributed to this phenotype (Schmeck et al., 2008). To date, NleC is the first bacterial protein shown to interact with and subvert the acetyltransferase p300, joining many viral effectors that target this host protein. Following identification of p300 as a host protein interactor to NleC, we aimed to determine which region of p300 was targeted by NleC for interaction.  We made  sequential truncations of p300 and performed co-immunoprecipitations with epitopetagged NleC and determined that the TAZ1 domain of p300 is required for immunoprecipitation of NleC.  The TAZ1 domain functions as a protein-protein  interaction scaffold containing a zinc-finger domain (De Guzman et al., 2005), which has been shown to interact with many different proteins, including NF-κB p65, TATAbinding protein, Stat-2, and p53 [for review on p300 interactions, see (Goodman et al., 2000)]. Since p300 acetylates p65 to enhance expression of genes downstream of κBcontaining enhancers (Chen et al., 2001), we initially assayed for global cellular acetylation during infection with EPEC strains and found that total acetylation was increased in Caco-2 cells during infection with EPEC ∆nleC. This observation further supports our data that p300 is affected by NleC during EPEC infection. In addition, it is likely that cleavage of p300 is a slower event than cleavage of p65 since interaction  145  between p65 and full-length NleC could not be observed (Muehlen et al., 2011). Interestingly, expression of the N-terminal region of p300 lacking the TAZ1 domain (Nur-KIX; NK) was present in lower levels in cells co-expressing NleC, compared to vector control.  We demonstrated that NleC was not pulled-down with this p300  truncation, suggesting that the cleavage site may be in the KIX region followed by release of the protein. Other truncations expressed variably regardless of NleC presence, which is likely due to stability of the truncated p300 fragments. During the course of our work, four elegant studies were published demonstrating that NleC degrades p65 via a Zn-metalloprotease domain (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011). We had also observed that infection with an EPEC ∆nleC strain caused significantly increased NF-κB activity in Caco-2 cells; however, the aforementioned studies led us to investigate whether p300 is also a substrate for the enzymatic activity of NleC. We initially observed that a small population of cells infected with WT and ∆nleC/nleC EPEC strains was lacking nuclear p300 staining by immunofluorescence. Quantification of infected cells retaining p300 demonstrated that significantly less cells infected with EPEC encoding nleC retain p300 staining. The relatively modest degradation of p300 by NleC, in comparison to p65, is likely due to intricate regulation of p300 degradation by NleC, since p300 controls many normal metabolic and physiological processes in the cell (Ghosh et al., 2007, Goodman et al., 2000), including apoptotic cell death (Tyteca et al., 2006).  We and others have  previously demonstrated that EPEC actively reduces premature cell death during infection (Shames et al., 2010, Hemrajani et al., 2010) and this may explain the subtle extent of NleC-mediated p300 degradation during EPEC infection.  146  The requirement of the metalloprotease domain for NleC-mediated p300 degradation was not surprising. We show that the same mutation of the catalytic glutamate (E184A) made by Baruch and colleagues does not result in decreased abundance of nuclear p300 and that treatment with the matrix metalloprotease inhibitor GM6001 abrogates the protease activity of NleC against p65 and p300. During our immunofluorescent analyses, we observed irregularly shaped nuclei in cells expressing enzymatically active NleC. This phenotype is very intriguing considering the important role p300 plays in the cell cycle and normal cellular physiology (Goodman et al., 2000). Further investigation is required to determine the significance of this observation and if it is linked to NleC-mediated subversion of p300. Degradation of p65 by NleC has a striking impact on NF-κB activity and IL-8 secretion during EPEC infection (Pearson et al., 2011, Muehlen et al., 2011, Yen et al., 2010, Baruch et al., 2011). p300 plays a role in expression