LYSINE FATTY ACYLATION IS A DYNAMICALLY REGULATED MODIFICATION THAT AFFECTS BACTERIAL PATHOGENESIS A Dissertation Presented to the Faculty of the Graduate School of Cornell University In Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy By Garrison Paul Komaniecki August, 2022 © Garrison Paul Komaniecki LYSINE FATTY ACYLATION IS A DYNAMICALLY REGULATED MODIFICATION THAT AFFECTS BACTERIAL PATHOGENESIS Garrison Paul Komaniecki Cornell University, 2022 Protein post-translation modifications expand the chemical, molecular, and physiological diversity encoded by the genetic code. Types of protein modifications are numerous and diverse to suit wide functional roles. Lysine fatty acylation (KFA) is the covalent addition of long-chain fatty acyl groups to the lysine side chain amine. Despite being discovered over 30 years ago, relatively little is known about KFA. Recently, several mammalian enzymes were found to be able to efficiently remove KFA from proteins and multiple bacterial pathogens were discovered to mediate pathogenicity in part by catalyzing the addition of KFA. This thesis discusses findings that reveal KFA as a newly appreciated battleground during bacterial infection. V. cholerae, the causative agent of the disease cholera, secretes MARTX, a multi-domain toxin protein that contains a Rho-GTPase inactivating domain (RID). RID catalyzes KFA addition to RhoA family GTPases. In chapter II, we found that HDAC11 can hydrolyze lysine fatty acylation on RID to decrease its activity towards mammalian substrates. Bone marrow derived macrophages that lack HDAC11 phagocytose less V. cholerae. Correspondingly, mice lacking HDAC11 are defective in being able to clear a V. cholerae infection. In chapter III, we find that SIRT2 can promote defense of S. flexneri through KFA hydrolysis. IcsB is a KFA transferase from S. flexneri that promotes bacterial survival by inhibiting host autophagic machinery. S. flexneri also secretes IpaJ which causes Golgi stress. Golgi stress activates the transcription factor CREB3 to upregulate SIRT2. SIRT2 then hydrolyzes IcsB-catalyzed KFA to promote bacterial elimination. Mice that lack SIRT2 are more susceptible to S. flexneri infection. Then, in chapter IV, we also identify three novel KFA transferase toxins from L. pnuemophila, the pathogen that causes a pneumonia known as Legionnaire’s disease. We go on to identify the substrates of the most active toxin, lpg1387, and establish that the endogenous lpg1387 modulates KFA during L. pnuemophila infection. BIOGRAPHICAL SKETCH Garrison Komaniecki, better known as Sonny, was born in Rochester, MN. He attended the University of Minnesota – Morris where he obtained an ACS-certified BA in chemistry with an emphasis on biochemistry. He graduated with honors and received several awards for academic achievement. He joined the Biochemistry, Molecular, and Cell Biology graduate program in the Department of Molecular Biology and Genetics at Cornell University as a graduate student in 2016. In 2017, he began his research on protein palmitoylation in the lab of Professor Hening Lin. Dedicated to my girls. Aspire to be inspired. ACKNOWLEDGEMENTS I would first like to thank my Ph.D. advisor, Hening Lin. Professor Lin has fostered a great environment for students to learn and find their own pathways to success. In this environment, I developed from a naïve student into an expert in my field with countless new skills in scientific exploration and communication. Many of these skills, such as logical thinking and problem solving undeterred by setbacks, I attribute directly to his guidance and tutelage. When I got ready to graduate and take my next step toward my dream job of being a professor at a primarily undergraduate university, I initially intended to pursue a post-doc position. Professor Lin thought that I had the potential and acumen to skip this step and move directly to a tenure- track position. Now, having accepted one of these positions, I truly appreciate how valuable it is to have an advisor who has even more confidence in my abilities than I do. For that I will always be grateful. I would like to thank the other members of my special committee Professor Maurine Linder and Professor Jeremy Baskin. They provided me with valuable insight and advice for both my research and professional progression over the years. I also want to thank Professor Gerald Feigenson for serving on my A-exam committee and providing mentorship during my first year at Cornell. I would like to thank the many people that I collaborated with over the years. Dr. Miao Wang and Dr. Yugang Zhang made crucial contributions concerning SIRT2 and Shigella defense. Dr. Ji Cao initially discovered HDAC11’s activity in hydrolyzing lysine fatty acylation and made important contributions in understanding HDAC11’s role in defense against V. cholerae. Professor Edward Seto worked with us on HDAC11 projects and provided the HDAC11 knock out mice for our V. cholerae studies. Professor Tobias Doerr and his graduate student Anna Weaver at Cornell provided us with all the V. cholerae strains and Professor Neal Alto from UTSW provided us the S. flexneri strains. Professor Yuxin Mao and his graduate student Wenjie Zeng at Cornell provided a great collaboration for studying L. pnuemophila toxins. I also want to acknowledge Professor Devanand Sarkar and his postdoc Dr. Rajesh Yetirajam from UVA as great collaborators on a project not discussed in this dissertation and Professor Masaki Fukata from NIPS for providing many useful plasmid constructs. I would like to thank all the other people that served as mentors to me during my academic career. Dr. Ji Cao provided much of my initial training in the lab and every other lab member at some point served as a valuable source of advice and inspiration during my graduate career. I also want to thank my undergraduate professors for their guidance, especially my advisor Professor Nancy Carpenter. Nancy has been a role model for how to teach material effectively and for the type of educator I aspire to be. I would like to thank all the students and trainees I have worked with in the lab and in the classroom for being unwitting lab rats as I experiment with teaching techniques and for being outstanding coworkers. I especially want to thank two undergrads that I worked with extensively, Laurynn Cooper-Jordan and Bryan Arpi, for being great learners and a joy to work with. Lastly, I would like to thank my family. My parents have been endlessly supportive of all aspects of my life from watching me perform in the high school musicals to spending long weekends at wrestling tournaments to helping me move across the country to start my academic career. They provided me with all the opportunities I could ever need and fostered the curiosity that would ultimately lead to my admission into the academy. Thank you to my wife, Katie, who sacrificed her own career as I pursued mine and moved 1,000 miles away from her family to do so. Katie has been my rock during hard times and has reminded me to be more than just a scientist – to always remember the bigger picture. Thank you to my daughters, Luella and Ezra, for being the bigger picture and for revealing to me a depth of love I never knew existed. TABLE OF CONTENTS ABSTRACT ...................................................................................... Error! Bookmark not defined. BIOGRAPHICAL SKETCH ....................................................................................................... 5 ACKNOWLEDGEMENTS ......................................................................................................... 7 TABLE OF CONTENTS ........................................................................................................... 10 LIST OF FIGURES .................................................................................................................... 14 LIST OF TABLES ...................................................................................................................... 22 CHAPTER 1 ................................................................................................................................ 23 LYSINE FATTY ACYLATION: REGULATORY ENZYMES, RESEARCH TOOLS, AND BIOLOGICAL FUNCTION ............................................................................................ 23 ABSTRACT .............................................................................................................................. 23 INTRODUCTION ..................................................................................................................... 24 FATTY ACYL HYDROLASES ............................................................................................... 25 SIRT6..................................................................................................................................... 26 SIRT2..................................................................................................................................... 29 HDAC11 ................................................................................................................................ 30 LYSINE FATTY ACYL TRANSFERASES ............................................................................ 32 RtxC proteins ......................................................................................................................... 33 RID ........................................................................................................................................ 35 IcsB ........................................................................................................................................ 36 NMT1 and NMT2 .................................................................................................................. 38 FUNCTIONS OF LYSINE FATTY ACYLATION ................................................................. 40 IL-1α ...................................................................................................................................... 40 Aquaporin-0 ........................................................................................................................... 42 TNF-α .................................................................................................................................... 44 R-Ras2 ................................................................................................................................... 45 K-Ras4a ................................................................................................................................. 45 RalB ....................................................................................................................................... 46 Arf6 ........................................................................................................................................ 47 SHMT2 .................................................................................................................................. 47 Gravin-α ................................................................................................................................. 48 RTX Toxins ........................................................................................................................... 49 TOOLS FOR STUDYING LYSINE FATTY ACYLATION .................................................. 50 Methods to detect endogenous KFA ..................................................................................... 50 Enzyme activity assays .......................................................................................................... 51 Measuring KFA with fatty acyl probes ................................................................................. 55 SUMMARY AND FUTURE QUESTIONS ............................................................................. 57 REFERENCES .......................................................................................................................... 58 CHAPTER 2 ................................................................................................................................ 70 HDAC11 COUNTERACTS VIBRIO CHOLERAE MARTX TOXIN BY HYDROLYZING LYSINE FATTY ACYLATION ................................................................................................ 70 ABSTRACT .............................................................................................................................. 70 INTRODUCTION ..................................................................................................................... 70 RESULTS.................................................................................................................................. 73 RID fatty acylation is important for activity ......................................................................... 73 HDAC11 hydrolyzes RID lysine fatty acylation ................................................................... 75 RID lysine fatty acylation regulates its cell rounding activity .............................................. 77 RID lysine fatty acylation is important for activity in V. cholerae MARTX ........................ 79 HDAC11 KO mice are more susceptible to V. cholera infection ......................................... 80 DISCUSSION ........................................................................................................................... 83 METHODS................................................................................................................................ 87 Reagents................................................................................................................................. 87 Cloning and mutagenesis ....................................................................................................... 88 Cell culture ............................................................................................................................ 88 Detection of fatty acylation by in-gel fluorescence ............................................................... 88 Expression and purification of human HDAC11 from HEK 293T cells ............................... 89 In vitro defatty acylation assay .............................................................................................. 90 Generation of mutant V. cholerae strains .............................................................................. 91 Detection of V. cholerae induced fatty acylation .................................................................. 92 Mouse inoculation by V. cholerae ......................................................................................... 93 Determination of phagocytic V. cholerae uptake by BMDMs .............................................. 93 ACKNOWLEDGEMENTS ...................................................................................................... 94 REFERENCES .......................................................................................................................... 94 CHAPTER 3 ................................................................................................................................ 99 GOLGI STRESS INDUCES SIRT2 TO COUNTERACT SHIGELLA INFECTION VIA DEFATTY ACYLATION .......................................................................................................... 99 ABSTRACT .............................................................................................................................. 99 INTRODUCTION ................................................................................................................... 100 RESULTS................................................................................................................................ 101 Golgi stress upregulates SIRT2 via CREB3 ........................................................................ 101 SIRT2 counteracts the action of IcsB. ................................................................................. 108 SIRT2 limits Shigella autophagosome escape..................................................................... 111 SIRT2 restricts Shigella infection in cells and in mice. ...................................................... 111 DISCUSSION ......................................................................................................................... 117 METHODS.............................................................................................................................. 119 Reagents, antibodies and plasmids. ..................................................................................... 119 Cell culture, transfection and transduction. ......................................................................... 120 Bacterial strains and infection. ............................................................................................ 121 Gentamycin killing assay..................................................................................................... 121 Western blot analysis. .......................................................................................................... 122 RT-PCR analysis of mRNA levels. ..................................................................................... 122 Luciferase assay for Sirt2 promoter activity........................................................................ 122 Chromatin Immunoprecipitation. ........................................................................................ 123 Detection of fatty acylation on protein of interest using Alk14. ......................................... 124 Defatty acylation assay by sirtuins in vitro. ........................................................................ 124 GDI pulldown. ..................................................................................................................... 124 Immunofluorescence. .......................................................................................................... 124 Intraperitoneal shigellosis mice model study. ..................................................................... 125 Intranasal shigellosis mice model study. ............................................................................. 125 ACKNOWLEDGEMENTS .................................................................................................... 126 REFERENCES ........................................................................................................................ 126 CHAPTER 4 .............................................................................................................................. 131 LYSINE FATTY ACYLATION CATALYZED BY A FAMILY OF LEGIONELLA PNUEMOPHILA TOXINS ...................................................................................................... 131 ABSTRACT ............................................................................................................................ 131 INTRODUCTION ................................................................................................................... 132 RESULTS................................................................................................................................ 134 DISCUSSION ......................................................................................................................... 143 METHODS.............................................................................................................................. 146 Detection of individual protein fatty acylation by in-gel fluorescence. .............................. 146 Detection of global protein fatty acylation by in-gel fluorescence ..................................... 147 Detection of endogenous protein fatty acylation by biotin click chemistry ........................ 148 SILAC labeling for lpg1387 lysine fatty acylation substrates ............................................. 149 Protein Identification by nano LC/MS/MS Analysis: ......................................................... 150 MS data analysis: ................................................................................................................. 151 Generation of lpg1387 knockout ......................................................................................... 152 Infection of HEK 293T cells by L. pnuemophila ................................................................ 154 ACKNOWLEDGEMENTS .................................................................................................... 155 REFERENCES ........................................................................................................................ 155 CHAPTER 5 .............................................................................................................................. 158 SUMMARY AND FUTURE DIRECTIONS .......................................................................... 158 SUMMARY ............................................................................................................................ 158 FUTURE DIRECTIONS......................................................................................................... 159 Further characterization of V. cholerae and L. pnuemophila KFAT toxins ........................ 159 Further exploration of HDAC11’s role during bacterial infection ...................................... 162 Discovery of additional KFAT toxins and characterization of Burkholderia BopA ........... 162 REFERENCES ........................................................................................................................ 163 LIST OF FIGURES Figure 1. 1. KFA. KFA is the modification of lysine side chains with long fatty acyl groups. How KFA affects a modified substrate is not always understood but known outcomes of KFA are diverse. ............................................................................................................... 25 Figure 1. 2. Structure of KFA hydrolases SIRT2 (A,B, PDB 4R8M) and SIRT6 (C,D, PDB 3ZG6) highlighting hydrophobic pockets that accommodate long-chain fatty groups (B,D). (A) Overall structure of SIRT2 in complex with a thiomyristoyl-lysine peptide. The peptide is shown in green. The blue oval highlight the overall active site of SIRT2 and the catalytic histidine residue is shown in stick representation. (B) Zoom-in view of the SIRT2 hydrophobic pocket that accommodates the myristoyl group. The hydrophobic side chains are shown in cyan stick representations. A surface representation showing the hydrophobic pocket in a slight different orientation is also shown. (C) Overall structure of SIRT6 in complex with a myristoyl-lysine peptide. The peptide is shown in green. The blue oval highlight the overall active site of SIRT2 and the catalytic histidine residue is shown in stick representation. (D) Zoom-in view of the SIRT2 hydrophobic pocket that accommodates the myristoyl group. The hydrophobic side chains are shown in cyan stick representations. A surface representation showing the hydrophobic pocket in a different orientation is also shown. .................................................................................................. 