of the IL-8 gene by NF-κB (Roebuck, 1999) and this prompted us to examine the impact of p300 on NleC-mediated repression of IL-8 secretion. The physiologically relevant intestinal epithelial cell line, Caco-2, was used for functional readout during EPEC infection. We hypothesized that overexpressing p300 would increase IL-8 secretion during WT EPEC infection. We were able to confirm previous work showing that infection with a ∆nleC strain of EPEC causes significantly more IL-8 secretion from infected cells than WT or the ∆nleC/nleC complemented strain. Indeed, overexpressing p300 in Caco-2 cells caused significantly greater IL-8 secretion compared to cells harboring empty vector during WT EPEC infection. Thus, increased levels of p300 may be overwhelming for the amount of NleC translocated by WT EPEC. In addition, p65 protein levels do not decrease in Caco-2  147  cells following infection with WT EPEC, and overexpression of nleC on a plasmid was required for cleavage of p65 to be observed (Baruch et al., 2011).  We had also  hypothesized that reduction of p300 contributes to dampening of IL-8 secretion during EPEC infection and to test this, we utilized siRNA to knock-down p300. During EPEC ∆nleC infection, reduction of p300 protein was sufficient to decrease IL-8 secretion to levels observed during WT EPEC infection. Several A/E pathogen T3S effector proteins play a role in decreasing IL-8 secretion during infection and dampening innate immune signaling is critical for these pathogens to efficiently cause disease in their host. Although others have observed that murine infection with C. rodentium ∆nleC does not cause greater bacterial burden or colon weights (Kelly et al., 2006), it is possible that investigation of other phenotypes in vivo may reveal the role of NleC during C. rodentium infection.  148  Chapter 6. Discussion Work presented in this thesis has detailed novel mechanisms by which A/E pathogen effector proteins can influence host cell death and innate immune signaling pathways during infection. EspZ interacts with CD98 and TIM17b to promote host cell survival during EPEC infection and NleC reduces IL-8 secretion by inhibition of NF-κB function and p300 degradation. In this chapter, we discuss other bacterial pathogens that utilize similar virulence mechanisms to EPEC and their ability to use effector proteins to efficiently cause disease. In addition, insight gained from the work presented in this thesis and future directions will be addressed. 6.1 Protection of host cells from cytotoxicity in bacterial pathogenesis Bacterial pathogens have evolved extraordinary mechanisms to efficiently infect host organisms. A majority of these pathogens do so by delivering virulence factors into host cells, which act to impair host defenses or utilize the host as a niche for replication. Although regulation of virulence factor expression by bacterial pathogens is a wellknown pathogenic mechanism (Mekalanos, 1992), the concept of host-protective virulence factors is emerging. In addition to that observed for EspZ in Chapter 2 and Chapter 3 of this thesis, several strategies by which pathogens appear to attenuate their own lethality towards host cells have been documented, suggesting that increased hostility and damage of host cells is not necessarily beneficial to the pathogen. Virulence is often defined as the ability of a pathogen to inflict damage on host cells, and the following discussion addresses the concept that increased virulence is not always advantageous for the pathogen, and moderating it to preserve host cells is a mechanism several pathogens use as part of their overall pathogenic strategy. This strategy is well 149  known for obligate intracellular pathogens, but has become an emerging theme in extracellular and facultative intracellular bacteria. In addition to A/E pathogens, Yersinia spp., Shigella flexneri, and Helicobacter pylori are well known for their ability to kill host cells.  