27 Figure 1. 3. Bacterial KFA transferases. (A) The genetic structure for a generic RTX toxin operon includes five genes. The RtxC toxin is modified with KFA by RtxA and secreted through a transmembrane spanning pore consisting of RtxB, RtxD, and TolC. In the extracellular space, RtxC binds Ca2+ with characteristic nonapeptide repeats and associates with mammalian cell membranes. Oligomerization of RtxC forms large lytic pores in the membrane. (B) The structure of the Vibrio cholerae MARTX toxin is illustrated. Interior domains vary from species to species, but the terminal RTX domains in a MARTX toxin allows for translocation through the cell membrane. Once in the cell the cysteine protease domain (CPD) is activated to cleave the toxin at several points releasing the other domains: the actin crosslinking domain (ACD), the α/β hydrolase domain (ABH), and the Rho-inactivating domain (RID). RID modifies RhoA-family GTPases with KFA to suppress multiple cellular processes. (C) During part of the pathogenesis of Shigella flexneri, bacteria reside in an intracellular vacuole that can be destroyed through autophagy involving CHMP5. IcsB blocks autophagic destruction by catalyzing KFA on CHMP5. Figure created with BioRender.com. ................................. 32 Figure 1. 4. Model for NMT KFA transferase activity. Rotation around the peptide backbone (bold) positions the amine of a lysine side chain similarly to the amine of an N-terminal glycine. Both are primary amines that can rotate freely during NMT catalysis. The superimposed myristoyl-lysine (green stick representation) and myristoyl-glycine (white stick representation) in human NMT2 (PDB 6PAU) and NMT1 (PDB 5O9V) structures overlap nicely, supporting this model. .............................................................................. 39 Figure 1. 5. Models of KFA regulation. (A) KFA on TNF-α directs it to the lysosome for degradation. SIRT6 hydrolyzes TNF-α KFA leading to higher levels on the plasma membrane and increased secretion of cleaved, soluble TNF-α. (B) KFA on the polybasic region of R-Ras2 results in increased plasma membrane localization, PI3K interaction, AKT signaling, and cell proliferation. SIRT6 hydrolyzes R-Ras2 KFA, releasing it from the membrane and decreasing cell proliferation. (C) K-Ras4a with KFA preferentially localizes at the plasma membrane. SIRT2 hydrolyzes K-Ras4a KFA. Cysteine palmitoylation on K-Ras4a retains membrane affinity, but instead localizes to endomembranes where K-Ras4a interacts with the signaling kinase A-Raf leading to downstream cellular transformation. (D) KFA on RalB also directs it to the plasma membrane to interact with Sec5 and Exo84 and promote cell migration. SIRT2 hydrolyzes RalB KFA, decreasing plasma membrane localization. (E) NMT-catalyzed KFA on K3 of Arf6 increases plasma membrane localization. GTP hydrolysis at the plasma membrane aided by a GTPase activating protein (GAP) causes Arf6 N-terminal glycine myristoylation to be sequestered. Arf6 is then trafficked to the endocytic recycling compartment (ERC) where SIRT2 hydrolyzes KFA and Arf6 is reloaded with GTP. NMT then starts the cycle again. Both the GTP and KFA dynamic cycle promote ERK signaling. Figure created with BioRender.com. ....................................................... 41 Figure 1. 6. Techniques for studying KFA enzyme activity. (A) [32P]NAD+ sirtuin assay. 32P (red atom) is incorporated into NAD+ to be used by a sirtuin with KFA hydrolase activity. During the hydrolysis mechanism, the sirtuin transfers the fatty acyl group from a modified protein to the cofactor releasing nicotinamide (NAM) and resulting in the formation of [32P]fatty acyl-ADPR. TLC plates are used to separate, visualize, and quantify radiolabeled species. (B) HPLC analysis of KFA enzyme activity on peptides. Unmodified and KFA-modified peptides are separated with HPLC and the area of the corresponding peaks are determined for quantification. (C) Fluorescence-based peptide KFA hydrolase assay. KFA is hydrolyzed from a lysine immediately preceding and AMC group. Trypsin hydrolyzes the amide bond following only the unmodified lysine releasing the AMC, which then exhibits fluorescence. (D) Acyl-cLIP assay for KFA enzymes. KFA on a peptide modified with fluorescein increases its affinity for micellar membranes. Fluorescein has slower tumbling near the bulky micelles, resulting in increased fluorescence polarization. Figure created with BioRender.com. ...................... 52 Figure 1. 7. Metabolic probes to examine KFA on a substrate protein. (A) Structure of KFA probes. Bioorthogonal probes for KFA can mimic myristic acid or palmitic acid. Both azide and alkyne probes are applicable though alkyne probes more closely resemble the endogenous fatty acids. (B) Metabolic labeling of KFA proteins can be analyzed with CuACC. Alkyne labeled proteins can be modified with fluorophores or affinity tags like biotin to analyze KFA levels with in-gel fluorescence, western blot, or mass spectrometry. ..................................................................................................................... 56 Figure 2. 1. RID has lysine fatty. A) HEK 293T cells were co-transfected with RID and its substrate Rac1. Fatty acylation was analyzed with Alk14 labeling followed by click chemistry. Rac1 shows fatty acylation in the presence of RID. RID shows fatty acylation. B) Rac1 was co-overexpressed with catalytically dead mutants of RID to analyze fatty acylation status. Both Rac1 and RID lack fatty acylation demonstrating that catalytic activity is necessary for RID modification as well as substrate modification. C) RID lysine point mutants were generated and assayed for fatty acylation. Mutation of K2816 to R abolishes RID fatty acylation. ................................................................................... 74 Figure 2. 2. RID lysine fatty acylation increases activity towards human substrate. A) Rac1 was co-overexpressed with WT and K2816R (KR) RID to assay activity of RID mutant. K2816R RID shows decreased activity compared to WT. B) Rac3 was co-overexpressed with WT, C3022A (CA), K2816R (KR), or K2816A (KA) RID. KR mutant RID showed decreased activity compared to WT whereas KA did not................................................. 75 Figure 2. 3. HDAC11 hydrolyzes RID lysine fatty acylation. A) HEK 293T cells were co- transfected with Rac1, RID, and HDAC11. Rac1 fatty acylation was assayed with Alk14 labeling and showed a decrease with the overexpression of WT HDAC11, but no change with catalytic dead HDAC11 (DA). B) Rac1 was purified from HEK 293T cells co- transfected with RID. Fatty acylated Rac1 was then treated with HDAC11 to assay in vitro activity towards Rac1. No change in Rac1 fatty acylation was detected with the addition of HDAC11. C) RID was co-transfected with HDAC11 in HEK 293T cells. RID fatty acylation was reduced in the presence of WT HDAC11 and unchanged in the presence of catalytic dead (DA) HDAC11. D) HDAC11’s activity towards fatty acylated RID was assayed in vitro by treating RID purified from transfected HEK 293T cells. WT, but not catalytic dead (YH), HDAC11 could decrease RID fatty acylation demonstrating that HDAC11 can hydrolyze RID lysine fatty acylation. E) Biochemical model of HDAC11 counteraction of RID. Graphic made with BioRender. .................................... 76 Figure 2. 4. Lysine fatty acylation regulates RID cell rounding activity. A) RID mutants were transfected into HEK 293T cells stably overexpressing GFP. WT RID caused a clear cell rounding phenotype where as C3022A and K2186R mutants did not. B) WT and catalytic dead (DA) HDAC11 were co-transfected into HEK 293T cells stably overexpressing GFP. WT HDAC11 was able to partially rescue RID-induced cell rounding whereas catalytic dead HDAC11 was not. ...................................................................................... 78 Figure 2. 5. Visualization of RID substrate fatty acylation from live V. cholerae. A) HEK 293T cells were transfected with RID substrate Rac3, incubated with Alk14, and infected with V. cholerae for the indicated times. V. cholerae infection caused an increase in Rac3 fatty acylation, demonstrating RID activity. NH2OH treatment showed no decrease in Rac3 fatty acylation ruling out cysteine fatty acylation. B) HEK 293T cells were transfected with the indicated GTPases, incubated with Alk14, and infected with V. cholerae for the indicated times. All GTPases show increased fatty acylation with V. cholerae infection, demonstrating RID activity................................................................ 79 Figure 2. 6. RID lysine fatty acylation affects activity in V. cholerae. A) Rac3 fatty acylation was assayed with RID point mutants V. cholerae strains. The RID catalytic dead C3022A (CA) strain showed no activity in Rac3 fatty acylation. The RID K2816R mutant (KR) showed decreased activity towards in Rac3 fatty acylation whereas the K2816A mutant was as active as WT. Actin crosslinking was used as a control to demonstrate equal MARTX translocation between mutants. B) Rac3 fatty acylation was assayed in HEK 293T cells stably overexpressing HA-HDAC11. Rac3 fatty acylation was decreased with HDAC11 overexpression. Actin crosslinking was used as a control for equal MARTX translocation between cell lines. C) Fatty acylation of RhoA-family GTPases was assayed for 6 hours in HEK 293T cells stably overexpressing HA-HDAC11. HDAC11 overexpression resulted in decreased fatty acylation for all RID substrates. ................... 81 Figure 2. 7. HDAC11 is beneficial during V. cholerae infection. A) Workflow of mouse V. cholerae infection protocol. Following 24 hour administration of streptomycin in the water source to eliminate mouse native gut microbiome, V. cholerae was administered via oral gavage. Mice were tracked for two weeks and fecal pellets isolated to count bacterial colonization. B) Average weight change of mice experimental groups over the course of two week infection. C) Quantification of bacteria isolated from FPs of WT and HDAC11 mice challenged with PBS or V. cholerae. D) Bacterial growth from serial dilutions of FP homogenates from WT and HDAC11 mice infected with V. cholerae. Values at left correspond to the fold dilution from FP homogenates. .............................. 83 Figure 2. 8. HDAC11 and RID affect V. cholerae phagocytosis by BMDMs. A) Representative fluorescence microscopy images of WT and HDAC11 BMDMs challenged with the indicated strains of V. cholerae expressing GFP. Nuclei are stained with DAPI. B) Quantification of A. At least 700 BMDMs were quantified for each sample. .............................................................................................................................. 84 Figure 2. 9. K2816 is near active site of V. vulnificus RID. The structure of the V. vulnificus RID domain has been solved. K2816 is near the putative active site residues C3022A (mutated to an alanine when solving the crystal structure) and H2782. .......................................... 86 Figure 3. 1. Golgi stress upregulates SIRT2. A) Immunoblots for SIRT2 in different cell lines treated with 5 g/ml BFA for 24 hrs. HSP90 blots are shown as loading controls. Representative images from three independent experiments are shown. B) In A549 cells, BFA treatment significantly increased SIRT2 protein level, while the ER stress inducers Tunicamycin (Tm) and Thapsigargin (Tg) only weakly induced SIRT2 expression. C) RT-PCR analysis for SIRT1-7 and HDAC11 mRNA level in A549 cells treated with 5 g/ml BFA for 24 hrs. Fold change in mRNA was calculated by comparing samples with BFA treatment to control DMSO. ................................................................................... 102 Figure 3. 2. CREB3 promotes SIRT2 transcription under Golgi stress. A) RT-PCR analysis for Sirt2 mRNA in A549 CREB3 KD cells treated with or without BFA. mRNA is normalized to DMSO treated control (shLuc) cells. Statistical evaluation was done by two-way ANOVA. (B) Immunoblots for SIRT2 protein levels in A549 control (shLuc) and CREB3 KD cells treated with or without BFA. HSP90 blot was used as the loading control. (C) Immunoblots for SIRT2 in control and CREB3 KD cells treated with BFA and rescued with CREB3 transfection. SIRT2 level relative to actin loading indicated below blots. ..................................................................................................................... 104 Figure 3. 3. CREB3 binds the Sirt2 promoter to regulate transcription. A) A schematic representation of SIRT2 promoter. The annotated regions are on Chromatin 19 complement[39,389,400-39,391,600], reference: GRCh37. B – C) SIRT2-promoter driven firefly transcription in cells under CREB3 N-terminal bZIP domain overexpression (B) or BFA treatment (C). The Renilla luciferase construct was used as an internal control. D) Firefly luciferase transcription in cells treated with BFA. Cells were transfected with an empty pGL3-basic vector or pGL3-basic vector with 994 base pairs of SIRT2 5’ UTR. E) Immunoprecipitation from cells treated with BFA using an α-CREB3 antibody. Blot shows full length and cleaved version of CREB3. F) qPCR of Sirt2 promoter from chromatin IP of cells treated with or without BFA. ............................... 105 Figure 3. 4. Shigella infection upregulates SIRT2 through Golgi stress. A) SIRT2 protein level is induced by IpaJ, showing by immunoblots for SIRT2 protein levels in HEK293T cells with or without Flag-IpaJ overexpression. HSP90 is the loading control. B) Immunoblots of SIRT2 in BMDMs infected with mock, wildtype or IpaJ deletion S. flexneri M90T from 6 to 8-week old wildtype C57BL/6J mice. C) Quantification of SIRT2 protein levels (normalized to actin) in B. Statistical evaluation was done by student t-test. D) Immunoblots of Sirt2 in bronchoalveolar lavage cells from 6 to 8-week old wildtype C57BL/6J mice intranasally infected with mock, wildtype or IpaJ deletion S. flexneri M90T. E) Quantification of SIRT2 protein levels (normalized to HSP90) in D. Statistical evaluation was done by one-way ANOVA. F) Immunoblots of HEK 293T cells transfected with empty vector or flag tagged EspG or VirA. ......................................... 107 Figure 3. 5. SIRT2 can remove IcsB-catalyzed lysine fatty acylation. A) In-gel fluorescence detection of lysine fatty acylation of Flag-tagged IcsB substrate proteins treated with 5 M of SIRT2, with or without 1 mM of NAD+ in vitro. B) In-gel fluorescence detection of lysine fatty acylation of Flag-tagged IcsB substrates in HEK293T cells that were also transfected with Flag-tagged IcsB and SIRT2. Representative images from at least 3 independent experiments are shown. EV, empty vector. WT, SIRT2 WT. HY, SIRT2 HY. FL, fluorescence (indicative of lysine fatty acylation level on substrate proteins). Flag, anti-Flag immunoblot (indicative of input level of substrate proteins). C) In-gel fluorescence detection of lysine fatty acylation of Flag-tagged CHMP5 in HEK293T cells that were co-transfected with Flag-tagged IcsB and SIRT2, empty vector (EV), SIRT6, SIRT7, or HDAC11. ....................................................................................................... 109 Figure 3. 6. SIRT2 -catalyzed defatty acylation promotes Rho GTPases and Rho GDI interaction. The disruption of Rho GTPase binding to RhoGDI by IcsB is rescued by SIRT2. HEK293T cells were transfected with plasmids encoding indicated proteins: IcsB, SIRT2 WT or HY mutant, and Flag-tagged Rac1 (A), Rac2 (B) or CDC42 (C). Cell lysates for each sample were subjected to GST-RhoGDI pulldown assay (shown in the first panel) and lysine fatty acylation labeling assay (shown in the last panel). The quantification of GDI enrichment of Rho GTPases, obtained by dividing Flag-Rho GTPase signal in GDI pulldown with Flag-Rho GTPase in the input (second panel), is shown at the bottom. ....................................................................................................... 110 Figure 3. 7. SIRT2 suppresses Shigella autophagosome escape. A) Effect of SIRT2 knockout on S. flexneri autophagosome escape. Representative images of MEF cells infected with S. flexneri from three biological replicates are shown. Scale bar: 15 m. White arrow: LC3-positive Shigella. B) Quantitation of data shown in A. Percentage of MEF cells containing LC3-positive shigella was quantified. Data is shown as mean ± SEM with >100 infected MEF cells counted for each experiment. Statistical evaluation was done using two-way ANOVA. Data are represented as mean ± SEM. *p<0.05. ns, not significant. C) Autophagy level (measured by ratio of LC3-II to LC3-I) is lower in SIRT2 knockout and knockdown cells upon S. flexneri infection. Immunoblotting analysis of LC3 in SIRT2 knockdown A549 cells infected with wildtype or IcsB deletion S. flexneri M90Tstrain at the same MOI. The quantification of LC3-II/LC3-I is shown at the bottom. ......................................................................................................................................... 112 Figure 3. 8. SIRT2 restricts Shigella infection in cells and in mice model. A) Effect of Sirt2 knockout on S. flexneri intracellular proliferation analyzed by gentamycin killing assay. MEF Sirt2+/+ and Sirt2-/- cells were infected with equal number of S. flexneri for 10 min. Intracellular S. flexneri number and MEF cell number were counted at indicated time points to get MOI value. Data are presented as mean ± SEM with three biological replicates, and each with two technical replicates. Statistical evaluation was done by an unpaired two-tailed Student’s t test. B) The protective effect of SIRT2 was suppressed when IpaJ is deleted. A549 control and SIRT2 KD cells were infected with equal number of wildtype and IpaJ deletion S. flexneri M90T cells for 10 min, then treated with gentamycin to kill extracellular bacteria. Intracellular S. flexneri number and mammalian cell number were counted to get MOI. Data are presented as mean ± SEM with nine biological replicates. Statistical evaluation was done using two-way ANOVA. C) Recoverable CFU in lung homogenates from 6 to 8-week old Sirt2+/+ and Sirt2-/- C57BL/6J mice intranasally infected with 1 million S. flexneri M90T wildtype strain. Statistical evaluation was done using unpaired two-tail Student’s t test. Data are presented as mean ± SEM with 3 mice per group. D) CREB3 KD cells transfected with WT or HY SIRT2 and infected with S. flexneri M90T for 10 mins before washing with PBS and replacing the media with 50 g/ml gentamicin for 6 hours. Cells were collected and CFU/cell determined. ............................................................................................... 114 Figure 3. 9. SIRT2 restricts Shigella infection in cell and in vivo by counteracting IcsB. (A) Effect of SIRT2 knockout and IcsB deletion on S. flexneri intracellular proliferation. MOI were determined after 10 min of infection and 6 hrs of incubation in the presence of 50 g/ml gentamicin. Data are represented as mean ± SEM with three biological replicates. Statistical evaluation was done by two-way ANOVA. ***p<0.001, ns, not significant. (B) CFU in lung homogenates from Sirt2+/+ and Sirt2-/- C57BL/6J mice infected with wildtype or IcsB deletion S. flexneri M90T for 3 days. Data are presented as mean ± SEM with 3 mice per group. Statistical evaluation was done using two-way ANOVA. *p<0.05, ns, not significant. (C) CFU in lung homogenates from Sirt2+/+ and Sirt2-/- C57BL/6J mice infected with wildtype or IcsB deletion S. flexneri M90T rescued with WT or C306A IcsB for 3 days. Data are presented as mean ± SEM with 3 mice per group. Statistical evaluation was done using two-way ANOVA. *p<0.05, ns, not significant. (D) Model depicting SIRT2 as a Golgi stress response protein that limits Shigella pathogenesis by counteracting Shigella-mediated host protein lysine fatty acylation…116 Figure 4. 1. Sequence alignment of L. pnuemophila toxins lpg1387, lpg0196, and lpg1797. Putative catalytic residues are colored in red. ................................................................. 133 Figure 4. 2. Structures of lpg0196, lpg1387, and 1797 predicted by Alphafold. Structures are overlapped to demonstrate structural similarities. The putative catalytic site and residues are shown below. ............................................................................................................ 134 Figure 4. 3. Fluorescence gels and immunoblots of Rac3 cotransfected with the indicated toxins to assay for fatty acylation using Alk14 labeling. HA = hydroxylamine. HA lpg1387 = H38A mutant. CA lpg1387 = C403A mutant. ................................................................ 137 Figure 4. 4. lpg1387 has KFA. A) In-gel fluorescence detection for lysine fatty acylation of indicated L. pneumophila proteins treated with or without hydroxylamine. Coomasie brilliant blue (CBB) stained gel was used to check protein loading. B) In-gel fluorescence detection of WT or catalytic dead mutants of lpg1387, H38A and C403A. C) In-gel fluorescence detection of WT or indicated lysine to arginine mutants of lpg1387 and with all four lysines mutated to arginine (4KR). D) In-gel fluorescence and immunoblots of lpg1387 co-transfected with indicated mammalian lysine fatty acyl hydrolase enzymes. E) In-gel fluorescence detection of flag-RheB lysine fatty acylation co-transfected with WT or 4KR lpg1387. HA = hydroxylamine. .................................................................. 138 Figure 4. 5. SILAC proteomic screen for lpg1387 substrates. A) Experimental setup for processing proteomic samples. B) Flow chart for selecting high confidence substrate hits from the SILAC screen. .................................................................................................. 139 Figure 4. 6. KFA of lpg1387 substrates. A) In-gel fluorescence and immunoblots of flag- tagged SILAC hits co-transfected with lpg1387. Coomasie brilliant blue (CBB) stained gel was used to check protein loading. B) In-gel fluorescence of WT and lysine to arginine mutant flag-RheB co-transfected with lpg1387. C) Diagram of RheB C-terminal sequence showing position of mutated lysines and terminal farnesylation. D) In-gel fluorescence and immunoblots of flag-RheB co-transfected with lpg1387 and indicated lysine fatty acyl hydrolases. E) In-gel fluorescence and immunoblots of flag-Rac3 co- transfected with lpg1387 and indicated lysine fatty acyl hydrolases. ............................. 141 Figure 4. 7. L. pnuemophila infection modulates KFA levels. A) In-gel fluorescence of whole cell lysates from HEK 293T cells incubated with Alk14 and infected with WT or lpg1387 KO (Δ) Lp02. Coomasie brilliant blue (CBB) stained gel was used to check protein loading. Red arrows indicate bands that increase in intensity with infection of both strains. Blue arrows indicate bands that increase in intensity only with infection by WT Lp02. B) In-gel fluorescence and immunoblots of flag-RheB transfected into HEK 293T cells infected with WT or lpg1387 KO (Δ) Lp02. C) In-gel fluorescence and immunoblots of WT or K178R flag-RheB transfected into HEK 293T cells infected with WT Lp02. “+HA” samples were incubated at 95 oC for five minutes with 0.66 M hydroxylamine. ............................................................................................................... 143 Figure 5. 1. Summary of KFA regulation reactions discussed in this work. ............................. 159 Figure 5. 2. A) In-gel fluorescence and immunoblots of CHMP5 co-transfected with RID. B) In- gel fluorescence of Rac3 in cells infected with V. cholerae. Panels at left correspond to Alk14-modified Rac3 reacted with TAMRA-N3 for in-gel fluorescence. Panels at right correspond to Alk14-modified Rac3 reacted with biotin-N3 then precipitated, resolubilized and purified with streptavidin. .................................................................. 160 Figure 5. 3. In-gel fluorescence and immunoblots of CHMP5 and IcsB co-transfected with HDAC11. ........................................................................................................................ 162 LIST OF TABLES Table 1. 1. Enzymes that regulate lysine fatty acylation .............................................................. 28 Table 1. 2. Proteins with lysine fatty acylation ............................................................................ 43 Table 4. 1. Top five hits from an HHpred structural search for lpg1387 from residues 152-355. The HDA1 coiled coil is formed from a single chain. The coiled coil structure in the other hits is formed from at least two chains. .......................................................................... 134 Table 4. 2. Proteins identified in SILAC proteomics screen for lpg1387 lysine fatty acylation substrates. Heavy to light ratio for both the forward are reverse experiment is shown along with a molecular detail for each protein. ............................................................... 140 Table 4. 3. Selected gene ontology biological process identified to be enriched in proteomic hits from SILAC screen for lpg1387 substrates. ................................................................... 139 CHAPTER 1 LYSINE FATTY ACYLATION: REGULATORY ENZYMES, RESEARCH TOOLS, AND BIOLOGICAL FUNCTION This is a revised version of the published paper: Komaniecki, G., and Lin, H. (2021) Lysine Fatty Acylation: Regulatory Enzymes, Research Tools, and Biological Function, Front Cell Dev Biol 9, 717503. ABSTRACT Post-translational acylation of lysine side chains is a common mechanism of protein regulation. Modification by long-chain fatty acyl groups is an understudied form of lysine acylation that has gained increasing attention recently due to the characterization of enzymes that catalyze the addition and removal this modification. In this review we summarize what has been learned about lysine fatty acylation in the approximately 30 years since its initial discovery. We report on what is known about the enzymes that regulate lysine fatty acylation and their physiological functions, including tumorigenesis and bacterial pathogenesis. We also cover the effect of lysine fatty acylation on reported substrates. Generally, lysine fatty acylation increases the affinity of proteins for specific cellular membranes, but the physiological outcome depends greatly on the molecular context. Finally, we will go over the experimental tools that have been used to study lysine fatty acylation. While much has been learned about lysine fatty acylation since its initial discovery, the full scope of its biological function has yet to be realized. INTRODUCTION Post-translational modification of proteins is a key biological regulatory mechanism that impacts every aspect of life. One class of post-translational modifications is protein lipidation which involves the attachment of hydrophobic moieties such as fatty acyl groups, isoprenoid lipids, or cholesterol 1. These hydrophobic species partition into the lipophilic environment of cellular membranes, bringing the modified protein to the membranes 2. N-terminal glycine myristoylation and cysteine palmitoylation and prenylation are well studied forms of protein lipidation known to regulate biological processes from development and cancer to inflammation and microbial pathogenesis 3. In comparison, fatty acylation of lysine residues is under- examined. However, recent progress in the understanding of the enzymes that regulate lysine fatty acylation and the effect of the modification on substrates has opened the door to exciting new research directions. Lysine fatty acylation (KFA) is the addition of long-chain fatty acyl groups to lysine side chains via amide bonds. Myristoylation (C14) and palmitoylation (C16) are the most common forms of KFA, but the identity of the endogenous acyl group is often unknown making fatty acylation a more inclusive and general description (Figure 1.1). KFA of mammalian proteins was first discovered in 1988 by Bursten et al. when studying the membrane affinity of IL-1α. Since this initial discovery, seven other human proteins and an entire class of bacterial proteins were identified to be modified by KFA. In addition, several proteins of both human and bacterial origin were found to be able to add or remove this modification. These discoveries, along with the tools that make them possible, will be covered in this chapter. Figure 1. 1. KFA. KFA is the modification of lysine side chains with long fatty acyl groups. How KFA affects a modified substrate is not always understood but known outcomes of KFA are diverse. FATTY ACYL HYDROLASES To date, all enzymes known to have KFA hydrolase activity fall into the lysine deacetylase (KDAC) family of proteins. This includes both Zn2+-dependent histone deacetylases (HDACs) and the NAD+-dependent sirtuins (SIRTs). Archaecterial sirtuins Sir2Af1 and Sir2Af2 from the archaea Archaeoglobus fulgidus, were also reported to be able to remove KFA, but there are no known substrates for this activity 4. The KFA hydrolases discussed below were originally assumed to only remove acetyl groups so it is possible that some of the bacterial deacetylases could also have KFA hydrolase activity 5. Sir2A from the malaria parasite Plasmodium falciparum has also been found to have KFA hydrolase activity, but again no substrates for this activity have been identified 6. Human enzymes HDAC8 and HDAC11 along with SIRT1, 2, 3, 6, and 7 can all remove KFA in vitro (Table 1.1). We will highlight the enzymes with known endogenous substrates for this activity in order of their discovery. SIRT6 The first mammalian protein identified to hydrolyze KFA is SIRT6 7. SIRT6 is involved in several physiological processes such as regulating immune signaling and suppressing tumorigenesis 8. SIRT6 is recruited to chromatin by DNA double strand breaks and by transcription factors such as Hif-1α to remodel chromatin and regulate gene expression 9. SIRT6 is best characterized as a lysine deacetylase. Targets of SIRT6 deacetylase activity include histone H3 and GCN5, which represses NF-kB levels and regulates glucose production, respectively 9b, 10. However, in vitro SIRT6 deacetylase activity is relatively weak compared to other sirtuin family members, raising the possibility of additional enzymatic activities 11. Jiang et al. explored alternative SIRT6 deacylation activities using various acyl-lysine peptides 7. Like previous studies, SIRT6 had very little deacetylation activity. However, SIRT6 was able to efficiently hydrolyze octanoyl, myristoyl, and palmitoyl lysine. A SIRT6 structure was obtained by co-crystallization with a myristoyl-lysine peptide revealing a hydrophobic groove in which bound the myristoyl group (Figure 1.2). Interestingly, free fatty acids were found to activate SIRT6 deacetylase activity but to inhibit deKFA activity 12. Together, these observations suggest that binding of fatty acyl groups, whether free or on a lysine, to the acyl pocket of SIRT6 may activate SIRT6. Figure 1. 2. Structure of KFA hydrolases SIRT2 (A,B, PDB 4R8M) and SIRT6 (C,D, PDB 3ZG6) highlighting hydrophobic pockets that accommodate long-chain fatty groups (B,D). (A) Overall structure of SIRT2 in complex with a thiomyristoyl-lysine peptide. The peptide is shown in green. The blue oval highlight the overall active site of SIRT2 and the catalytic histidine residue is shown in stick representation. (B) Zoom-in view of the SIRT2 hydrophobic pocket that accommodates the myristoyl group. The hydrophobic side chains are shown in cyan stick representations. A surface representation showing the hydrophobic pocket in a slight different orientation is also shown. (C) Overall structure of SIRT6 in complex with a myristoyl-lysine peptide. The peptide is shown in green. The blue oval highlight the overall active site of SIRT2 and the catalytic histidine residue is shown in stick representation. (D) Zoom-in view of the SIRT2 hydrophobic pocket that accommodates the myristoyl group. The hydrophobic side chains are shown in cyan stick representations. A surface representation showing the hydrophobic pocket in a different orientation is also shown. Table 1. 1. Enzymes that regulate lysine fatty acylation Name Species kcat/KM (s -1/M-1) Known References Substrates RtxC Many gram NA RtxA toxins 36 Family negative bacteria Lysine fatty RID V. cholerae NA RhoA-family 50 acyl GTPases transferases IcsB S. flexneri NA Several - see 53 citation. NMT1, Human NMT1: 144 a Arf6 16c, 64 NMT2 NMT2: 133 a HDAC8 Human 120 b NA 97 HDAC11 Human 1.54 x 104 b SHMT2 26a Lysine fatty SIRT1 Human 1.44 x 105 b NA 102 acyl SIRT2 Human 7.4 x 104 b K-Ras4a, RalB 16a, 16b hydrolases SIRT3 Human 2.51 x 105 b NA 102 SIRT6 Human 1.4 x 103 b TNF-α, R-Ras2 7, 80 SIRT7 Human > 167 c NA 103 a Myristoylation of Arf6 G2A peptide. Analysis using HPLC. b Demyristoylation Myr-H3K9 peptide. Analysis using HPLC. c Demyristoylation Myr-H3K9 peptide in the presence of rRNA. Analysis using HPLC. Given the reported tumor suppressor activity of SIRT6, it is attractive to imagine compounds that could selectively activate SIRT6 through interactions with the hydrophobic groove. Indeed, multiple studies have identified compounds able to activate SIRT6 deacetylation activity 13. Crystal structures consistently reveal SIRT6 activating compounds bound in the hydrophobic groove. It is therefore unsurprising that when tested in demyristoylation assays these compounds act as inhibitors 13a, 13c. SIRT6 deacetylation inhibitors have also been developed and have been shown to have potential efficacy in type II diabetes and multiple sclerosis models 14. These inhibitors were not tested against SIRT6 deKFA activity so what role SIRT6 deKFA activity has in these contexts is unclear. Thiomyristoyl peptides can inhibit SIRT6 deKFA by taking advantage of SIRT6 activity to generate a covalent stalled intermediate 15. Future SIRT6 inhibitors could use this as a starting point for more potent compounds. Such an approach has proven successful for SIRT2 inhibitors as will be discussed below. SIRT2 SIRT2 was the next enzyme found to have endogenous substrates for its deKFA activity 16. SIRT2 has been extensively studied due to its diverse physiological roles and its potential as a therapeutic target for certain cancers and neurological disorders 17. In cancer, SIRT2 can play both a tumor promoting and tumor suppressing role 17b. For instance, SIRT2 can promote breast cancer development by deacetylating Slug and aldehyde dehydrogenase 1A1 (ALDH1A1) 18. On the other hand, SIRT2 can reduce the activity of peroxiredoxin-1 (Prdx-1) through deacetylation which leads to breast cancer cells accumulating reactive oxygen species (ROS) and becoming less viable 19. Regardless, inhibition of SIRT2 leads to degradation of c-Myc and reduced growth in a broad variety of cancer cells, demonstrating its efficacy as a drug target 20. Unlike SIRT6, SIRT2 has strong in vitro deacetylase activity and has a plethora of reported deacetylase targets 12, 17a. While SIRT2 is a well-established deacetylase, it was also found to be able to hydrolyze lysine myristoylation with comparable efficiency to acetylation 12, 21. Similar to SIRT6, structural analysis of SIRT2 crystalized with a myristoyl-lysine peptide revealed a hydrophobic pocket that can readily accommodate a fatty acyl group (Figure 1.2) 21. SIRT2 inhibitors are numerous and diverse. Inhibitors of SIRT2 can be broadly categorized into two classes: activity-based and non-activity-based. Activity-based SIRT2 inhibitors usually have a peptide backbone and contain a thioacyl moiety that reacts with NAD+ in the SIRT2 active site to form a covalent stalled intermediate. Non-activity-based SIRT2 inhibitors function through a more typical manner by binding tightly at or near the active site. Direct comparison of SIRT2 inhibitors revealed that non-activity-based inhibitors AGK2, SirReal2, and Tenovin-6 can inhibit in vitro SIRT2 deacetylase activity, but not deKFA activity. Activity-based inhibitor TM was able to inhibit both activities in vitro 22, but not much in cells. However, simultaneous inhibition of SIRT2 deacetylase and deKFA activity can be achieved with a proteolytic targeting chimera (PROTAC) strategy to selectively degrade SIRT2 23. Crystal structures of inhibitors bound SIRT2 have been solved for several inhibitors 24. A recuring theme in these structures is the contribution of residues in the SIRT2 hydrophobic pocket for interaction with the inhibitors. While SIRT2 inhibitors have yet to be used in a clinical setting, cellular and mouse studies have yielded encouraging results for the use of SIRT2 inhibitors in treating disease. As mentioned above SIRT2i reduced growth of numerous different cancer cell lines 20. SIRT2i has also been shown to decrease α-synuclein toxicity in Parkinson’s disease models 25. The exact mechanism of action for this effect is still unclear and there is some debate about the causative or protective role for SIRT2 in Parkinson’s disease 17c. HDAC11 The most recent enzyme found to hydrolyze KFA is HDAC1126. HDAC11 is the newest member of the HDAC family and is the only class IV HDAC in humans. Since its discovery, HDAC11 has been found to play a role in neuronal function, immune regulation, and metabolic homeostasis 27. In addition, HDAC11 is overexpressed in several cancers and silencing HDAC11 can cause cell death in some cancer cell lines 28. In in vitro studies, HDAC11 lacks detectable deacetylation activity. Instead, HDAC11 is highly active towards KFA modified substrates 26. HDAC11 kinetics are similar to SIRT2 in hydrolyzing KFA, but HDAC11 is far more selective towards this unique modification. Unfortunately, no crystal structure has been obtained for HDAC11. HDAC11 modulation has been shown to affect acetylation of several proteins, but catalytic dead HDAC11 mutants were not utilized in any of these studies so whether or not HDAC11 has any bona-fide deacetylation substrates remains unclear 29. HDAC11 is reported to interact with HDAC6 so it is possible that changes in acetylation following HDAC11 modulation occur indirectly through HDAC6 or with the help of some unidentified cofactor or interacting partner that is lost in purification of HDAC11 for in vitro assays 30. Further work is necessary to determine whether and how HDAC11 regulates protein acetylation. Compared to SIRT6 and SIRT2, HDAC11 is understudied. The number of published inhibitors reflects this. A common strategy for creating inhibitors for Zn2+-dependent HDACs is to design a molecule that can chelate Zn2+ and has isoform specific interactions with residues surrounding the active site. This strategy was successfully employed in the design of SIS17, which has a fatty acyl moiety which likely interacts with HDAC11 in a similar manner to a KFA substrate 31. Another HDAC11 inhibitor, FT895, has shown promising anti-cancer activity in lung adenocarcinoma cells by suppressing Sox2 expression 32. Additionally, a natural product garcinol was shown to inhibit HDAC11 selectively 33. Garcinol has several reported biological activities. Although the relevance of HDAC11 inhibition in its various biological activities is unclear, it is interesting to note that biological effects of garcinol in mouse models share some commonality with HDAC11 knockout 33. Given HDAC11’s substrate specificity for KFA hydrolysis, use of HDAC11-specific inhibitors could help illuminate the role of KFA in a biological context. LYSINE FATTY ACYL TRANSFERASES One of the major impediments to studying KFA is the identification of human KFA transferase enzymes. Knowledge of how proteins acquire KFA would be invaluable in understanding the purpose of this modification. For instance, some bacterial toxins with known physiological importance take advantage of this activity during pathogenesis to promote infection (Figure 1.3). Although human lysine fatty acyl transferases are known, the acyl transferases for many of the reported KFA-modified proteins are still unknown. It is possible that such enzymes do exist but require more future effort to identify. However, an alternative explanation for the presence of endogenous KFA is that this modification simply arises from an S-N transfer. In this model a free lysine acts as a nucleophile to steal a fatty acyl group from a palmitoylated cysteine or palmitoyl-CoA. An amide bond is more stable than a thioester bond, making this reaction thermodynamically favorable. Several KFA transferases have been characterized and are reviewed below in order of discovery. Figure 1. 3. Bacterial KFA transferases. (A) The genetic structure for a generic RTX toxin operon includes five genes. The RtxC toxin is modified with KFA by RtxA and secreted through a transmembrane spanning pore consisting of RtxB, RtxD, and TolC. In the extracellular space, RtxC binds Ca2+ with characteristic nonapeptide repeats and associates with mammalian cell membranes. Oligomerization of RtxC forms large lytic pores in the membrane. (B) The structure of the Vibrio cholerae MARTX toxin is illustrated. Interior domains vary from species to species, but the terminal RTX domains in a MARTX toxin allows for translocation through the cell membrane. Once in the cell the cysteine protease domain (CPD) is activated to cleave the toxin at several points releasing the other domains: the actin crosslinking domain (ACD), the α/β hydrolase domain (ABH), and the Rho-inactivating domain (RID). RID modifies RhoA-family GTPases with KFA to suppress multiple cellular processes. (C) During part of the pathogenesis of Shigella flexneri, bacteria reside in an intracellular vacuole that can be destroyed through autophagy involving CHMP5. IcsB blocks autophagic destruction by catalyzing KFA on CHMP5. Figure created with BioRender.com. RtxC proteins Several species of bacterial pathogens employ a class of secreted toxin proteins known as Repeats in ToXin (RTX) toxins. RTX toxins function in various ways, but typically act as cytolysins by forming pores in the plasma membrane of mammalian cells 34. These proteins have anywhere from six to more than fifty repeats of a characteristic nonapeptide sequence G-G-X-G- (N/D)-D-X-(L/I/V/W/Y/F)-X that, when bound to Ca2+, forms a parallel β-roll motif 35. Structures in the RTX toxin operon vary, but typically consist of four genes: rtxA-D. rtxA encodes the actual toxin which is secreted through a type one secretion system encoded by rtxB, rtxD, and a third gene, tolC, located elsewhere on the bacterial chromosome. RtxA is synthesized as a protoxin that is activated by fatty acylation on one or two highly conserved lysine residues catalyzed by RtxC (Figure 1.3A) 36. The best characterized RtxC member is HlyC, which catalyzes KFA on the Escherichia coli hemolysin toxin HlyA. Initial characterization of the HlyA toxin revealed that HlyC activates proHlyA in a manner dependent on the acylated acyl carrier protein (acyl-ACP) 37. Follow up studies determined through mass spectrometry that HlyC activates HlyA by directing fatty acylation of two internal lysines 38. While the majority of HlyA lysine acylation is myristoylation (C14), HlyC apparently has some flexibility in substrate preference as saturated C15 and C17 acylation was also detected at appreciable amounts 39. Similar findings were made for the Bordetella pertussis CyaA toxin which is palmitoylated on a single lysine 40. Interestingly, when recombinantly expressed with its cognate acyl transferase CyaC in E. coli, CyaA is palmitoylated on a second lysine 41. This reveals that the number of lysines and the nature of the transferred acyl chain vary by each unique enzyme as well as the species background. More detailed information of RtxC enzymology was revealed using HlyC 42. HlyC acylation of proHlyA occurs through a ping pong mechanism involving two steps. In the proposed model, the acyl group from acyl-ACP is first transferred to His23 to form a covalent acyl-HlyC intermediate. This His is conserved throughout RtxC proteins. Then, the acyl group is transferred to the lysine residues on proHlyA. Ser20 is also important for optimal activity. While detailed enzymology has not been carried out for many RtxC proteins, the enzymatic mechanism is likely to be similar due to the conservation of relevant catalytic residues. Indeed, the analogous active site His and Ser in CyaC are necessary for its activity 43. Structural information for RtxC proteins is sparse but revealing. Despite unidentifiable sequence homology, ApxIC, the RtxC from Actinobacillus pleuropneumoniae, has conspicuous structural homology to the Gcn5-like N-acetyl transferase (GNAT) superfamily 44. GNAT proteins are well established acyl-transferases that use acyl-CoA as an acyl donor 45. The ApxIC structure is differentiated from other GNAT proteins by the lack of elements that