For Yersinia, death of infected  macrophages decreases cytokine release and enables the pathogen to propagate with minimal challenges from the immune system (Aepfelbacher et al., 2007). Two recent studies suggest that host cell cytotoxicity caused by Yersinia species is tightly regulated. Y. pestis, the etiologic agent of plague, and gastroenteritis-inducing Y. pseudotuberculosis and Y. enterocolitica all encode a cytotoxic virulence factor called YopJ/P (YopJ in the two former species and YopP in the latter), which is translocated into infected cells via a T3SS (Aepfelbacher et al., 2007, Mills et al., 1997). Altering the cytotoxicity of Y. pseudotuberculosis affects its virulence. Decreased secretion of YopJ was shown to enhance Y. pseudotuberculosis pathogenesis in vivo (Brodsky et al., 2008). Similarly for Y. pestis, enhanced cytotoxicity results in decreased incidence of pneumonic plague in vivo (Zauberman et al., 2009). Tight regulation of cytotoxicity by pathogenic Yersinia is an efficient virulence strategy. Increased apoptosis of infected immune cells decreases production of proinflammatory cytokines; however, some inflammation at the early stages of infection is thought to facilitate tissue damage necessary for movement of bacteria and infected cells to other sites of replication within the host (Brodsky et al., 2008). Helicobacter pylori causes apoptosis of infected gastric epithelial cells (Moss et al., 1996). Apoptosis induction by H. pylori has been linked to a secreted toxin called VacA, which induces cytochrome c release from mitochondria (Fig 6.1) (Galmiche et al., 2000). 150  Recently, it was determined that VacA-mediated apoptosis is counteracted by a type IV secreted protein called CagA by both blocking pinocytosis of VacA and inhibiting VacA mediated cytochrome c release from mitochondria (Oldani et al., 2009) (Fig 6.1). Interestingly, loss of CagA in a VacA+ H. pylori strain decreases bacterial colonization and the incidence of gastric hyperplasia, adenocarcinoma, and inflammation (Franco et al., 2008). Similarly to the aforementioned pathogens, H. pylori has evolved a delicate interplay between host-protective and -detrimental virulence factors that are able to finetune virulence while promoting their propagation. Shigella flexneri, the etiologic agent of bacillary dysentery, causes death of infected macrophages and epithelial cells (Carneiro et al., 2009). Despite this, several hostprotective strategies are employed by S. flexneri. The T3S effector OspE was recently found to enhance adhesion of infected host cells to the underlying extracellular matrix (Kim et al., 2009b). Whether OspE activates host cell survival pathways directly is unknown; however, its interaction with integrin-linked kinase inhibits sloughing of infected cells into the intestinal lumen (Kim et al., 2009b), consequently preventing anoikis of Shigella infected cells. An ospE mutant does not colonize as efficiently as WT S. flexneri in vivo, and OspE may thus enhance colonization by preventing premature release of infected cells (Kim et al., 2009b). Epithelial cells succumb to S. flexneri infection via necrotic cell death, which functions to release intracellular bacteria and enhance inflammation (Carneiro et al., 2009). Interestingly, survival pathways involving receptor-interacting protein 2 (Rip2)/IKKβ/NFκB are activated by nucleotide-binding oligomerization domain-containing protein 1 (NOD1) early during infection followed by mitochondrial dysfunction and necrotic cell death (Fig 6.1) (Carneiro et al., 2009). The  151  early expression of pro-survival genes may enable S. flexneri to postpone cell death in a similar manner to EPEC, thus ensuring greater bacterial load prior to dissemination. The mechanism(s) by which S. flexneri enhances NFκB-mediated pro-survival signals are unknown. In addition to EPEC, the aforementioned pathogens have evolved strategies to attenuate their own virulence towards host cells and investigation of pathogenic mechanisms utilized by other pathogens can provide insight into EPEC-mediated disease. In many of these scenarios, removal of host-protective mediators actually reduces pathogenicity of the bacteria. The observation that EPEC encodes a host-protective virulence factor, EspZ [see Chapters 2 & 3] that is essential for its pathogenesis, suggests that protecting host cells may be a key to the pathogenic strategies of other bacterial pathogens. The concept of host-protective virulence factors is only just emerging, and we believe host-protective virulence factors will become more apparent in other pathogenic strategies.  