typically interact with CoA, explaining why characterized RtxC proteins do not utilize acyl-CoA as the acyl donor. The conserved active site residues were found to reside within a deep surface groove. While further structural information is needed, this study, along with the fact that RtxC proteins are well conserved, opens the door for further understanding of these unique enzymes and raises the possibility of designing inhibitors to block toxin function. RID Most RtxA toxins have a similar structure and function. However, there is a subset of multifunctional RtxA toxins that are much larger called Multifunctional Autoprocessing Repeats in ToXin (MARTX) toxins. MARTX toxins contain the trademark nonapeptide repeats in both the N- and C-terminus, which forms a pore in the cell membrane that translocates the interior portion of the toxin encoding a modular array of effector domains into the cytoplasm 46. The interior domains vary in both number and type. However, a cysteine protease domain (CPD) is universally conserved in MARTX toxins. Once internalized, the CPD is activated and proteolytically cleaves the toxin at several points, releasing additional effector domains into the cell 47. One of these effectors present in multiple species was dubbed the Rho inactivation domain (RID). RID is present in several known MARTX toxins 46. The best characterized RID domain is the one in the Vibrio cholerae toxin, MARTXVc (Figure 1.3B). MARTXVc is known to induce cell rounding through an actin crosslinking domain (ACD) that covalently links actin monomers, preventing actin polymerization 48. However, cell rounding was still observed when this domain was knocked out. Follow up studies determined that this occurred through RID, which substantially decreases the amount of GTP-bound Rho GTPases Rho, Rac, and Cdc42 49. The mechanism by which RID functions was not known until a structure of the Vibrio vulnificus RID domain was solved 50. Clues from this structure allowed the authors to determine that RID catalyzes palmitoylation on lysines in the polybasic region of RhoA-family GTPases. This activity was dependent on C-terminal prenylation of the GTPases and could utilize palmitoyl- CoA as a acyl donor. Small GTPases are activated by guanine nucleotide exchange factors (GEFs) by stimulating the release of GDP, to allow the binding of GTP. RID-catalyzed KFA on Rac1 inhibits its interaction with GEFs, but how KFA prevents GEF interaction is unclear. Both the ACD and RID from MARTXVc lead to an obvious cell rounding phenotype by preventing actin dynamics. It is reasonable to wonder why V. cholerae would evolve to have two domains to carry out the same role. Woida and Satchell recently discovered that RID and a third domain in the MARTXVc, a α/β hydrolase (ABH) domain, function to suppress proinflammatory signaling 51. Cytoskeletal collapse caused by ACD activates mitogen-activated protein kinase (MAPK) signaling, leading to upregulation and enhanced secretion of proinflammatory cytokines. RID inactivation of Rac1 blocks MAPK signaling subsequent cytokine secretion (Figure 1.3B). RhoA-family GTPases also have other functions in immune processes, so RID could be playing a broad immunosuppressive function to prevent V. cholerae clearance by leukocytes 52. IcsB The Shigella flexneri toxin IcsB is another enzyme identified to catalyze KFA 53. Shigella are gram negative bacteria that cause a form of bacillary dysentery known as shigellosis. Shigella can colonize the intestinal epithelium through a complicated mechanism with the aid of toxin effectors secreted by a type three secretion system 54. Following ingestion, Shigella induce endocytosis by M cells in the epithelium of the colon 55. Endocytosed Shigella are transferred to resident macrophages where they employ a barrage of secreted effectors to escape the vacuole and replicate in the cytosol of the macrophage 56. This leads to lysis of the macrophage and dissemination of bacterial progeny. One of the critical effectors that enables this mode of infection is IcsB. IcsB promotes infection in several ways including preventing autophagic destruction and promoting cell lysis 57. How IcsB functions to modulate multiple cellular processes had been unclear, but it was predicted to be an enzyme through bioinformatic analysis 58. IcsB was found to be a potential homolog of RID and ectopic expression of IcsB was found to disrupt the actin cytoskeleton. In line with these observations, IcsB was also found to catalyze KFA on lysines in the polybasic region of Rho GTPases 53. The eighteen carbon stearoyl CoA seems to be the preferred acyl donor and prenylation of Rho GTPases is also necessary for IcsB activity. Proteomic analysis identified numerous IcsB substrates in addition to Rho GTPases. KFA on one of these substrates, CHMP5, was identified to inhibit the autophagic destruction of S. flexneri thus providing a mechanism for sustained intercellular survival (Figure 1.3C). While RID and IcsB are both bacterial toxins that catalyze KFA of host substrates, they differ in a few key aspects. IcsB transfected cells exhibit cell rounding whereas S. flexneri infected cells do not. Cells both introduced to RID alone as well as V. cholerae infected cells exhibit cell rounding. This discrepancy is in part due to the presence of the V. cholerae ACD as well as to the observation that S. felxneri activate actin polymerization for part of their pathogenesis. Additionally, while no proteomic search for substrates has been done, RID is only known to modify Rho GTPases while IcsB can modify many other substrates in addition to Rho GTPases. What role KFA plays on these other substrates is an area for future study. NMT1 and NMT2 N-myristoyltransferases (NMTs) have long been known to transfer a myristoyl group from myristoyl-CoA to the α-amine of an N-terminal glycine following cleavage of the initiator methionine 59. This modification happens co-translationally or after proteolytic events that result in a free N-terminal glycine such as caspase cleavage 60. There are no known enzymes that can hydrolyze N-terminal myristoylation and as such it is thought to be irreversible. Like KFA, N- terminal myristoylation increases a protein’s affinity for membranes. Knocking out NMT is embryonically lethal in mice and fruit flies 61. NMT levels are elevated in several cancers and NMT inhibition has advanced to clinical trials for the treatment of NMT deficient blood cancers 62. Additionally, several viruses utilize endogenous host NMTs for their replication and infectivity raising the possibility of using NMT inhibitors as an antiviral agent 63. Because NMTs catalyze the fatty acylation of a primary amine, they can do the same chemistry as a potential KFA transferase. Indeed, two groups found that the human NMT1 and NMT2 can catalyze KFA on lysines towards the N-terminal of a peptide 16c, 64. The proposed mechanism for this reaction requires rotation around the peptide bond to present the lysine ε- amine into the active site typically occupied by a glycine α-amine (Figure 1.4). Both amines can freely rotate during NMT catalysis and myristoyl-glycine and myristoyl-lysine are similarly positioned in the active site of NMT structures, supporting the proposed activity (Figure 1.4). The precise sequence requirements for this activity are still being clarified, but it is clear that this activity is strongest when lysine is closer to the N-terminus. While glycine is the preferred substrate, KFA is efficiently catalyzed on a lysine that is right after the glycine, especially when the N-terminal amine is blocked. KFA transferase activity is decreased when an extra glycine is added before the modified lysine and is absent when two additional glycines are inserted. This narrows the scope of potential substrates for NMT KFA transferase activity. Peptide experiments indicate that a good KFA transferase substrate for NMTs has a small amino acid at position two, a lysine at position three, a serine at position six, and lysine at position seven. This was confirmed when Arf6 was identified to be a substrate of NMT KFA transferase activity 16c. The Arf6 N-terminal sequence following initiator methionine cleavage is GKVLSKIF. Arf6 can be doubly myristoylated at both G2 and K3. It was proposed that NMTs can accommodate both myristoyl groups when one is inserted into a solvent channel in the protein structure. While Arf6 is currently the only known substrate for NMT KFA transferase activity, it is feasible that additional proteins with N-termini similar to Arf6 could also be modified 65. Figure 1. 4. Model for NMT KFA transferase activity. Rotation around the peptide backbone (bold) positions the amine of a lysine side chain similarly to the amine of an N-terminal glycine. Both are primary amines that can rotate freely during NMT catalysis. The superimposed myristoyl-lysine (green stick representation) and myristoyl-glycine (white stick representation) in human NMT2 (PDB 6PAU) and NMT1 (PDB 5O9V) structures overlap nicely, supporting this model. FUNCTIONS OF LYSINE FATTY ACYLATION A diverse set of proteins has been identified to have KFA (Table 1.2). How exactly KFA affects the modified protein is often unclear, but a common theme is a change in membrane affinity (Figure 1.5). If a protein is otherwise soluble, KFA can lead to membrane localization as is the case for IL-1α or SHMT2. For proteins that already have membrane targeting elements, KFA can change which cellular membranes they are localized to. This is the case for several small GTPases which are targeted to membranes through electrostatic interactions and other lipid modifications. In this section we review proteins known to have KFA, how KFA on the protein was found, and what is known about how KFA affects protein function. Not discussed in this section are the substrates for RID and IcsB toxins which are examined above or in other publications 50, 53. IL-1α Interleukins (ILs) are a family of secreted proteins used in cell-to-cell communication that regulate inflammatory signaling. The IL-1 family of proteins contains 11 different members that are synthesized as precursor proteins, requiring proteolytic processing for optimal biologic activity 66. IL-1α and IL-1β are closely related pro-inflammatory members of the IL-1 family. Despite obvious structural similarity between the two proteins, they differ in some key respects 67. While IL-1β functions exclusively as a secreted protein, IL-1α has activity both secreted and bound to the cell membrane 68. In examining its affinity for membranes, it was determined that IL-1α was doubly myristoylated on the N-terminal portion of the protein 69. This was determined through the incorporation of radiolabeled myristic acid. Incubation of synthetic peptides with monocyte lysates identified lysines 82 and 83 as sites of myristoylation 69a. Myristoylation was also observed for IL-1β, but to a much less extent. While the precise role KFA plays on IL-1α Figure 1. 5. Models of KFA regulation. (A) KFA on TNF-α directs it to the lysosome for degradation. SIRT6 hydrolyzes TNF-α KFA leading to higher levels on the plasma membrane and increased secretion of cleaved, soluble TNF-α. (B) KFA on the polybasic region of R-Ras2 results in increased plasma membrane localization, PI3K interaction, AKT signaling, and cell proliferation. SIRT6 hydrolyzes R-Ras2 KFA, releasing it from the membrane and decreasing cell proliferation. (C) K-Ras4a with KFA preferentially localizes at the plasma membrane. SIRT2 hydrolyzes K-Ras4a KFA. Cysteine palmitoylation on K-Ras4a retains membrane affinity, but instead localizes to endomembranes where K-Ras4a interacts with the signaling kinase A-Raf leading to downstream cellular transformation. (D) KFA on RalB also directs it to the plasma membrane to interact with Sec5 and Exo84 and promote cell migration. SIRT2 hydrolyzes RalB KFA, decreasing plasma membrane localization. (E) NMT-catalyzed KFA on K3 of Arf6 increases plasma membrane localization. GTP hydrolysis at the plasma membrane aided by a GTPase activating protein (GAP) causes Arf6 N-terminal glycine myristoylation to be sequestered. Arf6 is then trafficked to the endocytic recycling compartment (ERC) where SIRT2 hydrolyzes KFA and Arf6 is reloaded with GTP. NMT then starts the cycle again. Both the GTP and KFA dynamic cycle promote ERK signaling. Figure created with BioRender.com. function is unclear, only the uncleaved IL-1α, with modified lysines present, associates with the cell membrane 70. It is not known what enzymes could regulate IL-1α KFA, but acylation in the presence of lysates raises the possibility of an unidentified KFA transferase. Membrane associated IL-1α contributes to arthritis in a mouse model, so modulating IL-1α KFA could have therapeutic applications 71. Aquaporin-0 Aquaporins are a family of transmembrane channel proteins that facilitate the transport of water across the plasma membrane. Aquaporin-0 (AQP0) is highly abundant in the ocular lens where it plays an important role, not only in circulating water, but as a cell to cell adhesion protein for maintenance of proper tissue structure 72. There is very little protein turnover in lens proteins due to the loss of fiber cell organelles during differentiation, making proteins in this tissue an interesting model for aging. AQP0 is known to accumulate post-translational modifications with age 73. To determine what fatty acylations may be accumulating on AQP0, Schey et al. carried out mass spectrometric analysis of hydrophobic peptides from bovine and human lens tissue 74. They identified AQP0 as containing two fatty acylations: one on the N- terminal methionine and one on a highly conserved lysine. The identity of AQP0 KFA modification varies from C16 to C20 with up to 4 desaturations at ratios closely resembling the abundance of phosphoethanolamine lipids in lens membranes 75. This suggests that either the modification is accumulating non-enzymatically or that a potential KFA transferase has limited preference in acyl group. The effect KFA has on AQP0 function is unknown, but the modified protein partitions to detergent-resistant membrane fractions. This suggests a role in membrane domain targeting. Table 1. 2. Proteins with lysine fatty acylation KFA KFA Name Species Effect of KFA References transferase hydrolase IL-1α Human NA NA NA 69 Human, Aquaporin-0 NA NA NA 74 Bovine Decreased TNF-α Human NA SIRT6 TNF-α 77-78 secretion Increased cell R-Ras2 Human NA SIRT6 80 proliferation Decreased cell K-Ras4 Human NA SIRT2 16b proliferation H-Ras Human NA NA NA 16b Increased cell RalB Human NA SIRT2 16a migration Promotes Arf6 Human NMT1/NMT2 SIRT2 16c GTPase cycle Reduced ubiquitination SHMT2 Human NA HDAC11 and enhanced 26a signalling of INFαR1 Lipid raft Human, recruitment, Gravin-α NA HDAC11 106 mouse increased β-AR signalling GTPase inactivation; defect in actin RhoA Human RID NA polymerization; 50-51 GTPases reduced inflammatory signaling Defect in S. flexneri CHMP5* Human IcsB NA 53 autophagic destruction Increased Many gram binding to β2- RTX toxins negative RtxC NA integrins; 36 bacteria promotion of oligomerization *CHMP5 is one of many identified IcsB substrates. For a complete list see Liu et al., 2018. TNF-α Tumor necrosis factor α (TNF-α) is a proinflammatory cytokine secreted by several immune cells. TNF-α is translated with a single transmembrane domain that is cleaved at the plasma membrane to release a soluble form that binds TNF receptors (TNFRs). TNFR1 is universally expressed on almost every cell type meaning that TNF-α signaling is both ubiquitous and tightly regulated76. In a follow up study to the IL-1α findings highlighted above, TNF-α was also identified to have KFA 77. Modification is present on two lysines, K19 and K20, at the intracellular end of the transmembrane helix. SIRT6 was found to hydrolyze KFA on TNF-α which in turn promotes its secretion 7. KFA causes TNF-α to accumulate in the lysosome where it is degraded 78. In this context, KFA thus serves an anti-inflammatory role by decreasing the amount of secreted TNF-α (Figure 1.5A). In addition to TNF-α, a proteomic study revealed that SIRT6 deKFA activity regulates the secretion of numerous other proteins 79. Interestingly, SIRT6 deKFA activity decreases the secretion of ribosomal proteins via exosomes, but the detailed mechanism for how this happens is unknown. R-Ras2 The only other known endogenous substrate for SIRT6 deKFA activity is R-Ras2, a member of the Ras family of GTPases 80. Ras GTPases are known to be targeted to cellular membranes by their C-terminal hypervariable regions through cysteine lipidation and polybasic regions. R-Ras2 was identified to have KFA in its polybasic region using mass spectrometry. Mutating a patch of four lysine residues to arginine in the polybasic region of R-Ras2 abolished KFA. Immunofluorescence analysis demonstrated that KFA enhances R-Ras2 plasma membrane localization. There, R-Ras2 exist more in the GTP-bound state, interacts with PI3K, activates Akt signaling, and upregulates cell proliferation. SIRT6 KO MEFs are more proliferative than WT, an effect which is rescued by suppressing R-Ras2 level. Together this provides a model where SIRT6 acts as a tumor suppressor by inhibiting R-Ras2 and downregulating proliferative PI3K/Akt signaling (Figure 1.5B). K-Ras4a The first identified SIRT2 deKFA substrate was K-Ras4a 16b. After the identification of R-Ras2 as a KFA substrate, Jing et al. examined other small GTPases with polybasic regions similar to R-Ras2 for potential KFA modification. These included the well-known oncoproteins H-Ras, N-Ras, K-Ras4a, and K-Ras4b. Using mass spectrometry, they identified both H-Ras, and K-Ras4a as having KFA in their polybasic region. Sirtuins with known deKFA activity were screened against both H-Ras and K-Ras4a and SIRT2 was found to remove K-Ras4a KFA. Removal of H-Ras KFA was not observed for any of the screened enzymes. The oncogene KRAS (which is alternatively spliced into K-Ras4a and K-Ras4b) is the most frequently mutated gene in cancer, so identifying how modifications regulate its activity is of great interest 81. KFA on K- Ras4a was found to inhibit intracellular puncta localization, interaction with the signaling kinase A-Raf, and downstream cellular proliferation (Figure 1.5C). By removing K-Ras4a KFA, SIRT2 serves a tumor promoting role. RalB The identification of R-Ras2 and K-Ras4a as proteins with KFA motivated Spiegelmen et al. to further expand the search further into the of Ras subfamily small GTPases 16a. In so doing they identified RalB having KFA. Like R-Ras2 and K-Ras4a, RalB is modified with KFA in its C-terminal polybasic region. Mutating all eight lysines in this region to arginines abolished KFA signal. Screening enzymes with deKFA activity identified SIRT2 as being able to remove KFA from RalB. RalB has been identified to promote multiple cancer phenotypes 82. Studying the effect of RalB KFA on these phenotypes revealed that RalB KFA enhanced migration of A549 lung cancer cells but did not affect cell proliferation. KFA of RalB was also found to increase GTP loading, plasma membrane localization, and co-localization with its effector proteins Sec5 and Exo84 (Figure 1.5D). Contrary to the K-Ras4a mechanism, SIRT2 by removing RalB KFA, decreases RalB activation and cell migration. This juxtaposition underscores the diverse, context-dependent role of KFA and the enzymes that modulate it. Arf6 Arf6 is a small GTPase in the ADP-ribosylation factor (Arf) subfamily that has been found to be modified with KFA 16c. While members of the Ras subfamily of GTPases discussed above are anchored to membranes through lipidation and electrostatic interactions in their C-terminus, the Arf family is targeted to membranes via N-terminal glycine myristoylation catalysed by NMTs 83. Unlike KFA, there are no enzymes that are known to hydrolyse glycine myristoylation. For Arf proteins, regulating membrane association occurs through a nucleotide-dependent conformational shift that flips the myristoyl group back into the protein to sequester it in a hydrophobic pocket 84. Unlike Arf1-5, Arf6 has a lysine in the third position of its sequence. Interestingly, K3 was found to be modified with KFA by NMT enzymes and the modification is hydrolysed by SIRT2. NMT activity is greatest towards the active, GTP-bound, form of Arf6 where SIRT2 prefers the inactive, GDP-bound, form. This enzyme preference links a dynamic KFA cycle to the GTPase cycle. Indeed, inhibiting either SIRT2 or NMTs decreased phosphorylation of ERK, a downstream target of Arf6 activation. A cycle of dynamic KFA modification thus drives the Arf6 GTPase cycle (Figure 1.5E). SHMT2 Serine hydroxymethyltransferase 2 (SHMT2) is a key member of one-carbon metabolism. SHMT2 catalyses the reversible conversion of serine and THF to glycine and methylene-THF in the mitochondria. High levels of SHMT2 are associated with poor patient prognosis in several cancers and inhibitors that target both SHMT2 and SHMT1 (which catalyses the same reaction in the cytoplasm) have shown promising results in initial cellular and mouse studies 85. Additionally, SHMT2 was found to regulate immune signalling as a component of the BRISC deubiquitylase complex. The BRISC complex is known to hydrolyse K63-linked polyubiquitin chains on type I IFN receptor chain 1 (IFNαR1), a key receptor in the antiviral response. BRISC prevents degradation of the receptor and promotes recycling to the plasma membrane 86. SHMT2 was identified to have KFA during a search for substrates of HDAC11 26a. KFA on SHMT2 did not affect its in vitro activity in converting serine and THF to glycine and methylene-THF. Instead, KFA was found to be relevant in SHMT2’s role in the BRISC complex. Increasing SHMT2 KFA, through HDAC11 knock out or knock down, increased SHMT2 localization to the endosomes/lysosomes where the BRISC complex can hydrolyse IFNαR1 ubiquitination. Correspondingly, HDAC11 KO cells had more IFNαR1 on the cell surface following IFNα treatment. Further, HDAC11 KO cells had stronger downstream signalling following IFNα stimulation and HDAC11 KO mice also had a stronger antiviral response when challenged with vesicular stomatitis virus. In this context, KFA again controls intracellular localization of the substrate protein, here leading to enhanced cellular activity of a deubiquitylase complex. Inhibiting HDAC11 may therefor enhance antiviral responses. Gravin-α Gravin-α, also known as A-kinase anchoring protein 12 (AKAP12), is a scaffold protein thought to act as a signalosome hub by interacting with pools of phosphatases and kinases 104. Gravin-α is recruited to the plasma membrane in response to β-adrenergic receptor (AR) stimulation to compartmentalize protein kinase A (PKA) signalling and activate downstream signalling pathways important for cardiac function and adipose thermogenesis 105. Gravin-α was identified to have increased fatty acylation levels in a SILAC screen of HDAC11 knockdown mouse embryonic fibroblasts 106. Subsequently, two lysines within the PKA binding domain of gravin-α were identified to be myristoylated and regulated by HDAC11. Lysine myristoylation of gravin-α was found to be required for its role in activating β2- and β3-AR-mediated uncoupling protein 1 induction, an important step in the “beiging” of adipose tissue 107. Mechanistically, lysine myristoylation increases gravin-α localization to detergent-resistant membrane fractions with calveolin-1, a scaffolding protein critical to the activation of β3-AR signalling in adipocytes. These findings further cemented the role of KFA in increasing protein affinity for specific cellular membranes and provided evidence for the efficacy in inhibiting HDAC11 to promote energy expenditure during obesity treatment. RTX Toxins The RtxA family of proteins are toxins that are secreted by many pathogenic gram- negative bacteria 87. As discussed above, RtxA proteins, hereby referred to as RTX toxins, are activated via lysine fatty acylation catalyzed by RtxC proteins. Once secreted, RTX toxins function in a variety of different ways. The best-known mode of action for RTX toxins is cytolysis and hemolysis through formation of pores in the cell membrane 88. RTX toxins primarily target leukocytes and form pores in the cell membrane in a calcium dependent manner 89. Hydrophobic regions towards the N-terminus are believed to interact with the target cell membrane to form cation-selective pores, leading to cell lysis 90. KFA is necessary for the cytotoxicity of all known pore forming RTX toxins, but the mechanism by which KFA affects RTX toxin function is still unclear. Unacylated RTX toxins from E. coli and B. pertussis, HlyA and CyaA, respectively, are both able to form pores in liposomes and planar lipid bilayers 91. At concentrations insufficient to cause cell lysis, RTX toxins bind to β2-integrins and activate downstream signalling pathways that lead to apoptosis 92. KFA was found to increase the affinity to β2-integrin receptors for CyaC and the A. actinomycetemcomitans toxin LtxA 93. It was reported that KFA on HlyA is important for the formation of oligomers in erythrocyte membranes, but the physiological importance of RTX toxin oligomerization is unclear 94. It is conceivable that KFA on RTX toxins may promote localization to appropriate membrane subdomains such as lipid rafts, somehow increasing toxin function. For instance, the K. kingae RtxA binds cholesterol, a membrane component enriched in lipid rafts, for optimal activity and palmitoylation on cysteines is thought to target transmembrane proteins to lipid rafts 95. KFA on RTX toxins could also potentially play a role in maintaining an appropriate structural confirmation. Additional studies are necessary to identify the precise mechanism by which KFA regulates RTX toxins. TOOLS FOR STUDYING LYSINE FATTY ACYLATION Methods to detect endogenous KFA Research involving KFA is well suited to a chemical biology approach. The most commonly used approach when assaying KFA is the use of fatty acyl alkyne probes as discussed below. However, this approach has inherent limitations. First, one must add alkyne probes exogenously which is unfeasible for applications in mice or other tissue samples. Second, when used in cell cultures, a working concentration is ~50 μM which may be sufficient to unintentionally stimulate signaling pathways involving fatty acids or to increase protein lipidation due to artificially high amounts of fatty acids. One way to assay levels of endogenous KFA is to purify a protein of interest and to attempt to identify KFA with mass spectrometry 16c. Mass spectrometry can directly identify the modified residue but is technically challenging due to the relatively low abundance of KFA and the hydrophobicity of the modification which makes processing samples for MS difficult. Endogenous KFA can be indirectly detected using a [32P]NAD+ TLC assay if the modification is able to be hydrolyzed by a sirtuin 16b, 16c, 80. In this assay, a protein of interest is incubated with a sirtuin that has KFA hydrolase activity and radiolabeled [32P]NAD+. Sirtuin-catalyzed lysine deacylation results in the formation of [32P]O- acylADPR. Fatty acyl-ADPr is drastically more hydrophobic than NAD+ and can be easily separated on a TLC plate. Phosphorescence detection can then be used to analyze the presence of fatty acyl-ADPR and, by extension, KFA on the protein of interest (Figure 1.6A). Additional tools to study endogenous KFA could be inspired by research of similar post translational modifications. Studies concerning lysine acetylation commonly employ antibodies specific to acetyl-lysine. Antibodies that could specifically bind KFA would allow for purification and analysis of endogenously modified proteins without any exogenous treatment. Additional tools for studying KFA are necessary to fully realize the full scope of its biological importance. Enzyme activity assays Assays measuring in vitro enzyme activity are necessary for establishing enzyme kinetics, substrate preferences, and for testing inhibitors. Once the proper cofactors and buffer conditions are determined for an enzyme of interest, measuring its activity in regulating KFA can be straightforward. Assays for RtxC enzymes indirectly measure activity by taking advantage of the hemolytic action of their cognate RtxA proteins 87, 96. This is not a broadly applicable technique for KFA enzymes and as such will not be further elaborated on. Here we briefly introduce several proven strategies for measuring the activity of KFA hydrolases or transferases (Figure 1.6). Figure 1. 6. Techniques for studying KFA enzyme activity. (A) [32P]NAD+ sirtuin assay. 32P (red atom) is incorporated into NAD+ to be used by a sirtuin with KFA hydrolase activity. During the hydrolysis mechanism, the sirtuin transfers the fatty acyl group from a modified protein to the cofactor releasing nicotinamide (NAM) and resulting in the formation of [32P]fatty acyl-ADPR. TLC plates are used to separate, visualize, and quantify radiolabeled species. (B) HPLC analysis of KFA enzyme activity on peptides. Unmodified and KFA-modified peptides are separated with HPLC and the area of the corresponding peaks are determined for quantification. (C) Fluorescence-based peptide KFA hydrolase assay. KFA is hydrolyzed from a lysine immediately preceding and AMC group. Trypsin hydrolyzes the amide bond following only the unmodified lysine releasing the AMC, which then exhibits fluorescence. (D) Acyl-cLIP assay for KFA enzymes. KFA on a peptide modified with fluorescein increases its affinity for micellar membranes. Fluorescein has slower tumbling near the bulky micelles, resulting in increased fluorescence polarization. Figure created with BioRender.com. A commonly used approach to assaying KFA hydrolase activity is to incubate the enzyme of interest with a short peptide containing a KFA-modified lysine. After quenching the reaction, the products can be analyzed using HPLC or LC/MS (Figure 1.6B). For standard HPLC detection, the peptide ideally should have a strong chromophore which can be something as simple as a tryptophan residue. While this approach has been successfully implemented to study sirtuins, HDACs, and NMTs, it could theoretically be applied for studying RtxC enzymes 12, 16c, 21, 26a, 97. One of the benefits of this assay is that a peptide closely resembling the physiological protein substrate can be used. It allows for determination of an enzyme’s substrate specificity by assaying peptides of various amino acid sequences. Additionally, tandem MS can be used to confirm the site of modification. Drawbacks for this approach include the need for HPLC or LC/MS instruments and relatively low throughput. Acyl-peptide based assays for KFA hydrolases can also be analyzed with a fluorescence readout (Figure 1.6C). In this approach, peptides containing amino acids of choice with the acyl lysine at the C-terminal end followed by a fluorescent moiety such as 7-amino-4- methylcoumarin moiety (AMC) which will only fluoresce if released from the peptide 98. After incubation with enzyme the reaction is quenched by adding trypsin which cleaves amide bonds after lysines. Increased KFA hydrolase activity can thus be measured by increased fluorescence. This assay can be done quickly in a 96-well plate which allows researchers to test many different conditions at once and in replicates. This assay is well suited to screening libraries of compounds for enzyme inhibitors or activators. IC50 and EC50 values can be easily calculated by relating fluorescence values to the concentration of inhibitor or activator in the well. The drawback of this assay is the potential difficulty in synthesizing the appropriate peptide substrates. Furthermore, there are no amino acids on the C-terminal of the acyl-lysine, limiting the degree to which sequence specificity of a given enzyme can be ascertained. A similar technique is to modify a lysine with a fluorescent acyl group and to incorporate a fluorescence quenching moiety into the peptide. Hydrolysis separates the fluorophore from the quenching moiety and results in increased fluorescence. While this assay is also fast, the acyl group is necessarily bulky and may not have broad applicability 99. A promising novel technique for studying protein lipidation is acylation-coupled lipophilic induction of polarization (Acyl-cLIP) 100. In this assay, an enzymatic protein lipidation reaction is carried out on a fluorescein tagged peptide in the presence of detergent micelles. Changes in lipidation affect the peptides affinity for the micelles. When bound to the micelles, the fluorescein has decreased molecular tumbling and a corresponding increase in polarized fluorescence emission. The change in polarized fluorescence can be detected with fluorescence anisotropy measurements and this change can be attributed to the lipidation status of the peptide (Figure 1.6D). Drawbacks of this assay are the same as other peptide-based assays mentioned above. Benefits of this assay include a high throughput for kinetics and inhibitor studies, the ability to assay both lipid transferase and hydrolase enzymes, and a universal applicability to any type of lipid modification. Measuring KFA with fatty acyl probes If a target protein can be readily purified in large quantities, endogenous KFA levels can be analyzed with mass spectrometry, such as for APQ0 74. However, this approach is not broadly applicable for reasons outlined above. More commonly used tools for studying levels of KFA on a target protein involve the addition of exogenous fatty acid analogs (Figure 1.7A). An important caveat for all approaches that involve the incorporation of fatty acids is that more than just lysines can be fatty acylated. Cysteine palmitoylation and glycine myristoylation are both abundant modifications that involve fatty acyl groups and must be ruled out as the source of fatty acylation signal. Cysteine palmitoylation can be hydrolyzed by hydroxylamine while lysine acylations cannot. Treating a fatty acylated sample with hydroxylamine can thus eliminate an experimental signal from cysteine fatty acylation. Glycine myristoylation cannot be removed with hydroxylamine but does requires a glycine at the most N-terminal position so a cursory examination of the protein’s sequence can reveal if the modification is possible. One class of exogenous fatty acyl probes is radioactive-isotope-labeled fatty acids. Cells treated with radioactive fatty acids can then form radioactive fatty acyl-CoA for use by KFA transferases. A protein of interest can then be purified and monitored for radioactivity to assay for fatty acylation 69b. 3H- and 14C-labeled fatty acids are the best choice for this application as the isotopes can be incorporated into the fatty acid without modifying the structure. However, these isotopes have relatively low signal and must be monitored for a long time. Figure 1. 7. Metabolic probes to examine KFA on a substrate protein. (A) Structure of KFA probes. Bioorthogonal probes for KFA can mimic myristic acid or palmitic acid. Both azide and alkyne probes are applicable though alkyne probes more closely resemble the endogenous fatty acids. (B) Metabolic labeling of KFA proteins can be analyzed with CuACC. Alkyne labeled proteins can be modified with fluorophores or affinity tags like biotin to analyze KFA levels with in-gel fluorescence, western blot, or mass spectrometry. Clickable fatty acid analogs have proven to be a more powerful tool for studying KFA. After treating cells with these probes, proteins are modified with fluorophores or biotin via click chemistry to track levels of fatty acylation using fluorescence scanning, western blot, or various mass spectrometry approaches (Figure 1.7B). While azido fatty acids have been implemented for the study of cysteine palmitoylation, alkyne fatty acids have lower background and more closely resemble the structure of the endogenous fatty acids 101. There is a suite of fatty acyl alkyne probes that analogize natural fatty acids myristic acid, palmitic acid, and more (Figure 1.7A). In addition to analyzing a protein of interest, clickable fatty acid probes can help identify substrates of KFA transferases or hydrolases as has been done for IcsB and HDAC11 26a, 53. In this SILAC (stable isotope labeling with amino acids in cell culture) experiment, cells are cultured in media containing normal or isotopically heavy amino acids. KFA modulating enzymes are then overexpressed or suppressed and cells are treated with fatty acid alkyne probes. After probe incubation, cells are lysed and lysates from heavy and light media are mixed. Alkyne-labeled proteins are modified with biotin via click chemistry then treated with hydroxylamine to remove cysteine palmitoylation signal. The remaining biotinylated proteins are enriched with streptavidin and analyzed via MS. Further proteomic analysis in this way is necessary to fully appreciate the scope of KFA during various biological processes. SUMMARY AND FUTURE QUESTIONS Since the initial discovery of KFA on IL-1α, significant progress has been made in studying KFA. This is especially true of the last 10 or so years as several enzymes were identified to add or remove KFA. Additionally, implementation of clickable probes has allowed for the identification of many new proteins with KFA and has hastened the pace of discovery. We now know that KFA can regulate a variety of biological processes such as protein secretion, tumorigenesis, and immune signaling. The knowledge gained surrounding KFA can now serve as a basis for further exploration. Multiple species of pathogenic bacteria utilize KFA to enhance their pathogenesis. While mammalian cells are not yet known to have KFA transferases that act on a broad range of substrates, the presence of multiple enzymes with strong KFA hydrolase activity suggests an evolutionarily beneficial role for this activity. Is it possible that during certain bacterial infections KFA hydrolase activity serves a protective role against toxins like RID and IcsB? Is it possible that KFA hydrolase enzymes could remove RTX toxin KFA to inactivate the pore-forming toxin? Additional lines of future study are myriad. In addition to bacteria, could other pathogens modulate KFA during infection? In addition to infection, KFA has already been shown to modulate tumor phenotypes. What other diseases could be regulated by KFA? Can KFA guide the creation of therapeutics to counteract these diseases? Addressing these questions, will require identifying more proteins modified by KFA and additional enzymes that regulate this modification, as well as the development of small molecules targeting the regulatory enzymes. Future work needs to address these issues to fully understand the biological impact of KFA. REFERENCES 1. Jiang, H.; Zhang, X.; Chen, X.; Aramsangtienchai, P.; Tong, Z.; Lin, H., Protein Lipidation: Occurrence, Mechanisms, Biological Functions, and Enabling Technologies. Chem Rev 2018, 118 (3), 919-988. 2. Resh, M. D., Fatty acylation of proteins: new insights into membrane targeting of myristoylated and palmitoylated proteins. Biochim Biophys Acta 1999, 1451 (1), 1-16. 3. (a) Wang, B.; Dai, T.; Sun, W.; Wei, Y.; Ren, J.; Zhang, L.; Zhang, M.; Zhou, F., Protein N-myristoylation: functions and mechanisms in control of innate immunity. Cell Mol Immunol 2021, 18 (4), 878-888; (b) Schlott, A. C.; Holder, A. A.; Tate, E. W., N- Myristoylation as a Drug Target in Malaria: Exploring the Role of N-Myristoyltransferase Substrates in the Inhibitor Mode of Action. ACS Infect Dis 2018, 4 (4), 449-457; (c) Resh, M. D., Trafficking and signaling by fatty acylated and prenylated proteins. Nat Chem Biol 2006, 2 (11), 584-90. 4. Ringel, A. E.; Roman, C.; Wolberger, C., Alternate deacylating specificities of the archaeal sirtuins Sir2Af1 and Sir2Af2. Protein Sci 2014, 23 (12), 1686-97. 5. Gregoretti, I. V.; Lee, Y. M.; Goodson, H. V., Molecular evolution of the histone deacetylase family: functional implications of phylogenetic analysis. J Mol Biol 2004, 338 (1), 17-31. 6. Zhu, A. Y.; Zhou, Y.; Khan, S.; Deitsch, K. W.; Hao, Q.; Lin, H., Plasmodium falciparum Sir2A preferentially hydrolyzes medium and long chain fatty acyl lysine. ACS Chem Biol 2012, 7 (1), 155-9. 7. Jiang, H.; Khan, S.; Wang, Y.; Charron, G.; He, B.; Sebastian, C.; Du, J.; Kim, R.; Ge, E.; Mostoslavsky, R.; Hang, H. C.; Hao, Q.; Lin, H., SIRT6 regulates TNF-alpha secretion through hydrolysis of long-chain fatty acyl lysine. Nature 2013, 496 (7443), 110-3. 8. Chang, A. R.; Ferrer, C. M.; Mostoslavsky, R., SIRT6, a Mammalian Deacylase with Multitasking Abilities. Physiol Rev 2020, 100 (1), 145-169. 9. (a) Toiber, D.; Erdel, F.; Bouazoune, K.; Silberman, D. M.; Zhong, L.; Mulligan, P.; Sebastian, C.; Cosentino, C.; Martinez-Pastor, B.; Giacosa, S.; D'Urso, A.; Naar, A. M.; Kingston, R.; Rippe, K.; Mostoslavsky, R., SIRT6 recruits SNF2H to DNA break sites, preventing genomic instability through chromatin remodeling. Mol Cell 2013, 51 (4), 454- 68; (b) Zhong, L.; D'Urso, A.; Toiber, D.; Sebastian, C.; Henry, R. E.; Vadysirisack, D. D.; Guimaraes, A.; Marinelli, B.; Wikstrom, J. D.; Nir, T.; Clish, C. B.; Vaitheesvaran, B.; Iliopoulos, O.; Kurland, I.; Dor, Y.; Weissleder, R.; Shirihai, O. S.; Ellisen, L. W.; Espinosa, J. M.; Mostoslavsky, R., The histone deacetylase Sirt6 regulates glucose homeostasis via Hif1alpha. Cell 2010, 140 (2), 280-93. 10. (a) Dominy, J. E., Jr.; Lee, Y.; Jedrychowski, M. P.; Chim, H.; Jurczak, M. J.; Camporez, J. P.; Ruan, H. B.; Feldman, J.; Pierce, K.; Mostoslavsky, R.; Denu, J. M.; Clish, C. B.; Yang, X.; Shulman, G. I.; Gygi, S. P.; Puigserver, P., The deacetylase Sirt6 activates the acetyltransferase GCN5 and suppresses hepatic gluconeogenesis. Mol Cell 2012, 48 (6), 900-13; (b) Kawahara, T. L.; Michishita, E.; Adler, A. S.; Damian, M.; Berber, E.; Lin, M.; McCord, R. A.; Ongaigui, K. C.; Boxer, L. D.; Chang, H. Y.; Chua, K. F., SIRT6 links histone H3 lysine 9 deacetylation to NF-kappaB-dependent gene expression and organismal life span. Cell 2009, 136 (1), 62-74; (c) Michishita, E.; McCord, R. A.; Berber, E.; Kioi, M.; Padilla-Nash, H.; Damian, M.; Cheung, P.; Kusumoto, R.; Kawahara, T. L.; Barrett, J. C.; Chang, H. Y.; Bohr, V. A.; Ried, T.; Gozani, O.; Chua, K. F., SIRT6 is a histone H3 lysine 9 deacetylase that modulates telomeric chromatin. Nature 2008, 452 (7186), 492-6; (d) Michishita, E.; McCord, R. A.; Boxer, L. D.; Barber, M. F.; Hong, T.; Gozani, O.; Chua, K. F., Cell cycle-dependent deacetylation of telomeric histone H3 lysine K56 by human SIRT6. Cell Cycle 2009, 8 (16), 2664-6. 11. Pan, P. W.; Feldman, J. L.; Devries, M. K.; Dong, A.; Edwards, A. M.; Denu, J. M., Structure and biochemical functions of SIRT6. J Biol Chem 2011, 286 (16), 14575-87. 12. Feldman, J. L.; Baeza, J.; Denu, J. M., Activation of the protein deacetylase SIRT6 by long-chain fatty acids and widespread deacylation by mammalian sirtuins. J Biol Chem 2013, 288 (43), 31350-6. 13. (a) You, W.; Zheng, W.; Weiss, S.; Chua, K. F.; Steegborn, C., Structural basis for the activation and inhibition of Sirtuin 6 by quercetin and its derivatives. Sci Rep 2019, 9 (1), 19176; (b) Tenhunen, J.; Kucera, T.; Huovinen, M.; Kublbeck, J.; Bisenieks, E.; Vigante, B.; Ogle, Z.; Duburs, G.; Dolezal, M.; Moaddel, R.; Lahtela-Kakkonen, M.; Rahnasto- Rilla, M., Screening of SIRT6 inhibitors and activators: A novel activator has an impact on breast cancer cells. Biomed Pharmacother 2021, 138, 111452; (c) You, W.; Rotili, D.; Li, T. M.; Kambach, C.; Meleshin, M.; Schutkowski, M.; Chua, K. F.; Mai, A.; Steegborn, C., Structural Basis of Sirtuin 6 Activation by Synthetic Small Molecules. Angew Chem Int Ed Engl 2017, 56 (4), 1007-1011. 14. (a) Ferrara, G.; Benzi, A.; Sturla, L.; Marubbi, D.; Frumento, D.; Spinelli, S.; Abbotto, E.; Ivaldi, F.; von Holtey, M.; Murone, M.; Nencioni, A.; Uccelli, A.; Bruzzone, S., Sirt6 inhibition delays the onset of experimental autoimmune encephalomyelitis by reducing dendritic cell migration. J Neuroinflammation 2020, 17 (1), 228; (b) Sociali, G.; Magnone, M.; Ravera, S.; Damonte, P.; Vigliarolo, T.; Von Holtey, M.; Vellone, V. G.; Millo, E.; Caffa, I.; Cea, M.; Parenti, M. D.; Del Rio, A.; Murone, M.; Mostoslavsky, R.; Grozio, A.; Nencioni, A.; Bruzzone, S., Pharmacological Sirt6 inhibition improves glucose tolerance in a type 2 diabetes mouse model. FASEB J 2017, 31 (7), 3138-3149; (c) Parenti, M. D.; Grozio, A.; Bauer, I.; Galeno, L.; Damonte, P.; Millo, E.; Sociali, G.; Franceschi, C.; Ballestrero, A.; Bruzzone, S.; Del Rio, A.; Nencioni, A., Discovery of novel and selective SIRT6 inhibitors. J Med Chem 2014, 57 (11), 4796-804. 15. He, B.; Hu, J.; Zhang, X.; Lin, H., Thiomyristoyl peptides as cell-permeable Sirt6 inhibitors. Org Biomol Chem 2014, 12 (38), 7498-502. 16. (a) Spiegelman, N. A.; Zhang, X.; Jing, H.; Cao, J.; Kotliar, I. B.; Aramsangtienchai, P.; Wang, M.; Tong, Z.; Rosch, K. M.; Lin, H., SIRT2 and Lysine Fatty Acylation Regulate the Activity of RalB and Cell Migration. ACS Chem Biol 2019, 14 (9), 2014-2023; (b) Jing, H.; Zhang, X.; Wisner, S. A.; Chen, X.; Spiegelman, N. A.; Linder, M. E.; Lin, H., SIRT2 and lysine fatty acylation regulate the transforming activity of K-Ras4a. Elife 2017, 6; (c) Kosciuk, T.; Price, I. R.; Zhang, X.; Zhu, C.; Johnson, K. N.; Zhang, S.; Halaby, S. L.; Komaniecki, G. P.; Yang, M.; DeHart, C. J.; Thomas, P. M.; Kelleher, N. L.; Fromme, J. C.; Lin, H., NMT1 and NMT2 are lysine myristoyltransferases regulating the ARF6 GTPase cycle. Nat Commun 2020, 11 (1), 1067. 