Future  studies into pathogenic mechanisms of virulent bacteria will likely reveal important roles for effectors or regulatory mechanisms that help the host cell and promote bacterial pathogenesis.  152  Fig 6.1 Strategies evolved by bacterial pathogens to restrain virulence (A) EPEC injects effector proteins into IECs via a T3SS. EspF localizes to mitochondria and causes release of cytochrome c into the host cell cytosol, which results in apoptotic death of the host cells. NleH interacts with BI-1, which inhibits release of cytochrome c from mitochondria. EspZ interacts with TIM17b and CD98 to promote host cell survival. CD98 enhances survival via FAK-mediated signaling during EPEC infection. Localization of NleH and EspZ in host cells during early stages of EPEC infection is unclear and has been portrayed as shown for simplicity. (B) Helicobacter pylori injects virulence factors into gastric epithelial cells via a type IV secretion system in addition to secreting soluble toxins. VacA is an H. pylori secreted toxin that enters cells by pinocytosis and penetrates intracellular endosome trafficking pathways. VacA causes release of cytochrome c from mitochondria of infected cells, thus mediating host cell apoptosis. CagA is a type IV secreted virulence factor, which prevents both pinocytosis/trafficking and cytochrome c release by VacA. Functions of CagA are dependent on its phosphorylation state, not depicted here. (C) S. flexneri enters IECs from their basolateral surface and then resides in the cell cytoplasm. Prosurvival signaling is initiated by NOD1 activation of Rip2 signaling, which results in expression of pro-survival genes, including Bcl-2, via NF-κB activation and nuclear translocation. Conversely, S. flexneri facilitates a decrease in the Bcl-2/Bnip3 ratio, which leads to CypD-mediated disruption of mitochondria and oxidative stress-induced necrotic cell death.  153  6.2 Interplay between A/E T3S effectors in host cells as a mechanism for highly regulated cytotoxicity? The hierarchy and temporal regulation of effector secretion by A/E pathogens dictates the abundance and order in which these proteins are injected into infected host cells. Regulation of A/E pathogen effector gene expression and secretion has been investigated previously but is still largely unknown (Deng et al., 2005, Mills et al., 2008). Several T3S effector proteins localize to host mitochondria, but the order of their incorporation and their duration at this organelle remains elusive. EspF and Map cause mitochondrial dysfunction and both rely on host mitochondrial import machinery for their localization (Nagai et al., 2005, Papatheodorou et al., 2006).  EspZ also localizes to host  mitochondria and interacts with a member of the IMM transport complex, TIM17B (see Chapter 3). Since EspZ antagonizes cytotoxicity and is translocated prior to both EspF and Map (Mills et al., 2008), it is tempting to speculate that EspZ may limit access of cytotoxic effectors into the mitochondria or modulate their function once there. Since TIM17B plays a role in host cell death in response to WT EPEC infection (see Chapter 3), it may be involved in effector import. Despite the ability of EspZ to facilitate host cell survival, the eventual outcome of infection is host cell death (Crane et al., 1999). The contribution of several virulence factors to host cell death underlies its importance for infection. It is tempting to speculate that host cell death can facilitate detachment of intimately attached bacteria into the gut lumen where they can be subsequently shed into the environment. However, premature shedding of the pathogen would not enable a high enough bacterial burden to efficiently colonize a naïve host especially since EPEC requires a high infectious dose for healthy adults (Nataro et al., 1998). Highly regulated  154  EPEC-mediated host cell death involving interplay between several effectors was recently demonstrated by Dean and colleagues. These authors demonstrated that EspG1 and EspG2 facilitate activation of the host cysteine protease calpain, which results in potent host cell death and that Tir could abrogate this phenotype independently of its interaction with Intimin (Dean et al., 2010a). Mechanisms used by Tir to antagonize