17. (a) Wang, Y.; Yang, J.; Hong, T.; Chen, X.; Cui, L., SIRT2: Controversy and multiple roles in disease and physiology. Ageing Res Rev 2019, 55, 100961; (b) Chen, G.; Huang, P.; Hu, C., The role of SIRT2 in cancer: A novel therapeutic target. Int J Cancer 2020, 147 (12), 3297-3304; (c) Liu, Y.; Zhang, Y.; Zhu, K.; Chi, S.; Wang, C.; Xie, A., Emerging Role of Sirtuin 2 in Parkinson's Disease. Front Aging Neurosci 2019, 11, 372. 18. (a) Zhou, W.; Ni, T. K.; Wronski, A.; Glass, B.; Skibinski, A.; Beck, A.; Kuperwasser, C., The SIRT2 Deacetylase Stabilizes Slug to Control Malignancy of Basal-like Breast Cancer. Cell Rep 2016, 17 (5), 1302-1317; (b) Zhao, D.; Mo, Y.; Li, M. T.; Zou, S. W.; Cheng, Z. L.; Sun, Y. P.; Xiong, Y.; Guan, K. L.; Lei, Q. Y., NOTCH-induced aldehyde dehydrogenase 1A1 deacetylation promotes breast cancer stem cells. J Clin Invest 2014, 124 (12), 5453-65. 19. Fiskus, W.; Coothankandaswamy, V.; Chen, J.; Ma, H.; Ha, K.; Saenz, D. T.; Krieger, S. S.; Mill, C. P.; Sun, B.; Huang, P.; Mumm, J. S.; Melnick, A. M.; Bhalla, K. N., SIRT2 Deacetylates and Inhibits the Peroxidase Activity of Peroxiredoxin-1 to Sensitize Breast Cancer Cells to Oxidant Stress-Inducing Agents. Cancer Res 2016, 76 (18), 5467-78. 20. Jing, H.; Hu, J.; He, B.; Negron Abril, Y. L.; Stupinski, J.; Weiser, K.; Carbonaro, M.; Chiang, Y. L.; Southard, T.; Giannakakou, P.; Weiss, R. S.; Lin, H., A SIRT2-Selective Inhibitor Promotes c-Myc Oncoprotein Degradation and Exhibits Broad Anticancer Activity. Cancer Cell 2016, 29 (5), 767-768. 21. Teng, Y. B.; Jing, H.; Aramsangtienchai, P.; He, B.; Khan, S.; Hu, J.; Lin, H.; Hao, Q., Efficient demyristoylase activity of SIRT2 revealed by kinetic and structural studies. Sci Rep 2015, 5, 8529. 22. Spiegelman, N. A.; Price, I. R.; Jing, H.; Wang, M.; Yang, M.; Cao, J.; Hong, J. Y.; Zhang, X.; Aramsangtienchai, P.; Sadhukhan, S.; Lin, H., Direct Comparison of SIRT2 Inhibitors: Potency, Specificity, Activity-Dependent Inhibition, and On-Target Anticancer Activities. ChemMedChem 2018, 13 (18), 1890-1894. 23. (a) Hong, J. Y.; Jing, H.; Price, I. R.; Cao, J.; Bai, J. J.; Lin, H., Simultaneous Inhibition of SIRT2 Deacetylase and Defatty acylase Activities via a PROTAC Strategy. ACS Med Chem Lett 2020, 11 (11), 2305-2311; (b) Schiedel, M.; Herp, D.; Hammelmann, S.; Swyter, S.; Lehotzky, A.; Robaa, D.; Olah, J.; Ovadi, J.; Sippl, W.; Jung, M., Chemically Induced Degradation of Sirtuin 2 (Sirt2) by a Proteolysis Targeting Chimera (PROTAC) Based on Sirtuin Rearranging Ligands (SirReals). J Med Chem 2018, 61 (2), 482-491. 24. (a) Mellini, P.; Itoh, Y.; Tsumoto, H.; Li, Y.; Suzuki, M.; Tokuda, N.; Kakizawa, T.; Miura, Y.; Takeuchi, J.; Lahtela-Kakkonen, M.; Suzuki, T., Potent mechanism-based sirtuin-2-selective inhibition by an in situ-generated occupant of the substrate-binding site, "selectivity pocket" and NAD(+)-binding site. Chem Sci 2017, 8 (9), 6400-6408; (b) Moniot, S.; Forgione, M.; Lucidi, A.; Hailu, G. S.; Nebbioso, A.; Carafa, V.; Baratta, F.; Altucci, L.; Giacche, N.; Passeri, D.; Pellicciari, R.; Mai, A.; Steegborn, C.; Rotili, D., Development of 1,2,4-Oxadiazoles as Potent and Selective Inhibitors of the Human Deacetylase Sirtuin 2: Structure-Activity Relationship, X-ray Crystal Structure, and Anticancer Activity. J Med Chem 2017, 60 (6), 2344-2360; (c) Rumpf, T.; Schiedel, M.; Karaman, B.; Roessler, C.; North, B. J.; Lehotzky, A.; Olah, J.; Ladwein, K. I.; Schmidtkunz, K.; Gajer, M.; Pannek, M.; Steegborn, C.; Sinclair, D. A.; Gerhardt, S.; Ovadi, J.; Schutkowski, M.; Sippl, W.; Einsle, O.; Jung, M., Selective Sirt2 inhibition by ligand-induced rearrangement of the active site. Nat Commun 2015, 6, 6263; (d) Hong, J. Y.; Price, I. R.; Bai, J. J.; Lin, H., A Glycoconjugated SIRT2 Inhibitor with Aqueous Solubility Allows Structure-Based Design of SIRT2 Inhibitors. ACS Chem Biol 2019, 14 (8), 1802-1810. 25. Outeiro, T. F.; Kontopoulos, E.; Altmann, S. M.; Kufareva, I.; Strathearn, K. E.; Amore, A. M.; Volk, C. B.; Maxwell, M. M.; Rochet, J. C.; McLean, P. J.; Young, A. B.; Abagyan, R.; Feany, M. B.; Hyman, B. T.; Kazantsev, A. G., Sirtuin 2 inhibitors rescue alpha- synuclein-mediated toxicity in models of Parkinson's disease. Science 2007, 317 (5837), 516-9. 26. (a) Cao, J.; Sun, L.; Aramsangtienchai, P.; Spiegelman, N. A.; Zhang, X.; Huang, W.; Seto, E.; Lin, H., HDAC11 regulates type I interferon signaling through defatty acylation of SHMT2. Proc Natl Acad Sci U S A 2019, 116 (12), 5487-5492; (b) Kutil, Z.; Novakova, Z.; Meleshin, M.; Mikesova, J.; Schutkowski, M.; Barinka, C., Histone Deacetylase 11 Is a Fatty-Acid Deacylase. ACS Chem Biol 2018, 13 (3), 685-693; (c) Moreno-Yruela, C.; Galleano, I.; Madsen, A. S.; Olsen, C. A., Histone Deacetylase 11 Is an epsilon-N- Myristoyllysine Hydrolase. Cell Chem Biol 2018, 25 (7), 849-856 e8. 27. (a) Yanginlar, C.; Logie, C., HDAC11 is a regulator of diverse immune functions. Biochim Biophys Acta 2018, 1861 (1), 54-59; (b) Yang, H.; Chen, L.; Sun, Q.; Yao, F.; Muhammad, S.; Sun, C., The role of HDAC11 in obesity-related metabolic disorders: A critical review. J Cell Physiol 2021; (c) Nunez-Alvarez, Y.; Suelves, M., HDAC11: a multifaceted histone deacetylase with proficient fatty deacylase activity and its roles in physiological processes. FEBS J 2021; (d) Bagchi, R. A.; Ferguson, B. S.; Stratton, M. S.; Hu, T.; Cavasin, M. A.; Sun, L.; Lin, Y. H.; Liu, D.; Londono, P.; Song, K.; Pino, M. F.; Sparks, L. M.; Smith, S. R.; Scherer, P. E.; Collins, S.; Seto, E.; McKinsey, T. A., HDAC11 suppresses the thermogenic program of adipose tissue via BRD2. Jci Insight 2018, 3 (15); (e) Sun, L.; Marin de Evsikova, C.; Bian, K.; Achille, A.; Telles, E.; Pei, H.; Seto, E., Programming and Regulation of Metabolic Homeostasis by HDAC11. EBioMedicine 2018, 33, 157-168; (f) Sun, L.; Telles, E.; Karl, M.; Cheng, F.; Luetteke, N.; Sotomayor, E. M.; Miller, R. H.; Seto, E., Loss of HDAC11 ameliorates clinical symptoms in a multiple sclerosis mouse model. Life Sci Alliance 2018, 1 (5), e201800039. 28. (a) Deubzer, H. E.; Schier, M. C.; Oehme, I.; Lodrini, M.; Haendler, B.; Sommer, A.; Witt, O., HDAC11 is a novel drug target in carcinomas. Int J Cancer 2013, 132 (9), 2200-8; (b) Thole, T. M.; Lodrini, M.; Fabian, J.; Wuenschel, J.; Pfeil, S.; Hielscher, T.; Kopp- Schneider, A.; Heinicke, U.; Fulda, S.; Witt, O.; Eggert, A.; Fischer, M.; Deubzer, H. E., Neuroblastoma cells depend on HDAC11 for mitotic cell cycle progression and survival. Cell Death Dis 2017, 8 (3), e2635. 29. (a) Glozak, M. A.; Seto, E., Acetylation/deacetylation modulates the stability of DNA replication licensing factor Cdt1. J Biol Chem 2009, 284 (17), 11446-53; (b) Yuan, Y.; Zhao, K.; Yao, Y.; Liu, C.; Chen, Y.; Li, J.; Wang, Y.; Pei, R.; Chen, J.; Hu, X.; Zhou, Y.; Wu, C.; Chen, X., HDAC11 restricts HBV replication through epigenetic repression of cccDNA transcription. Antiviral Res 2019, 172, 104619; (c) Gong, D.; Zeng, Z.; Yi, F.; Wu, J., Inhibition of histone deacetylase 11 promotes human liver cancer cell apoptosis. Am J Transl Res 2019, 11 (2), 983-990; (d) Wang, W.; Fu, L.; Li, S.; Xu, Z.; Li, X., Histone deacetylase 11 suppresses p53 expression in pituitary tumor cells. Cell Biol Int 2017, 41 (12), 1290-1295. 30. Gao, L.; Cueto, M. A.; Asselbergs, F.; Atadja, P., Cloning and functional characterization of HDAC11, a novel member of the human histone deacetylase family. J Biol Chem 2002, 277 (28), 25748-55. 31. Son, S. I.; Cao, J.; Zhu, C. L.; Miller, S. P.; Lin, H., Activity-Guided Design of HDAC11- Specific Inhibitors. ACS Chem Biol 2019, 14 (7), 1393-1397. 32. (a) Bora-Singhal, N.; Mohankumar, D.; Saha, B.; Colin, C. M.; Lee, J. Y.; Martin, M. W.; Zheng, X.; Coppola, D.; Chellappan, S., Novel HDAC11 inhibitors suppress lung adenocarcinoma stem cell self-renewal and overcome drug resistance by suppressing Sox2. Sci Rep 2020, 10 (1), 4722; (b) Martin, M. W.; Lee, J. Y.; Lancia, D. R., Jr.; Ng, P. Y.; Han, B.; Thomason, J. R.; Lynes, M. S.; Marshall, C. G.; Conti, C.; Collis, A.; Morales, M. A.; Doshi, K.; Rudnitskaya, A.; Yao, L.; Zheng, X., Discovery of novel N-hydroxy-2- arylisoindoline-4-carboxamides as potent and selective inhibitors of HDAC11. Bioorg Med Chem Lett 2018, 28 (12), 2143-2147. 33. Son, S. I.; Su, D.; Ho, T. T.; Lin, H., Garcinol Is an HDAC11 Inhibitor. ACS Chem Biol 2020, 15 (11), 2866-2871. 34. Benz, R., Channel formation by RTX-toxins of pathogenic bacteria: Basis of their biological activity. Biochim Biophys Acta 2016, 1858 (3), 526-37. 35. (a) Baumann, U.; Wu, S.; Flaherty, K. M.; McKay, D. B., Three-dimensional structure of the alkaline protease of Pseudomonas aeruginosa: a two-domain protein with a calcium binding parallel beta roll motif. EMBO J 1993, 12 (9), 3357-64; (b) Baumann, U., Structure-Function Relationships of the Repeat Domains of RTX Toxins. Toxins (Basel) 2019, 11 (11). 36. Benz, R., RTX-Toxins. Toxins (Basel) 2020, 12 (6). 37. Issartel, J. P.; Koronakis, V.; Hughes, C., Activation of Escherichia coli prohaemolysin to the mature toxin by acyl carrier protein-dependent fatty acylation. Nature 1991, 351 (6329), 759-61. 38. Stanley, P.; Packman, L. C.; Koronakis, V.; Hughes, C., Fatty acylation of two internal lysine residues required for the toxic activity of Escherichia coli hemolysin. Science 1994, 266 (5193), 1992-6. 39. Lim, K. B.; Walker, C. R.; Guo, L.; Pellett, S.; Shabanowitz, J.; Hunt, D. F.; Hewlett, E. L.; Ludwig, A.; Goebel, W.; Welch, R. A.; Hackett, M., Escherichia coli alpha-hemolysin (HlyA) is heterogeneously acylated in vivo with 14-, 15-, and 17-carbon fatty acids. J Biol Chem 2000, 275 (47), 36698-702. 40. Hackett, M.; Guo, L.; Shabanowitz, J.; Hunt, D. F.; Hewlett, E. L., Internal lysine palmitoylation in adenylate cyclase toxin from Bordetella pertussis. Science 1994, 266 (5184), 433-5. 41. Hackett, M.; Walker, C. B.; Guo, L.; Gray, M. C.; Van Cuyk, S.; Ullmann, A.; Shabanowitz, J.; Hunt, D. F.; Hewlett, E. L.; Sebo, P., Hemolytic, but not cell-invasive activity, of adenylate cyclase toxin is selectively affected by differential fatty acylation in Escherichia coli. J Biol Chem 1995, 270 (35), 20250-3. 42. (a) Trent, M. S.; Worsham, L. M.; Ernst-Fonberg, M. L., HlyC, the internal protein acyltransferase that activates hemolysin toxin: role of conserved histidine, serine, and cysteine residues in enzymatic activity as probed by chemical modification and site- directed mutagenesis. Biochemistry 1999, 38 (11), 3433-9; (b) Worsham, L. M.; Trent, M. S.; Earls, L.; Jolly, C.; Ernst-Fonberg, M. L., Insights into the catalytic mechanism of HlyC, the internal protein acyltransferase that activates Escherichia coli hemolysin toxin. Biochemistry 2001, 40 (45), 13607-16. 43. Basar, T.; Havlicek, V.; Bezouskova, S.; Hackett, M.; Sebo, P., Acylation of lysine 983 is sufficient for toxin activity of Bordetella pertussis adenylate cyclase. Substitutions of alanine 140 modulate acylation site selectivity of the toxin acyltransferase CyaC. J Biol Chem 2001, 276 (1), 348-54. 44. Greene, N. P.; Crow, A.; Hughes, C.; Koronakis, V., Structure of a bacterial toxin- activating acyltransferase. Proc Natl Acad Sci U S A 2015, 112 (23), E3058-66. 45. Salah Ud-Din, A. I.; Tikhomirova, A.; Roujeinikova, A., Structure and Functional Diversity of GCN5-Related N-Acetyltransferases (GNAT). Int J Mol Sci 2016, 17 (7). 46. Satchell, K. J., MARTX, multifunctional autoprocessing repeats-in-toxin toxins. Infect Immun 2007, 75 (11), 5079-84. 47. Satchell, K. J., Structure and function of MARTX toxins and other large repetitive RTX proteins. Annu Rev Microbiol 2011, 65, 71-90. 48. Cordero, C. L.; Kudryashov, D. S.; Reisler, E.; Satchell, K. J., The Actin cross-linking domain of the Vibrio cholerae RTX toxin directly catalyzes the covalent cross-linking of actin. J Biol Chem 2006, 281 (43), 32366-74. 49. Sheahan, K. L.; Satchell, K. J., Inactivation of small Rho GTPases by the multifunctional RTX toxin from Vibrio cholerae. Cell Microbiol 2007, 9 (5), 1324-35. 50. Zhou, Y.; Huang, C.; Yin, L.; Wan, M.; Wang, X.; Li, L.; Liu, Y.; Wang, Z.; Fu, P.; Zhang, N.; Chen, S.; Liu, X.; Shao, F.; Zhu, Y., N(epsilon)-Fatty acylation of Rho GTPases by a MARTX toxin effector. Science 2017, 358 (6362), 528-531. 51. Woida, P. J.; Satchell, K. J. F., The Vibrio cholerae MARTX toxin silences the inflammatory response to cytoskeletal damage before inducing actin cytoskeleton collapse. Sci Signal 2020, 13 (614). 52. (a) Bros, M.; Haas, K.; Moll, L.; Grabbe, S., RhoA as a Key Regulator of Innate and Adaptive Immunity. Cells 2019, 8 (7); (b) Biro, M.; Munoz, M. A.; Weninger, W., Targeting Rho-GTPases in immune cell migration and inflammation. Br J Pharmacol 2014, 171 (24), 5491-506; (c) Guo, F., RhoA and Cdc42 in T cells: Are they targetable for T cell-mediated inflammatory diseases? Precis Clin Med 2021, 4 (1), 56-61. 53. Liu, W.; Zhou, Y.; Peng, T.; Zhou, P.; Ding, X.; Li, Z.; Zhong, H.; Xu, Y.; Chen, S.; Hang, H. C.; Shao, F., N(epsilon)-fatty acylation of multiple membrane-associated proteins by Shigella IcsB effector to modulate host function. Nat Microbiol 2018. 54. Parsot, C., Shigella type III secretion effectors: how, where, when, for what purposes? Curr Opin Microbiol 2009, 12 (1), 110-6. 55. Wassef, J. S.; Keren, D. F.; Mailloux, J. L., Role of M cells in initial antigen uptake and in ulcer formation in the rabbit intestinal loop model of shigellosis. Infect Immun 1989, 57 (3), 858-63. 56. Zychlinsky, A.; Prevost, M. C.; Sansonetti, P. J., Shigella flexneri induces apoptosis in infected macrophages. Nature 1992, 358 (6382), 167-9. 57. (a) Allaoui, A.; Mounier, J.; Prevost, M. C.; Sansonetti, P. J.; Parsot, C., icsB: a Shigella flexneri virulence gene necessary for the lysis of protrusions during intercellular spread. Mol Microbiol 1992, 6 (12), 1605-16; (b) Ogawa, M.; Yoshimori, T.; Suzuki, T.; Sagara, H.; Mizushima, N.; Sasakawa, C., Escape of intracellular Shigella from autophagy. Science 2005, 307 (5710), 727-31. 58. Pei, J.; Grishin, N. V., The Rho GTPase inactivation domain in Vibrio cholerae MARTX toxin has a circularly permuted papain-like thiol protease fold. Proteins 2009, 77 (2), 413- 9. 59. Towler, D. A.; Adams, S. P.; Eubanks, S. R.; Towery, D. S.; Jackson-Machelski, E.; Glaser, L.; Gordon, J. I., Purification and characterization of yeast myristoyl CoA:protein N-myristoyltransferase. Proc Natl Acad Sci U S A 1987, 84 (9), 2708-12. 60. (a) Farazi, T. A.; Waksman, G.; Gordon, J. I., The biology and enzymology of protein N- myristoylation. J Biol Chem 2001, 276 (43), 39501-4; (b) Yuan, M.; Song, Z. H.; Ying, M. D.; Zhu, H.; He, Q. J.; Yang, B.; Cao, J., N-myristoylation: from cell biology to translational medicine. Acta Pharmacol Sin 2020, 41 (8), 1005-1015. 61. (a) Yang, S. H.; Shrivastav, A.; Kosinski, C.; Sharma, R. K.; Chen, M. H.; Berthiaume, L. G.; Peters, L. L.; Chuang, P. T.; Young, S. G.; Bergo, M. O., N-myristoyltransferase 1 is essential in early mouse development. J Biol Chem 2005, 280 (19), 18990-5; (b) Ntwasa, M.; Aapies, S.; Schiffmann, D. A.; Gay, N. J., Drosophila embryos lacking N- myristoyltransferase have multiple developmental defects. Exp Cell Res 2001, 262 (2), 134-44. 62. (a) Selvakumar, P.; Lakshmikuttyamma, A.; Shrivastav, A.; Das, S. B.; Dimmock, J. R.; Sharma, R. K., Potential role of N-myristoyltransferase in cancer. Prog Lipid Res 2007, 46 (1), 1-36; (b) Berthiaume, L. G.; Beauchamp, E. Epigenetic Silencing of NMT2. 2018. 63. Mousnier, A.; Bell, A. S.; Swieboda, D. P.; Morales-Sanfrutos, J.; Perez-Dorado, I.; Brannigan, J. A.; Newman, J.; Ritzefeld, M.; Hutton, J. A.; Guedan, A.; Asfor, A. S.; Robinson, S. W.; Hopkins-Navratilova, I.; Wilkinson, A. J.; Johnston, S. L.; Leatherbarrow, R. J.; Tuthill, T. J.; Solari, R.; Tate, E. W., Fragment-derived inhibitors of human N-myristoyltransferase block capsid assembly and replication of the common cold virus. Nat Chem 2018, 10 (6), 599-606. 64. Dian, C.; Perez-Dorado, I.; Riviere, F.; Asensio, T.; Legrand, P.; Ritzefeld, M.; Shen, M.; Cota, E.; Meinnel, T.; Tate, E. W.; Giglione, C., High-resolution snapshots of human N- myristoyltransferase in action illuminate a mechanism promoting N-terminal Lys and Gly myristoylation. Nat Commun 2020, 11 (1), 1132. 65. Kosciuk, T.; Lin, H., N-Myristoyltransferase as a Glycine and Lysine Myristoyltransferase in Cancer, Immunity, and Infections. ACS Chem Biol 2020, 15 (7), 1747-1758. 66. Afonina, I. S.; Muller, C.; Martin, S. J.; Beyaert, R., Proteolytic Processing of Interleukin- 1 Family Cytokines: Variations on a Common Theme. Immunity 2015, 42 (6), 991-1004. 67. (a) Priestle, J. P.; Schar, H. P.; Grutter, M. G., Crystallographic refinement of interleukin 1 beta at 2.0 A resolution. Proc Natl Acad Sci U S A 1989, 86 (24), 9667-71; (b) Ren, X.; Gelinas, A. D.; von Carlowitz, I.; Janjic, N.; Pyle, A. M., Structural basis for IL-1alpha recognition by a modified DNA aptamer that specifically inhibits IL-1alpha signaling. Nat Commun 2017, 8 (1), 810. 68. Kim, B.; Lee, Y.; Kim, E.; Kwak, A.; Ryoo, S.; Bae, S. H.; Azam, T.; Kim, S.; Dinarello, C. A., The Interleukin-1alpha Precursor is Biologically Active and is Likely a Key Alarmin in the IL-1 Family of Cytokines. Front Immunol 2013, 4, 391. 69. (a) Stevenson, F. T.; Bursten, S. L.; Fanton, C.; Locksley, R. M.; Lovett, D. H., The 31- kDa precursor of interleukin 1 alpha is myristoylated on specific lysines within the 16-kDa N-terminal propiece. Proc Natl Acad Sci U S A 1993, 90 (15), 7245-9; (b) Bursten, S. L.; Locksley, R. M.; Ryan, J. L.; Lovett, D. H., Acylation of monocyte and glomerular mesangial cell proteins. Myristyl acylation of the interleukin 1 precursors. J Clin Invest 1988, 82 (5), 1479-88. 70. Beuscher, H. U.; Colten, H. R., Structure and function of membrane IL-1. Mol Immunol 1988, 25 (11), 1189-99. 71. Niki, Y.; Yamada, H.; Kikuchi, T.; Toyama, Y.; Matsumoto, H.; Fujikawa, K.; Tada, N., Membrane-associated IL-1 contributes to chronic synovitis and cartilage destruction in human IL-1 alpha transgenic mice. J Immunol 2004, 172 (1), 577-84. 72. (a) Sindhu Kumari, S.; Gupta, N.; Shiels, A.; FitzGerald, P. G.; Menon, A. G.; Mathias, R. T.; Varadaraj, K., Role of Aquaporin 0 in lens biomechanics. Biochem Biophys Res Commun 2015, 462 (4), 339-45; (b) Mathias, R. T.; Rae, J. L.; Baldo, G. J., Physiological properties of the normal lens. Physiol Rev 1997, 77 (1), 21-50. 73. Ball, L. E.; Garland, D. L.; Crouch, R. K.; Schey, K. L., Post-translational modifications of aquaporin 0 (AQP0) in the normal human lens: spatial and temporal occurrence. Biochemistry 2004, 43 (30), 9856-65. 74. Schey, K. L.; Gutierrez, D. B.; Wang, Z.; Wei, J.; Grey, A. C., Novel fatty acid acylation of lens integral membrane protein aquaporin-0. Biochemistry 2010, 49 (45), 9858-65. 75. Ismail, V. S.; Mosely, J. A.; Tapodi, A.; Quinlan, R. A.; Sanderson, J. M., The lipidation profile of aquaporin-0 correlates with the acyl composition of phosphoethanolamine lipids in lens membranes. Biochim Biophys Acta 2016, 1858 (11), 2763-2768. 76. Locksley, R. M.; Killeen, N.; Lenardo, M. J., The TNF and TNF receptor superfamilies: integrating mammalian biology. Cell 2001, 104 (4), 487-501. 77. Stevenson, F. T.; Bursten, S. L.; Locksley, R. M.; Lovett, D. H., Myristyl acylation of the tumor necrosis factor alpha precursor on specific lysine residues. J Exp Med 1992, 176 (4), 1053-62. 78. Jiang, H.; Zhang, X.; Lin, H., Lysine fatty acylation promotes lysosomal targeting of TNF- alpha. Sci Rep 2016, 6, 24371. 79. Zhang, X.; Khan, S.; Jiang, H.; Antonyak, M. A.; Chen, X.; Spiegelman, N. A.; Shrimp, J. H.; Cerione, R. A.; Lin, H., Identifying the functional contribution of the defatty acylase activity of SIRT6. Nat Chem Biol 2016, 12 (8), 614-20. 80. Zhang, X.; Spiegelman, N. A.; Nelson, O. D.; Jing, H.; Lin, H., SIRT6 regulates Ras- related protein R-Ras2 by lysine defatty acylation. Elife 2017, 6. 81. Simanshu, D. K.; Nissley, D. V.; McCormick, F., RAS Proteins and Their Regulators in Human Disease. Cell 2017, 170 (1), 17-33. 82. (a) Martin, T. D.; Samuel, J. C.; Routh, E. D.; Der, C. J.; Yeh, J. J., Activation and involvement of Ral GTPases in colorectal cancer. Cancer Res 2011, 71 (1), 206-15; (b) Guin, S.; Ru, Y.; Wynes, M. W.; Mishra, R.; Lu, X.; Owens, C.; Barn, A. E.; Vasu, V. T.; Hirsch, F. R.; Kern, J. A.; Theodorescu, D., Contributions of KRAS and RAL in non- small-cell lung cancer growth and progression. J Thorac Oncol 2013, 8 (12), 1492-501; (c) Tecleab, A.; Zhang, X.; Sebti, S. M., Ral GTPase down-regulation stabilizes and reactivates p53 to inhibit malignant transformation. J Biol Chem 2014, 289 (45), 31296- 309. 83. Gillingham, A. K.; Munro, S., The small G proteins of the Arf family and their regulators. Annu Rev Cell Dev Biol 2007, 23, 579-611. 84. Goldberg, J., Structural basis for activation of ARF GTPase: mechanisms of guanine nucleotide exchange and GTP-myristoyl switching. Cell 1998, 95 (2), 237-48. 85. Cuthbertson, C. R.; Arabzada, Z.; Bankhead, A., 3rd; Kyani, A.; Neamati, N., A Review of Small-Molecule Inhibitors of One-Carbon Enzymes: SHMT2 and MTHFD2 in the Spotlight. ACS Pharmacol Transl Sci 2021, 4 (2), 624-646. 86. (a) Zheng, H.; Gupta, V.; Patterson-Fortin, J.; Bhattacharya, S.; Katlinski, K.; Wu, J.; Varghese, B.; Carbone, C. J.; Aressy, B.; Fuchs, S. Y.; Greenberg, R. A., A BRISC-SHMT complex deubiquitinates IFNAR1 and regulates interferon responses. Cell Rep 2013, 5 (1), 180-93; (b) Rabl, J.; Bunker, R. D.; Schenk, A. D.; Cavadini, S.; Gill, M. E.; Abdulrahman, W.; Andres-Pons, A.; Luijsterburg, M. S.; Ibrahim, A. F. M.; Branigan, E.; Aguirre, J. D.; Marceau, A. H.; Guerillon, C.; Bouwmeester, T.; Hassiepen, U.; Peters, A.; Renatus, M.; Gelman, L.; Rubin, S. M.; Mailand, N.; van Attikum, H.; Hay, R. T.; Thoma, N. H., Structural Basis of BRCC36 Function in DNA Repair and Immune Regulation. Mol Cell 2019, 75 (3), 483-497 e9; (c) Walden, M.; Tian, L.; Ross, R. L.; Sykora, U. M.; Byrne, D. P.; Hesketh, E. L.; Masandi, S. K.; Cassel, J.; George, R.; Ault, J. R.; El Oualid, F.; Pawlowski, K.; Salvino, J. M.; Eyers, P. A.; Ranson, N. A.; Del Galdo, F.; Greenberg, R. A.; Zeqiraj, E., Metabolic control of BRISC-SHMT2 assembly regulates immune signalling. Nature 2019, 570 (7760), 194-199; (d) Rabl, J., BRCA1-A and BRISC: Multifunctional Molecular Machines for Ubiquitin Signaling. Biomolecules 2020, 10 (11). 87. Linhartova, I.; Bumba, L.; Masin, J.; Basler, M.; Osicka, R.; Kamanova, J.; Prochazkova, K.; Adkins, I.; Hejnova-Holubova, J.; Sadilkova, L.; Morova, J.; Sebo, P., RTX proteins: a highly diverse family secreted by a common mechanism. FEMS Microbiol Rev 2010, 34 (6), 1076-112. 