EspG-mediated calpain activation and host cell death have not been explored and investigation may reveal novel signaling cascades subverted by Tir to promote host cell survival during EPEC infection. Understanding the individual contributions of cytotoxic and pro-survival effectors is the first step to understanding their role in EPEC pathogenesis; however, examination of their interplay during infection will provide further insight into pathogenic mechanisms of EPEC and the contribution of host cell cytotoxicity to infection by the A/E pathogens. Since several effectors that influence cytotoxicity are required for full virulence in vivo, this phenotype warrants further investigation.  6.3 Subversion of innate immune signaling by bacterial pathogens: a recurring theme Host organisms have evolved complex strategies by which to combat bacterial pathogens. Following bacterial breach of physical barriers in the host, the first line of defense is the innate immune system, which recognizes PAMPs and mounts an initial pro-inflammatory response and primes the adaptive immune system.  155  Many bacterial pathogens have  evolved elegant mechanisms by which to subvert the innate immune system and promote their own propagation. In addition to intricate regulation of host cell death, A/E pathogens and Yersinia spp. actively inhibit innate immune signaling. Work presented in this thesis has demonstrated that the A/E effector NleC abrogates NF-κB activity and subsequently IL-8 secretion, in part through degradation of the HAT p300. NleC also degrades p65 and may have many other targets in host cells (Yen et al., 2010, Pearson et al., 2011, Muehlen et al., 2011, Baruch et al., 2011). Yersinia spp. are also extracellular pathogens that impair innate immune signaling. Whereas A/E pathogens appear to use several effector proteins to inhibit NF-κB activity and IL-8 secretion, Yersinia spp. utilize complimentary mechanisms to limit pro-inflammatory signaling. Yersinia YopP/J is a de-ubiquitinase and acetyl-transferase that inhibits phosphorylation of IKKβ and degradation of IκBα, thus maintaining NF-κB dimers in the nucleus. In addition, the translocators LcrV and YopB contribute to reducing of pro-inflammatory signaling during Yersinia infection (Sodhi et al., 2005).  LcrV inhibits production of TNF-α and INFγ and promotes  production of IL-10, an anti-inflammatory cytokine (Brubaker, 2003). Together with LcrV, YopB contributes to decreasing TNF-α production and macrophage activation (Sodhi et al., 2005). Another parallel between Yersinia and A/E pathogens is the ability to interfere with MAPK signaling in order to subvert the innate immune system. EPEC NleC actively impairs activation of the p38 MAPK and YopP/J inhibits activation of multiple host MAPKs (Sham et al., 2011, Matsumoto et al., 2009). Yersinia infection of macrophages also facilitates apoptotic cell death through the activity of YopP/J as mentioned above (Zheng et al., 2011).  The A/E effectors NleH1 and NleH2 also  156  modulate host cell apoptosis and innate immune signaling, although NleH1/2 are antiapoptotic (Royan et al., 2010, Hemrajani et al., 2010). Yersinia spp. and A/E pathogens have both evolved to suppress innate immune signaling; however, several bacterial pathogens subvert the innate immune system to exacerbate inflammation. In contrast to Yersinia and A/E pathogens, Shigella hijacks the host innate immune system by increasing inflammation. Tissue destruction and increased recruitment of proinflammatory cells characterize the initial immune response to S. flexneri. Interestingly, Shigella can both up- and down- regulate the innate immune system, but appears to ultimately use tissue destruction and severe diarrhea as a mechanism for transmission to naïve hosts (Phalipon et al., 2007). Shigella initially invade professional phagocytes and promote caspase-1-dependent cell death termed pyroptosis (Ashida et al., 2009). Subsequently, Shigella is able to invade IECs via their basolateral membranes and triggers increased IL-8 production (Pedron et al., 2003). The increased IL-8 secretion functions to attract neutrophils to the sites of infection, which likely mediate the extreme inflammation observed during Shigella infection (Phalipon et al., 2007). In stark contrast to the A/E pathogens, Shigella induction of NF-κB activity in IECs is not counteracted by translocated effector proteins but instead, the bacteria promote host NOD1 activation resulting in enhanced NF-κB activity (Girardin et al., 2001).  