88. Ostolaza, H.; Gonzalez-Bullon, D.; Uribe, K. B.; Martin, C.; Amuategi, J.; Fernandez- Martinez, X., Membrane Permeabilization by Pore-Forming RTX Toxins: What Kind of Lesions Do These Toxins Form? Toxins (Basel) 2019, 11 (6). 89. Knapp, O.; Maier, E.; Polleichtner, G.; Masin, J.; Sebo, P.; Benz, R., Channel formation in model membranes by the adenylate cyclase toxin of Bordetella pertussis: effect of calcium. Biochemistry 2003, 42 (26), 8077-84. 90. (a) Welch, R. A., RTX toxin structure and function: a story of numerous anomalies and few analogies in toxin biology. Curr Top Microbiol Immunol 2001, 257, 85-111; (b) Iwaki, M.; Ullmann, A.; Sebo, P., Identification by in vitro complementation of regions required for cell-invasive activity of Bordetella pertussis adenylate cyclase toxin. Mol Microbiol 1995, 17 (6), 1015-24. 91. (a) Masin, J.; Basler, M.; Knapp, O.; El-Azami-El-Idrissi, M.; Maier, E.; Konopasek, I.; Benz, R.; Leclerc, C.; Sebo, P., Acylation of lysine 860 allows tight binding and cytotoxicity of Bordetella adenylate cyclase on CD11b-expressing cells. Biochemistry 2005, 44 (38), 12759-66; (b) Ludwig, A.; Garcia, F.; Bauer, S.; Jarchau, T.; Benz, R.; Hoppe, J.; Goebel, W., Analysis of the in vivo activation of hemolysin (HlyA) from Escherichia coli. J Bacteriol 1996, 178 (18), 5422-30. 92. (a) Atapattu, D. N.; Czuprynski, C. J., Mannheimia haemolytica leukotoxin binds to lipid rafts in bovine lymphoblastoid cells and is internalized in a dynamin-2- and clathrin- dependent manner. Infect Immun 2007, 75 (10), 4719-27; (b) Frey, J., RTX Toxins of Animal Pathogens and Their Role as Antigens in Vaccines and Diagnostics. Toxins (Basel) 2019, 11 (12). 93. (a) Balashova, N. V.; Shah, C.; Patel, J. K.; Megalla, S.; Kachlany, S. C., Aggregatibacter actinomycetemcomitans LtxC is required for leukotoxin activity and initial interaction between toxin and host cells. Gene 2009, 443 (1-2), 42-7; (b) El-Azami-El-Idrissi, M.; Bauche, C.; Loucka, J.; Osicka, R.; Sebo, P.; Ladant, D.; Leclerc, C., Interaction of Bordetella pertussis adenylate cyclase with CD11b/CD18: Role of toxin acylation and identification of the main integrin interaction domain. J Biol Chem 2003, 278 (40), 38514- 21. 94. Herlax, V.; Mate, S.; Rimoldi, O.; Bakas, L., Relevance of fatty acid covalently bound to Escherichia coli alpha-hemolysin and membrane microdomains in the oligomerization process. J Biol Chem 2009, 284 (37), 25199-210. 95. (a) Osickova, A.; Balashova, N.; Masin, J.; Sulc, M.; Roderova, J.; Wald, T.; Brown, A. C.; Koufos, E.; Chang, E. H.; Giannakakis, A.; Lally, E. T.; Osicka, R., Cytotoxic activity of Kingella kingae RtxA toxin depends on post-translational acylation of lysine residues and cholesterol binding. Emerg Microbes Infect 2018, 7 (1), 178; (b) Levental, I.; Lingwood, D.; Grzybek, M.; Coskun, U.; Simons, K., Palmitoylation regulates raft affinity for the majority of integral raft proteins. Proc Natl Acad Sci U S A 2010, 107 (51), 22050- 4. 96. (a) Bellalou, J.; Sakamoto, H.; Ladant, D.; Geoffroy, C.; Ullmann, A., Deletions affecting hemolytic and toxin activities of Bordetella pertussis adenylate cyclase. Infect Immun 1990, 58 (10), 3242-7; (b) Osickova, A.; Khaliq, H.; Masin, J.; Jurnecka, D.; Sukova, A.; Fiser, R.; Holubova, J.; Stanek, O.; Sebo, P.; Osicka, R., Acyltransferase-mediated selection of the length of the fatty acyl chain and of the acylation site governs activation of bacterial RTX toxins. J Biol Chem 2020, 295 (28), 9268-9280. 97. Aramsangtienchai, P.; Spiegelman, N. A.; He, B.; Miller, S. P.; Dai, L.; Zhao, Y.; Lin, H., HDAC8 Catalyzes the Hydrolysis of Long Chain Fatty Acyl Lysine. ACS Chem Biol 2016, 11 (10), 2685-2692. 98. Young Hong, J.; Cao, J.; Lin, H., Fluorogenic Assays for the Defatty acylase Activity of Sirtuins. Methods Mol Biol 2019, 2009, 129-136. 99. Kutil, Z.; Mikesova, J.; Zessin, M.; Meleshin, M.; Novakova, Z.; Alquicer, G.; Kozikowski, A.; Sippl, W.; Barinka, C.; Schutkowski, M., Continuous Activity Assay for HDAC11 Enabling Reevaluation of HDAC Inhibitors. ACS Omega 2019, 4 (22), 19895- 19904. 100. Lanyon-Hogg, T.; Ritzefeld, M.; Sefer, L.; Bickel, J. K.; Rudolf, A. F.; Panyain, N.; Bineva-Todd, G.; Ocasio, C. A.; O'Reilly, N.; Siebold, C.; Magee, A. I.; Tate, E. W., Acylation-coupled lipophilic induction of polarisation (Acyl-cLIP): a universal assay for lipid transferase and hydrolase enzymes. Chem Sci 2019, 10 (39), 8995-9000. 101. Speers, A. E.; Cravatt, B. F., Profiling enzyme activities in vivo using click chemistry methods. Chem Biol 2004, 11 (4), 535-46. 102. Gai, W.; Li, H.; Jiang, H.; Long, Y.; Liu, D., Crystal structures of SIRT3 reveal that the alpha2-alpha3 loop and alpha3-helix affect the interaction with long-chain acyl lysine. FEBS Lett 2016, 590 (17), 3019-28. 103. Tong, Z.; Wang, M.; Wang, Y.; Kim, D. D.; Grenier, J. K.; Cao, J.; Sadhukhan, S.; Hao, Q.; Lin, H., SIRT7 Is an RNA-Activated Protein Lysine Deacylase. ACS Chem Biol 2017, 12 (1), 300-310. 104. Qasim, H.; McConnell, B. K., AKAP12 Signaling Complex: Impacts of Compartmentalizing cAMP-Dependent Signaling Pathways in the Heart and Various Signaling Systems. J Am Heart Assoc 2020, 9 (13), e016615. 105. Madamanchi, A., Beta-adrenergic receptor signaling in cardiac function and heart failure. Mcgill J Med 2007, 10 (2), 99-104. 106. Bagchi, R. A.; Robinson, E. L.; Hu, T.; Cao, J.; Hong, J. Y.; Tharp, C. A.; Qasim, H.; Gavin, K. M.; Pires da Silva, J.; Major, J. L.; McConnell, B. K.; Seto, E.; Lin, H.; McKinsey, T. A., Reversible lysine fatty acylation of an anchoring protein mediates adipocyte adrenergic signaling. Proc Natl Acad Sci U S A 2022, 119 (7). 107. Ikeda, K.; Yamada, T., UCP1 Dependent and Independent Thermogenesis in Brown and Beige Adipocytes. Front Endocrinol (Lausanne) 2020, 11, 498. CHAPTER 2 HDAC11 COUNTERACTS VIBRIO CHOLERAE MARTX TOXIN BY HYDROLYZING LYSINE FATTY ACYLATION This is a revised version of an in-progress manuscript: Komaniecki, G., Cao, J., Weaver, A., Doerr, T., Seto, E., Lin, H. (2022) HDAC11 Counteracts Vibrio cholerae MARTX toxin by Hydrolyzing Lysine Fatty acylation on Rho-Inactivating Domain. GK did biochemistry and mouse experiments. JC did biochemistry experiments. AW and TD created and supplied V. cholerae strains. ES supplied HDAC11 knockout mice. GK and HL wrote the manuscript. ABSTRACT Vibrio cholerae, a gram-negative bacteria, is the causative agent of cholera which affects an estimated 3-5 million people every year. During infection, V. cholerae secretes a MARTX toxin that is processed in mammalian cells to release several toxin domains including RID, a lysine fatty acyl transferase. In this study we found that RID itself is regulated by self-promoted lysine fatty acylation. RID lysine fatty acylation increases its activity towards its mammalian substrates. HDAC11, a lysine defatty acylase, removes this modification from RID, decreasing its activity towards mammalian substrates. HDAC11 knockout leads to increased levels of V. cholerae in mice following infection and decreased phagocytic uptake in BMDMs. Our study reveals lysine fatty acylation as a reversible post-translational modification that mediates host-pathogen interactions and demonstrates a beneficial role for HDAC11 during V. cholerae infection. INTRODUCTION Vibrio cholerae is a gram negative bacteria and the causative agent of cholera, a diarrheal affliction that affects an estimated 3-5 million people a year and causes an estimated 28,800- 130,000 deaths a year, primarily in the developing world.1 V. cholerae is contracted by drinking contaminated water sources. It colonizes crypts between villi in the small intestine. V. cholerae secrets multiple toxic effector proteins that promote virulence. The best studied of these effectors is cholera toxin (CTX), an ADP-ribosyl transferase that modifies G-proteins leading to higher intercellular cAMP concentration and ion and water efflux from the cell.2 Another effector is the Multifunctional Autoprocessing Repeat in Toxin (MARTX) which is also present in several other bacterial species.3 MARTX is a large peptide that is proteolytically processed into multiple smaller toxin effectors once the holoprotein is inserted into a mammalian cell membrane.4 The V. cholerae MARTX has five major domains. The N and C termini contain characteristic RTX motifs present in all MARTX toxins and are thought to allow translocation of the interior portion of the toxin into host cells while remaining associated to the cell membrane. Once translocated, a cysteine protease domain cleaves the toxin at multiple places releasing an actin crosslinking domain (ACD) that catalyzes an atypical glutamate-lysine linkage between actin monomers,5 an α/β hydrolase with PI3P-specific phospholipase A1 activity,6 and a Rho-inactivating domain (RID). RID is known to disrupt Rho-family GTPases which in turn interferes with actin polymerization. It was recently reported that RID does this by catalyzing lysine fatty acylation on the polybasic region of these GTPases.7 Additionally, IcsB, a toxin from another pathogen, Shigella flexneri, was also found to have lysine fatty acyl transferase activity in human cells, revealing that this activity could be widespread in human pathogens.8 Histone deacetylases (HDACs) are a large family of highly conserved proteins best known for catalyzing the removal of acetyl groups from lysines of histones and other proteins.9 This activity is necessary to regulate gene expression and also other biological processes. However, despite a large degree of sequence similarity, some members of the HDAC family have very little or no deacetylase activity.10 Previous work has shown that one such protein, HDAC11, can efficiently hydrolyze long-chain fatty acyl groups of fourteen (myristoyl) and sixteen (palmitoyl) carbons from lysine residues, collectively known as lysine fatty acylation.11-13 HDAC11 is the newest member of the HDAC family, having only been discovered in 2002.14 HDAC11 protein is abundant in kidney, heart, brain, skeletal muscle, and testis. However, most findings concerning HDAC11 functionality is centered around leukocytes and adipocytes. HDAC11 has diverse effects in immune cells.15 HDAC11 has been shown to localize to the IL- 10 promoter, suppressing expression of this anti-inflammatory cytokine.16 Additionally, it suppresses Treg cells and myeloid-derived suppressor cells. 17, 18 HDAC11 is also highly expressed in neutrophils but HDAC11 knockout also increases the migratory and phagocytotic activity of neutrophils.19 Given HDAC11’s strong activity towards long chain fatty acyl lysine, we hypothesized that this activity could play a role in regulating immune function. Currently, only two proteins, gravin-α and SHMT2, are known HDAC11 defatty acylation substrates. HDAC11 defatty acylation of gravin-α blocks β-AR–mediated thermogenic gene expression.20 Defatty acylation of SHMT2 by HDAC11 suppresses type I interferon signaling, which adds further credence towards the idea that HDAC11 is an immunomodulatory protein.11 If more HDAC11 substrates can be identified, it would help link the interesting enzymatic activity of HDAC11 to its important physiological roles. Given that HDAC11 has been found to have strong lysine defatty acylase activity and the RID toxin has been found to have lysine fatty acyl transferase activity, we wondered whether HDAC11 could work against RID and serve a protective role during V. cholerae infection. We found that, while HDAC11 cannot reverse lysine fatty acylation of RID substrates, HDAC11 can decrease RID activity by removing previously unidentified lysine fatty acylation on RID. In this way, HDAC11 can decrease the toxicity of RID and serve a protective anti-toxin role. This study directly links HDAC11 enzymatic activity to a protective immunological function and identifies an exciting new mechanism of defense centering on an underappreciated protein modification. RESULTS RID fatty acylation is important for activity Previous findings revealed that the RID domain from Vibrio MARTX toxins has lysine fatty acyltransferase activity.7 While some human proteins have been found to contain lysine fatty acylation, this was the first report of an enzyme that can catalyze this modification on human proteins in a physiologically relevant context. To study RID fatty acyl tansferase activity, we utilized metabolic labeling with palmitic acid analog, Alk14, followed by click chemistry using an azide-conjugated fluorophore. When RID and its substrate Rac1 were co-transfected in HEK 293T cells, Rac1 one indeed showed increased fatty acylation (Figure 2.1A). However, we were surprised to find that RID also gave a fluorescent signal, indicating that it was fatty acylated as well (Figure 2.1A). Moreover, catalytic dead mutants of RID show no fatty acylation signal when transfected in HEK 293T cells, suggesting that RID can fatty acylate itself (Figure 2.1B). Given the known activity of RID, we hypothesized that it may be fatty acylated on a lysine residue. To explore this possibility, we mutated every lysine in the RID sequence individually. We found that K2816R mutation abolished RID fatty acylation (Figure 2.1C). Figure 2. 1. RID has lysine fatty acylation. A) HEK 293T cells were co-transfected with RID and its substrate Rac1. Fatty acylation was analyzed with Alk14 labeling followed by click chemistry. Rac1 shows fatty acylation in the presence of RID. RID shows fatty acylation. B) Rac1 was co-overexpressed with catalytically dead mutants of RID to analyze fatty acylation status. Both Rac1 and RID lack fatty acylation demonstrating that catalytic activity is necessary for RID modification as well as substrate modification. C) RID lysine point mutants were generated and assayed for fatty acylation. Mutation of K2816 to R abolishes RID fatty acylation. We next asked if fatty acylation of RID was important for its activity. Indeed, when co- overexpressed with Rac1 or Rac3 in HEK 293T cells, K2816R RID showed decreased activity when compared with WT (Figure 2.2). These results show that RID lysine fatty acylation is important for its lysine fatty acyl transferase activity in cells. Figure 2. 2. RID lysine fatty acylation increases activity towards human substrate. A) Rac1 was co-overexpressed with WT and K2816R (KR) RID to assay activity of RID mutant. K2816R RID shows decreased activity compared to WT. B) Rac3 was co-overexpressed with WT, C3022A (CA), K2816R (KR), or K2816A (KA) RID. KR mutant RID showed decreased activity compared to WT whereas KA did not. Fatty acylation of Rac3 results in an increase in SDS- PAGE mobility resulting in a lower molecular weight band. Lc = antibody light chain. HDAC11 hydrolyzes RID lysine fatty acylation Several previous reports have characterized HDAC11 as a lysine defatty acylase.11-13 We wondered whether HDAC11 could function to oppose RID by removing fatty acyl groups from RID substrates. To test this, we first co-transfected WT or catalytic dead HDAC11 with Rac1 and RID in HEK 293T cells. WT HDAC11 decreased RID-catalyzed Rac1 fatty acylation Figure 2. 3. HDAC11 hydrolyzes RID lysine fatty acylation. A) HEK 293T cells were co- transfected with Rac1, RID, and HDAC11. Rac1 fatty acylation was assayed with Alk14 labeling and showed a decrease with the overexpression of WT HDAC11, but no change with catalytic dead HDAC11 D181A (DA). B) Rac1 was purified from HEK 293T cells co-transfected with RID. Fatty acylated Rac1 was then treated with HDAC11 to assay in vitro activity towards Rac1. No change in Rac1 fatty acylation was detected with the addition of HDAC11 WT or Y304H (YH). C) RID was co-transfected with HDAC11 in HEK 293T cells. RID fatty acylation was reduced in the presence of WT HDAC11 and unchanged in the presence of catalytic dead D181A (DA) HDAC11. D) HDAC11’s activity towards fatty acylated RID was assayed in vitro by treating RID purified from transfected HEK 293T cells. WT, but not catalytic dead Y304H (YH), HDAC11 could decrease RID fatty acylation demonstrating that HDAC11 can hydrolyze RID lysine fatty acylation. E) Biochemical model of HDAC11 counteraction of RID. Graphic made with BioRender. whereas the catalytic dead mutant did not (Figure 2.3A). To confirm that HDAC11 directly hydrolyzed lysine fatty acylation on RID substrates, we treated Rac1 purified from HEK 293T cells co-transfected with RID. However, HDAC11 treatment did not decrease Rac1 fatty acylation in the in vitro treatment, suggesting that HDAC11 might not work directly on RID substrates (Figure 2.3B). We next asked if HDAC11 could hydrolyze lysine fatty acylation on RID and thus decrease RID activity. WT HDAC11 decreased RID fatty acylation when co-transfected in HEK 293T cells whereas catalytic dead HDAC11 had no effect (Figure 2.3C). When purified HDAC11 was added to RID purified from transfected HEK 293T cells, HDAC11 was able to decrease RID fatty acylation, demonstrating that RID is a HDAC11 substrate (Figure 2.3D). These results demonstrate that HDAC11 can decrease RID substrate lysine fatty acylation indirectly by hydrolyzing RID lysine fatty acylation and decreasing RID activity (Figure 2.3E). RID lysine fatty acylation regulates its cell rounding activity By inactivating RhoA-family GTPases, RID blocks downstream actin polymerization, which leads to cell rounding.21 To assay the effect of RID fatty acylation on cell rounding, we transfected HEK 293T cells stably expressing GFP to allow examination of cell shape. When WT RID was transfected a robust cell rounding phenotype was observed, whereas the catalytic dead C3022A mutant had no effect on cell morphology (Figure 2.4A). The K2816R mutant also had no effect on cell morphology, demonstrating that RID lysine fatty acylation is important for this physiological effect. Moreover, WT HDAC11 was able to rescue some of the cell rounding caused by RID when co-transfected whereas catalytic dead HDAC11 was not (Figure 2.4B). These results show that RID lysine fatty acylation is important for its cell rounding capability and that HDAC11 can decrease this phenotype. Figure 2. 4. Lysine fatty acylation regulates RID cell rounding activity. A) RID mutants were transfected into HEK 293T cells stably overexpressing GFP. WT RID caused a clear cell rounding phenotype where as C3022A and K2186R mutants did not. B) WT or catalytic dead (DA) HDAC11 were co-transfected with RID into HEK 293T cells stably overexpressing GFP. WT HDAC11 was able to partially rescue RID-induced cell rounding whereas catalytic dead D181A (DA) HDAC11 was not. Figure 2. 5. Visualization of RID substrate fatty acylation from live V. cholerae. A) HEK 293T cells were transfected with RID substrate Rac3, incubated with Alk14, and infected with V. cholerae for the indicated times. V. cholerae infection caused an increase in Rac3 fatty acylation, demonstrating RID activity. NH2OH treatment showed no decrease in Rac3 fatty acylation ruling out cysteine fatty acylation. B) HEK 293T cells were transfected with the indicated GTPases, incubated with Alk14, and infected with V. cholerae for the indicated times. All GTPases show increased fatty acylation with V. cholerae infection, demonstrating RID activity. Lc = antibody light chain. RID lysine fatty acylation is important for activity in V. cholerae MARTX After discovering that lysine fatty acylation is important for RID activity when transfected, we next wanted to determine if this is relevant in the context of the native MARTX protein. To do this, we first developed an assay to study RID activity during V. cholerae infection. When V. cholerae in log phase growth was added to HEK 293T cell media in the presence of Alk14, RID substrate fatty acylation could be visualized with click chemistry (Figure 2.5A). The fluorescence signal was resistant to NH2OH treatment which can hydrolyze thioesters. This rules out the possibility of the signal being cysteine palmitoylation. RID activity was found to be pronounced towards Rac3 in this context, but other substrates can be modified as well (Figure 2.5B). We then generated mutant V. cholerae strains with point mutations in the endogenous RID sequence. Indeed, the previously reported catalytic dead C3022A mutant strain was unable to increase Rac3 fatty acylation. The K2816R mutant was not catalytic dead, but had decreased activity compared to WT V. cholerae, suggesting that RID fatty acylation at this site is important for activity in the endogenous protein as well (Figure 2.6A). The presence of high molecular weight crosslinked actin from the V. cholerae MARTX actin crosslinking domain was used as control to ensure MARTX translocation is consistent between mutants. We next sought to assay if HDAC11 can reduce RID substrate fatty acylation in this infection model. To do this, we looked at the level of Rac3 fatty acylation in the presence of V. cholerae with HEK 293T cells stably expressing HDAC11 compared to control cells. In the cells stably expressing HDAC11, Rac3 fatty acylation is decreased compared to the control (Figure 2.6B). This observation was true for all tested RhoA-family GTPases tested but was most noticeable with Rac3 (Figure 2.6C). HDAC11 KO mice are more susceptible to V. cholera infection To test the physiological importance of HDAC11 during V. cholerae infection, we infected WT and HDAC11 KO mice and tracked the number of bacteria in fecal pellets (FPs). Mice native gut microbiome was first cleared with streptomycin and bacteria were administered via oral gavage (Figure 2.7A).22 Overall activity level and body weight change was consistent between experimental groups (Figure 2.7B). Bacterial colonization was tracked over two weeks during which WT mice were able to decrease the bacterial load more effectively (Figure 2.7C- D). Indeed, some WT mice were able to completely clear the V. cholerae by the end of the experiment (Figure 2.7D). Figure 2. 6. RID lysine fatty acylation affects activity in V. cholerae. A) Rac3 fatty acylation was assayed with RID point mutants V. cholerae strains. The RID catalytic dead C3022A (CA) strain showed no activity in Rac3 fatty acylation. The RID K2816R mutant (KR) showed decreased activity towards in Rac3 fatty acylation whereas the K2816A mutant was as active as WT. Actin crosslinking was used as a control to demonstrate equal MARTX translocation between mutants. B) Rac3 fatty acylation was assayed in HEK 293T cells stably overexpressing HA-HDAC11. Rac3 fatty acylation was decreased with HDAC11 overexpression. Actin crosslinking was used as a control for equal MARTX translocation between cell lines. C) Fatty acylation of RhoA-family GTPases was assayed for 6 hours in HEK 293T cells stably overexpressing HA-HDAC11. HDAC11 overexpression resulted in decreased fatty acylation for all RID substrates. Figure legend on following page. Figure 2. 