Since prolonged  inflammation is likely detrimental to the pathogen, Shigella utilizes an arsenal of T3S effector proteins to usurp the inflammatory response during infection (Ashida et al., 2009). The effector OspF translocates to host cell nuclei where it inhibits multiple MAPKs resulting in inhibition of histone H3 phosphorylation at Ser10 (Arbibe et al., 2007).  H3Ser10 phosphorylation enhances transcription of genes containing κB-  157  enhancer elements, thus increasing NF-κB-mediated gene expression (Wolter et al., 2008). Although we have shown that NleC-mediated degradation of p300 reduces NFκB activity and IL-8 secretion (Shames et al., 2011a), the global effect of NleC on host gene transcription has not been examined. To further regulate inflammation during Shigella infection, the effector OspG interacts with ubiquitinated E2 ubiquitinconjugating enzymes and inhibits transfer of ubiquitin moieties to phosphorylated IκBα thus using a recurring mechanism of inhibiting NF-κB liberation from IκBα (Kim et al., 2005). It is becoming increasingly apparent that intricate regulation of the host innate immune system is an important virulence strategy of bacterial pathogens. For A/E pathogens, innate immune signaling is temporarily inhibited until eventual pathogen clearance in non-susceptible hosts. Studies with C. rodentium in resistant mice have demonstrated a role for the adaptive immune system and a robust immune response for pathogen clearance (Higgins et al., 1999, Lebeis et al., 2007, Vallance et al., 2002, Simmons et al., 2002). Decreasing of the innate immune system, in addition to host cell cytotoxicity (see section 6.1), may increase time for the pathogen to replicate. Support for this hypothesis comes from the observation that infection with a C. rodentium quorum sensing mutant is ‘hyper-virulent’ and decreases survival of susceptible hosts at early time points (Coulthurst et al., 2007). Although the bacterial load of the quorum sensing mutant was not quantified, premature mortality of the host would limit pathogen dissemination. When the pathogens reach a high enough level of colonization, the signals to subvert the innate immune system may be overwhelmed thus resulting in pathogen clearance and potentially increased shedding. Alternatively, increased quorum sensing in C. rodentium  158  may facilitate inhibition of immune and cytotoxicity repression mechanisms. Ultimately, subversion of the host immune system is an important pathogenic mechanism employed by many bacterial pathogens and its investigation is continuously providing insight into techniques evolved by pathogens to facilitate their survival in the host environment.  6.4 Uncovering roles for bacterial effector proteins Our understanding of the pathogenic mechanisms of virulent bacteria is constantly increasing. It is apparent that many gastrointestinal pathogens encode secretion systems by which to inject bacterial proteins directly into infected host cells. These proteins have evolved to subvert host responses to infection and therefore promote bacterial colonization.  However, the functions of many of these effectors have yet to be  elucidated. A majority of gastrointestinal pathogens must invade host cells to cause disease, but the A/E pathogens are primarily extracellular and thus depend on their arsenal of injected proteins to inhibit their clearance by the host. This thesis has outlined novel functions for two A/E effector proteins, EspZ and NleC. Inhibition of host cell death by EspZ is likely critical for successful infection in vivo; however, this hypothesis has yet to be tested. NleC plays a role in inhibition of host NF-κB signaling IL-8 secretion together with several other effector proteins. The number of effector proteins dedicated to this phenotype indicates that it is an important pathogenic mechanism. In this case, targeting a host cell pathway would be a better mechanism to combat pathogenesis. Ultimately, the work presented in this thesis has added valuable insights into the function of A/E pathogen effector proteins during disease and has further  159  demonstrated that effector proteins can have multiple functions during bacterial infections.  This work has provided a basis for future directions to increase the  understanding of EspZ and NleC during A/E pathogen infection. In addition, much insight has been gained by these studies into the contribution of EspZ and NleC to the overall pathogenic mechanism of A/E pathogens. Studies presented in this thesis have greatly contributed to the field of T3S effector biology. Prior to this work, the function of EspZ was entirely unknown despite its ability to impair virulence and colonization of C. rodentium in vivo (Deng et al., 2004). Using cell culture models of infections, we were able to identify two host proteins targeted by EspZ and a molecular mechanism for decreasing cytotoxicity during EPEC infection. Further investigation of EspZ function during EPEC infection could follow several directions. The ability of EspZ to influence amino acid transport by CD98 heterodimers has yet to be examined and could provide insight into mechanisms of diarrhea production during EPEC infection.  In addition, a more detailed mechanism of EspZ-mediated  protection of host cells at mitochondria should be examined. TIM17B has not been associated with cytotoxicity prior to the work presented in Chapter 3 and further investigation may reveal a novel role for protein import complexes in eukaryotic cell death. Future work on this project could also focus on the ability of EspZ to impact transport of cytotoxic effector proteins EspF and Map into mitochondria in the presence or absence of TIM17B. The severity of cell death induced in host cells by the EPEC ∆espZ strain has provoked speculation further into the pathogenic mechanism of EPEC and why EspZ is crucial for virulence. The severe virulence “attenuation” observed in vivo following infection with C. rodentium ∆espZ may actually result from “hyper-  160  virulence” of the pathogen, as observed for EPEC in cell culture (Shames et al., 2011b, Shames et al., 2010). Infected IECs may undergo very rapid cell death at the onset of infection with a ∆espZ strain, resulting in the previously observed attenuation of virulence (Deng et al., 2004) and it would be very interesting to discern if this is responsible for impaired colonization of the ∆espZ strain.  Use of a Cre-Lox gene  excision system in C. rodentium whereby the espZ gene could be removed following specific induction would enable examination of cell death following initial colonization. However, investigation at early time points would be more relevant to the role of EspZ during infection based on its early translocation by EPEC (Mills et al., 2008). Host cell death is likely crucial for dissemination of A/E pathogens based on their intimate attachment to host cells; however, premature cell death would prevent optimal levels of colonization. Further studies on EspZ will likely provide very important insights into A/E pathogen virulence and many of these studies will be based from the first known function of EspZ during infection, which is presented in this thesis. The studies presented on the function of NleC revealed a novel mechanism used bacterial pathogens to lessen secretion of IL-8 during infection and identified NleC as the first bacterial virulence factor to target the HAT p300. Since the characterization of NleC as a Zn-metalloprotease occurred following our SILAC screen for interacting partners, the experiment could be modified to identify even more substrates of NleC. Use of the E184A catalytic mutant as bait in SILAC experiments would enable identification of targets that are cleaved at a greater rate than p300/CBP. I believe NleC likely has many more targets that have yet to be identified and may reside in different regions of the host cell. In addition, the homology between NleC and AIP56 from P. damselae suggests that  161  NleC may have pro-apoptotic functions. If NleC contributes to host cell apoptosis, the role of known substrates such as p65 and p300 should be examined. The influence of NleC on p300 may not be limited to proteolysis. Use of WT NleC to identify targets by SILAC enabled identification of proteins that can interact for prolonged periods. Since the NleC target p65 was not identified in our screen, the interaction and subsequent degradation of p65 is likely much more rapid. Further investigation into the kinetics of the metalloprotease activity of NleC may shed light on its function and differential substrate degradation. I believe further work on NleC will provide insight into a new class of T3S effector proteins using protease activity to regulate host cell processes. 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Summary  of  proteins  identified  by  MS  as  specifically  immunoprecipitated with HA2EspZ in SILAC experiments Protein Name HA-SepZ  Averagea 0.17  S.D.b 0.03  ubiquitin and ribosomal protein S27a precursor  0.18  0.04  4F2 cell-surface antigen heavy chain Heat shock protein HSP 90-beta  0.25 0.29  0.19 0.05  Heat shock protein HSP 90-alpha 2 Actin, alpha skeletal muscle  0.29 0.46  0.04 0.06  Sodium/potassium-transporting ATPase subunit beta-3 Tubulin alpha-3 chain Actin, cytoplasmic 1  0.46 0.47 0.49  0.15 0.13 0.05  Isoform 1 of Heat shock cognate 71 kDa protein Lung cancer oncogene 7 Heat shock 70 kDa protein 1 ADP/ATP translocase 2 60S acidic ribosomal protein P0  0.50 0.54 0.60 0.60 0.61  0.16 0.34 0.10 0.18 0.14  L-lactate dehydrogenase B chain Elongation factor 1-alpha 1 Tubulin beta-2 chain Alpha-enolase, lung specific 40S ribosomal protein SA enolase 1  0.62 0.63 0.64 0.64 0.69 0.69  0.03 0.22 0.18 0.11 0.15 0.16  Isoform Long of Sodium/potassium-transporting ATPase 0.72 alpha-1 chain precursor Tubulin beta-2C chain 0.74  0.16 0.13  Glyceraldehyde-3-phosphate dehydrogenase  0.76  0.14  Isoform A of Phosphate carrier protein, mitochondrial precursor 0.76 40S ribosomal protein S3 0.80 Protein DJ-1 0.81  0.02 0.22 0.02  184  Averagea  Protein Name  S.D.b  Isoform 1 of Sodium/potassium-transporting ATPase subunit beta-1 0.82 40S ribosomal protein S16 0.86  0.23 0.13  Nucleoside diphosphate kinase A 40S ribosomal protein S25  0.86 0.90  0.31 0.23  Malate dehydrogenase, mitochondrial precursor Peroxiredoxin-1 40S ribosomal protein S18  0.91 0.92 0.94  0.11 0.12 0.12  Hypoxanthine-guanine phosphoribosyltransferase peptidylprolyl isomerase A-like Thioredoxin 60S acidic ribosomal protein P2  0.95 0.95 0.95 0.98  0.19 0.13 0.05 0.39  Stress-70 protein, mitochondrial precursor  0.98  0.17  F-actin capping protein alpha-1 subunit  1.00  0.06  1-phosphatidylinositol-4,5-bisphosphate phosphodiesterase 1.05 beta 3 Crk-like protein 1.06 Peroxiredoxin-2 1.06 Isoform 2 of Nucleophosmin 1.07  0.25 0.34 0.13 0.13  Isoform 3 of Tyrosine-protein phosphatase non-receptor type 13 1.10  0.06  Isoform 2 of Protein-L-isoaspartate(D-aspartate) methyltransferase  1.11  0.02  1.11  0.07  Leucine zipper-EF-hand-containing transmembrane protein 1.13 1, mitochondrial precursor  0.11  F-actin capping protein alpha-2 subunit  1.16  0.33  Isoform Short of RNA-binding protein FUS  1.21  0.28  Eukaryotic translation initiation factor 3 subunit 1 Nucleolin  1.21 1.27  0.16 0.16  Macrophage migration inhibitory factor  185  O-  Protein Name Succinate dehydrogenase [ubiquinone] flavoprotein subunit, mitochondrial precursor Heat shock protein 60 Histone H2A type 1-B Histone H4 a – Averaged SILAC ratio (heavy isotope:light isotope).  Averagea  S.D.b  1.39 1.52 2.27 3.17  0.10 0.14 0.36 0.16  b – S.D.:standard deviation as determined by quantification of several peptides from either 2 or 3 independent immunoprecipitation experiments.  186  Appendix B. Peptides of E1A-associated protein p300 identified in NleC SILAC experiment Peptide K.HWEFSSLR.R K.RLQEWYK.K K.IFKPEELR.Q K.QLSELLR.G R.DAAYYSYQNR.Y K.NPMDLSTIK.R R.QDPESLPFR.Q R.LQQAQLMR.R K.SNPQLMAAFIK.Q R.QDPESLPFRQPVDPQLLGIPDYFDIVK.N R.DAFLTLAR.D M.AENLLDGPPNPK.R R.QALMPTLEALYR.Q K.LVQAIFPTPDPAALK.D R.MENLVAYAK.K K.QSMVNSLPTFPTDIK.N K.WGLGLDDEGSSQGEPQSK.S R.QPVDPQLLGIPDYFDIVK.N K.QAASTSGPTPAASQALNPQAQK.Q K.SPLSQGDSSAPSLPK.Q R.FVDSGEMSESFPYR.T R.SEMMEEDLQGASQVK.E R.SALSSELSLVGDTTGDTLEK.F  187  IonsScore mass error (Da) 27 0.0014 29 0.0009 31 0.001 33 0.0002 36 0.0016 37 0.0014 41 0.0013 42 0.0004 43 0.0022 45 0.0113 46 0.0009 48 0.0012 50 0.0026 50 -0.0011 56 0.0011 61 -0.0008 62 0.0016 62 0.005 67 -0.0018 72 0.0008 81 -0.0008 96 0.003 109 -0.0021  

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