7. HDAC11 is beneficial during V. cholerae infection. A) Workflow of mouse V. cholerae infection protocol. Following 24 hour administration of streptomycin in the water source to eliminate mouse native gut microbiome, V. cholerae was administered via oral gavage. Mice were tracked for two weeks and fecal pellets isolated to count bacterial colonization. B) Average weight change of mice experimental groups over the course of two week infection. C) Quantification of bacteria isolated from FPs of WT and HDAC11 mice challenged with PBS or V. cholerae. D) Bacterial growth from serial dilutions of FP homogenates from WT and HDAC11 mice infected with V. cholerae. Values at left correspond to the fold dilution from FP homogenates. We next asked what the mechanism is by which HDAC11 promotes clearance of V. cholerae. Actin polymerization plays a key function during phagocytosis so we sought to determine if HDAC11 could promote phagocytic uptake of V. cholerae. We engineered GFP expressing V. cholerae strains and administered them to bone marrow derived macrophages (BMDMs) isolated from WT and HDAC11 KO mice. We then used fluorescence microscopy to count phagocytosed V. cholerae (Figure 2.8A). HDAC11 KO BMDMs phagocytosed less WT V. cholerae than the WT BMDMs (Figure 2.8B). Additionally, V. cholerae with a catalytic dead RID (C3022A) has higher levels of phagocytosis. Together, this data suggests that HDAC11 contributes to defense of V. cholerae infection by counteracting RID activity to enhance phagocytosis. DISCUSSION In this study, we discovered a unique role for HDAC11 in fighting V. cholerae infection. HDAC11 has previously been connected to the immune system. Several reports have shown a Figure 2. 8. HDAC11 and RID affect V. cholerae phagocytosis by BMDMs. A) Representative fluorescence microscopy images of WT and HDAC11 BMDMs challenged with the indicated strains of V. cholerae expressing GFP. Nuclei are stained with DAPI. B) Quantification of A. At least 700 BMDMs were quantified for each sample. pro-inflammatory role for HDAC11 as it suppresses IL-10 expression and Treg cell development and regulates the migratory and phagocytotic activity of neutrophils.16, 17, 19 However, HDAC11 also suppresses type I interferon signaling via the defatty acylation of SHMT2, which suggests an anti-inflammatory role.11 Furthermore, HDAC11 knockout in mice has been shown to have several beneficial effects, including the resistance to high-fat diet induced weight gain, increased brown adipose tissue formation, and decreased inflammation.23-25 These contrasting observations make one wonder what the major physiological benefit of having HDAC11 is. Our study suggests that HDAC11 could help to fight bacteria that modulate lysine fatty acylation. This could potentially also help explain why HDAC11 has pro-inflammation roles but at the same time suppresses type I interferon signaling, which is mainly an anti-viral response.11, 16, 17, 19 We identified a previously unknown lysine fatty acylation site on the MARTX RID that is important for its activity. The modified lysine is near the active site based on the V. vulnificus RID crystal structure and is highly conserved in RIDs from multiple species (Figure 2.9).26 Curiously, K2816A RID is more active than K2816R for both transiently overexpressed RID and endogenous bacterial RID (Figures 2.2B, 2.6A). Arginine more closely resembles a free lysine sidechain which could explain this discrepancy and the previously reported cell rounding activity of K2816A RID.26 The lack of fatty acylation on catalytically dead RID mutants suggest that this enzyme is self-modified.7 By catalyzing its own lysine fatty acylation, RID increases its activity towards its human protein substrates. RID has been shown to inhibit actin polymerization. Actin remodeling is an important part of phagocytosis. We found that V. cholerae with an inactive RID have higher levels of phagocytosis by BMDMs and that HDAC11 KO decreases phagocytosis. However, phagocytosis of the K2816R mutant resembles the WT bacteria. These physiological results are not in line with the biochemical data, raising the possibility of an alternative biochemical mechanism during phagocytosis. One potential explanation is that HDAC11 can remove KFA from additional unexplored RID substrates to directly counteract the RID. ΔMARTX bacteria have increased phagocytosis compared to WT bacteria and HDAC11 KO increases phagocytosis of this strain (Figure 2.8B). This suggests that HDAC11 has a unique role during V. cholerae infection related to the MARTX toxin. HDAC11 could be activated by MARTX to be pro-phagocytic in opposition to RID but be could otherwise suppress phagocytosis. Additional studies of HDAC11’s role in macrophages and phagocytosis of other bacteria could illuminate a novel physiological activity. The V. cholerae MARTX also contains an ACD that crosslinks actin and suppresses actin dynamics.5 This may mask the anti-phagocytic role of RID when ACD is delivered in conjunction. However, some clinically reported V. cholerae strains contain a MARTX protein which does not have ACD which may make RID regulation even more impactful.27 RID has also been found to play an anti-inflammatory role during infection so HDAC11 and RID lysine fatty acylation could be regulating this function as well.28 Figure 2. 9. K2816 is near active site of V. vulnificus RID. The structure of the V. vulnificus RID domain has been solved. K2816 is near the putative active site residues C3022A (mutated to an alanine when solving the crystal structure) and H2782. Lysine fatty acylation has been shown to be important in the regulation of bacterial toxins, including the secretion and/or toxicity of many RTX toxins.29, 30 In mammals, lysine fatty acylation has been increasingly recognized as a relevant regulatory post-translational modification.31 Although the mammalian lysine fatty acyl transferases remain enigmatic for most human lysine fatty acylated proteins,32 several HDAC family members have been shown to have lysine defatty acylase activity, including the NAD-dependent sirtuins.11, 32-36 In this study, we initially sought to investigate whether the fatty acylation of RID substrates can be reversed by HDAC11, but instead identified a novel mode of regulation for RID itself. By demonstrating that HDAC11 regulates RID activity through lysine defatty acylation, this study provides a novel mechanism of host-pathogen interaction via protein lysine fatty acylation. In addition to RID, the Shigella flexneri toxin IcsB also catalyzes lysine fatty acylation. It would interesting to investigate whether IcsB is similarly regulated and what role HDAC11 could play during S. flexneri infection. Pathogenic bacteria use many protein posttranslational modifications to disrupt host immune defense mechanism.37 In most cases, it is not clear whether the host can counteract these pathogen-mediated PTMs. Lysine fatty acylation is a clear example of pathogen-mediated PTM that mammalian host is able to counteract with HDACs. METHODS Reagents. Anti-Flag affinity gel (catalog # A2220) and anti-Flag antibody conjugated with horseradish peroxidase (A8592) were purchased from Sigma-Aldrich. Antibodies against β- actin (sc-4777), and HA (sc516102) were purchased from Santa Cruz Biotechnology. Protease inhibitor cocktail was purchased from Sigma-Aldrich. Streptavidin agarose beads, ECL plus western blotting detection reagent and universal nuclease for cell lysis were purchased from Thermo Scientific Pierce. Polyethylenimine (PEI) was purchased from Polysciences (24765). Cloning and mutagenesis. Full length human RhoA-family GTPases with N-terminal Flag tag was cloned into pCMV5 vector for transient overexpression. Full length human HDAC11 and V. cholerae RID with C-terminal Flag tag was cloned into pCMV4a vector for transient overexpression. For generation of stably overexpressing HEK 293T cells lines, HDAC11 with C-terminal Flag tag was cloned into pCDH vector. Cell culture. HEK 293T cells were cultures in DMEM media (11965–092, Gibco) with 10% calf serum (C8056, Sigma-Aldrich). For stable overexpression, lentivirus was generated by co-transfection of pCDH containing the desired gene, pCMV-dR8.2, and pMD2.G plasmids into HEK293T cells. The cell medium was collected 48 h after transfection and used to infect cells of interest. After 72 h, infected cells were further treated with 1.5 mg/mL puromycin to select for stable overexpression cells. Empty pCDH vector was used as a negative control. Detection of fatty acylation by in-gel fluorescence. Indicated plasmids were transfected into HEK 293T cells using PEI transfection reagent following the manufacturer's protocol. The pCMV-Tag 4a empty vector was used as the negative control. After overnight transfection, cells were treated with 50 μM Alk14 for 6 hours. The cells were washed twice with ice cold PBS and collected by centrifugation at 1000 g for 5 min. Cells were then lysed in 1 mL 1% NP-40 lysis buffer (25 mM Tris-HCl, pH 7.8, 150 mM NaCl, 10% glycerol and 1% NP-40) with protease inhibitor cocktail (1:100 dilution) at 4 °C for 30 min. After centrifugation at 17,000 g for 30 min, the supernatant was collected and protein concentration was determined with Bradford assay (23200, Thermo Fisher). Lysates at a concentration of 0.5-2 mg/mL was incubated with 20 μL of anti-Flag affinity gel per mg of protein at 4 °C for 2 h. The affinity gel was washed three times with washing buffer (25 mM Tris-HCl, pH 7.8, 150 mM NaCl, 0.2% NP-40). Beads were drained with gel loading tips and resuspended in 20 μL of washing buffer. TAMRA-N3 (47130, Lumiprobe, 1 μL of 2 mM solution in DMF), Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (T2993, Tcichemicals, 1 μL of 10 mM solution in DMF), CuSO4 (1 μL of 40 mM solution in H2O) and Tris(2-carboxyethyl)phosphine (TCEP) (580560, Millipore, 1 μL of 40 mM solution in H2O) were added into the reaction mixture in the order listed. The click chemistry reaction was allowed to proceed at room temperature for 30 min. The reaction was quenched by adding 10 μL of 6x SDS loading dye to achieve a final concentration of ~2x and then boiled at 95 ˚C for 5 min. For samples that need to be treated with hydroxylamine to remove cysteine fatty acylation, 10 μL of the quenched reaction was treated with 2 uL of 4 M hydroxylamine (438227, Sigma, pH 7.4) and boiled at 95 ˚C for another 5 min. The samples were then resolved by 12% SDS-PAGE. The gel was then incubated with destaining buffer (50% CH3OH, 40% water and 10% acetic acid) by shaking 2-8 hours at 4˚C and then incubated in water for 2 hours. The gel was scanned to record the rhodamine fluorescence signal using a Typhoon 7000 Variable Mode Imager (GE Healthcare Life Sciences). After scanning, the gel was stained with Coomassie Brilliant Blue (CBB) (B7920, Sigma) to check for protein loading. Expression and purification of human HDAC11 from HEK 293T cells. HDAC11 plasmids (WT, and Y304H) were transfected into HEK 293T cells using PEI transfection reagent following the manufacturer's protocol. The pCMV-Tag 4a empty vector was used as the negative control. The cells were washed twice with ice cold PBS and collected by centrifugation at 1000 g for 5 min. Cells were then lysed in 1% NP-40 lysis buffer (25 mM Tris-HCl, pH 7.8, 150 mM NaCl, 10% glycerol and 1% NP-40) with protease inhibitor cocktail (1:100 dilution) at 4 °C for 30 min. After centrifugation at 17,000 g for 30 min, the supernatant was collected and protein concentration was determined with Bradford assay (23200, Thermo Fisher). Lysates at a concentration of ~2 mg/mL was incubated with 20 μL of anti-flag affinity gel per mg of protein at 4 °C for 2 h. The affinity gel was washed three times with washing buffer (25 mM Tris-HCl, pH 7.8, 150 mM NaCl, 0.2% NP-40) and then eluted three times with 300 μM of triple Flag peptide (F4799, Sigma-Aldrich) (dissolved in 25 mM Tris-HCl, pH 7.4, 150 mM NaCl, and 10% glycerol). The elutions were combined and concentrated with 10 kDa centrifugal spin filters according to the manufacturer’s directions (UFC501096, Millipore). Eluted HDAC11 was used in the in vitro assay. In vitro defatty acylation assay. Plasmids of the indicated HDAC11 substrates were transfected into HEK 293T cells using PEI transfection reagent. After 24 h, the cells were treated with 50 μM of Alk14 for 6 h. Cells were harvested, and flag-tagged protein was affinity purified using anti-flag affinity gel as described above. After washing the flag affinity gel three times with washing buffer (25 mM Tris-HCl, pH 7.4, 150 mM NaCl, 0.2% NP-40), 50 μL of triple flag peptide solution (300 μM in 25 mM Tris-HCl, pH 7.4, 150 mM NaCl) was added to the affinity gel and incubated with rocking for 30 min at 4 °C to elute indicated protein. The elution step was repeated two times and elutions were pooled. 50 μL of purified substrate was incubated with 300 μmol of HDAC11 purified from HEK 293T cells or buffer for 1 h at 37 °C. After incubation, protein was precipitated by adding 200 μL of cold methanol, 75 μL of cold chloroform, and 150 μL of cold water to each sample. After vortexing, the samples were centrifuged at 17,000 g for 20 min at 4 °C. The supernatant was gently removed by pipetting, and 1 mL cold methanol was added to the pellet. The samples were again vortexed and spun down at 17,000 × g for 10 min at 4 °C. Then methanol was removed, and the wash step was repeated. After the second methanol wash, the protein pellets were air-dried for 20 min. Protein was then re-solubilized in 20 μL of 4% SDS buffer (50 mM triethanolamine at pH 7.4, 150 mM NaCl, 4% (w/v) SDS) followed by the click chemistry reaction using TAMRA-N3 described above. The samples were resolved by 12% SDS-PAGE and TAMRA fluorescence was scanned as indicated above. Generation of mutant V. cholerae strains. A summary of all strains, plasmids, and primers used in this study can be found in Tables S1-S2. The V. cholerae strains in this study are a derivative of streptomycin-resistant biovar O1 El Tor E7946.38 E. coli DH5α λpir was used for general cloning, while E. coli SM10 was used for conjugation into V. cholerae.39 Plasmids were constructed using Gibson assembly.40 Chimeric DNA fragments used for chitin-inducible natural transformation were generated via splicing by overlap extension (SOE) PCR.41, 42 Chromosomal mutations were validated by Sanger sequencing of the relevant locus. RtxA was inactivated by replacing the genetic region encoding the RID, residues 2778- 3108, with a trimethoprim resistance cassette via chitin-inducible natural transformation.42, 43 Briefly, V. cholerae was grown to early stationary phase (OD600 0.8-1) in LB, pelleted, then resuspended to an OD600 of 1.0 in artificial seawater (Amazon B00NQH210G). Cells were diluted to OD600 0.1 in 1 ml artificial seawater containing 80 mg autoclaved chitin (Sigma 1398-61-4) in a 1.5 ml tube and incubated without agitation at 30°C for 24 hrs. At this time, 550 μl of supernatant were removed and replaced with 300 μl artificial seawater containing ~1 ng of SOE PCR amplified trimethoprim resistance cassette sandwiched by 2.3kb of the up and downstream flanking sequence of the RID-encoding region. After DNA addition, the cell and chitin mixture was very gently inverted to mix and incubated at 30°C for 24 hrs. Next, the cell and chitin mixture was vortexed vigorously with 1 ml LB, transferred to a culture tube, and incubated at 37°C for 3 hrs before selection on LB plates containing 50 μg/ml trimethoprim. The RtxA RID K2816A mutant was similarly generated by co-transformation of a SOE PCR product restoring the RID-encoding region with the K96A mutation and a secondary SOE PCR amplified chloramphenicol resistance cassette targeting pseudogene vc1807 as a positive selection for DNA uptake. Transformants were selected on LB containing 5 μg/mL chloramphenicol, then screened for loss of trimethoprim resistance. The RtxA RID K2816R mutant was generated using the pCVD442 ampR/sacB allelic exchange system.44 The RID-encoding region of rtxA (plus 500bp flanking either side of the RID-encoding region) was amplified in two parts such that the K96R mutation could be introduced by primers during amplification and stitched together during Gibson assembly into the SmaI site of the suicide vector pCVD442. Conjugation into V. cholerae from E. coli was performed by mixing and pelleting equal volumes of recipient V. cholerae and SM10 donor LB overnight cultures, spotting the mixed pellet onto LB followed by incubation at 37°C for 3 hrs. The first round of selection was performed on LB + streptomycin (200 μg ml-1) + ampicillin (100 μg ml-1) at 30°C followed by counter selection on salt-free LB + 10% sucrose + streptomycin at room temperature. To achieve constitutive expression of GFP, gfp-mut3 under the lac promoter was integrated into the native lacZ locus (vc2338) using pCVD442 derivative pJZ111.45 As V. cholerae encodes no native lacI, expression from Plac is constitutive. Selection for double crossover events was performed as described for pCVD442. Detection of V. cholerae induced fatty acylation. Indicated V. cholerae strains were grown in 5 mL LB overnight with shaking at 37 °C. The next day, V. cholerae strains were diluted 1:100 in fresh LB and grown with shaking at 37 °C to log phase (OD600 = 0.6 – 1.0). After diluting the overnight V. cholerae cultures, 50 M of Alk14 was added to mammalian cells and incubated for 2 hours. During this 2 hour period, once V. cholerae strains reached desired OD600, different strains were normalized with LB to have an equal OD600 of at least 0.6. Then, 1 mL of the V. cholerae culture was centrifuged at 17,000 G for 1 min. LB was carefully pipetted off and the bacterial pellet was resuspended with 1 mL of sterile PBS. After mammalian cells were incubated with Alk14 for 2 hours, the V. cholerae in PBS was diluted 1:500 into cell culture media and added to mammalian cells for 6 hours or otherwise indicated. No antibiotics was added in the cell culture media to allow good V. cholerae growth. After 6 hours, cells were collected and the fatty acylation of target proteins was assayed as indicated above. Mouse inoculation by V. cholerae. The procedure was adapted from work previously reported.22 Indicated V. cholerae strains were grown in 5 mL LB overnight with shaking at 37 °C. The same day (24 hours before inoculation), WT or HDAC11 knockout C57BL/6 mice were given water with 5 mg/mL streptomycin ad libitum to clear out the native gut microbiome. The next day, V. cholerae strains were diluted 1:100 in fresh LB and grown with shaking at 37 °C to log phase (OD600 = 0.6 – 1.0). Bacteria were pelleted, washed twice with sterile PBS, and resuspended to an OD600 of 6.5. Bacteria were administered via oral gavage. Mice were first weighed then given 10 μL of bacterial suspension per gram mouse weight (e.g. 210 μL for a 21 g mouse) corresponding to approximately 5 × 106 bacteria per gram of mouse. Mice were observed over the course of the experiment to track weight change and overall fitness and water was changed to new streptomycin every 48 hours. To track bacterial colonization, one average sized (~0.5 cm) fresh fecal (FP) pellet was collected and placed in sterile PBS. The FP was homogenized on a TissueLyser LT (Qiagen, without use of steel beads) for 5 min at 50/s and used to make serial dilutions in sterile PBS. Dilutions were spread on LB agar plates with 100 μg/mL streptomycin to enumerate CFU. Determination of phagocytic V. cholerae uptake by BMDMs. Bone marrow derived macrophages were isolated from WT or HDAC11 knockout C57BL/6 mice. Monocytes isolated from the bone marrow were cultured for 6 days (changing media after 3 days) in DMEM 10% FBS supplemented with GlutaMAX (Thermo Fisher), Pen/Strep (Gibco), and 20 ng/mL mouse M-CSF (Thermo Fisher). 5 × 105 BMDMs were then plated onto a 35 mm imaging dish (Mattek). Meanwhile, indicated GFP-expressing V. cholerae strains were grown in 5 mL LB overnight with shaking at 37 °C. The next day, V. cholerae strains were diluted 1:100 in fresh LB and grown with shaking at 37 °C to log phase (OD600 = 0.6 – 1.0). Bacteria were pelleted and washed twice with sterile PBS and resuspended to OD600 = 0.6. 50 μL of bacteria were added to the BMDM cells for an MOI of approximately 30. BMDMs were incubated with bacteria at 37 °C for 1 hour to allow for internalization. Extracellular bacteria were then washed away with 1 mL PBS (five times), fixed with 3.7% formaldehyde, washed 3 times with 1 mL PBS, and labeled with DAPI fluoromount (Thermo Fisher). Image acquisition and GFP puncta enumeration was carried out on Cytation5 (Biotek). ACKNOWLEDGEMENTS This work is supported in part by a grant from NIH/NIAID R01AI153110. REFERENCES 1. Harris, J. B.; LaRocque, R. C.; Qadri, F.; Ryan, E. T.; Calderwood, S. B., Cholera. Lancet 2012, 379 (9835), 2466-2476. 2. Muanprasat, C.; Chatsudthipong, V., Cholera: pathophysiology and emerging therapeutic targets. Future Med Chem 2013, 5 (7), 781-98. 3. Satchell, K. J., Structure and function of MARTX toxins and other large repetitive RTX proteins. Annu Rev Microbiol 2011, 65, 71-90. 4. Fullner, K. J.; Boucher, J. C.; Hanes, M. A.; Haines, G. K., 3rd; Meehan, B. M.; Walchle, C.; Sansonetti, P. J.; Mekalanos, J. J., The contribution of accessory toxins of Vibrio cholerae O1 El Tor to the proinflammatory response in a murine pulmonary cholera model. J Exp Med 2002, 195 (11), 1455-62. 5. Kudryashov, D. S.; Durer, Z. A.; Ytterberg, A. J.; Sawaya, M. R.; Pashkov, I.; Prochazkova, K.; Yeates, T. O.; Loo, R. R.; Loo, J. A.; Satchell, K. J.; Reisler, E., Connecting actin monomers by iso-peptide bond is a toxicity mechanism of the Vibrio cholerae MARTX toxin. Proc Natl Acad Sci U S A 2008, 105 (47), 18537-42. 6. Agarwal, S.; Kim, H.; Chan, R. B.; Agarwal, S.; Williamson, R.; Cho, W.; Paolo, G. D.; Satchell, K. J., Autophagy and endosomal trafficking inhibition by Vibrio cholerae MARTX toxin phosphatidylinositol-3-phosphate-specific phospholipase A1 activity. Nat Commun 2015, 6, 8745. 7. Zhou, Y.; Huang, C.; Yin, L.; Wan, M.; Wang, X.; Li, L.; Liu, Y.; Wang, Z.; Fu, P.; Zhang, N.; Chen, S.; Liu, X.; Shao, F.; Zhu, Y., N(epsilon)-Fatty acylation of Rho GTPases by a MARTX toxin effector. Science 2017, 358 (6362), 528-531. 8. Liu, W.; Zhou, Y.; Peng, T.; Zhou, P.; Ding, X.; Li, Z.; Zhong, H.; Xu, Y.; Chen, S.; Hang, H. C.; Shao, F., N(epsilon)-fatty acylation of multiple membrane-associated proteins by Shigella IcsB effector to modulate host function. Nat Microbiol 2018. 9. Yang, X. J.; Seto, E., The Rpd3/Hda1 family of lysine deacetylases: from bacteria and yeast to mice and men. Nat Rev Mol Cell Biol 2008, 9 (3), 206-18. 10. Seto, E.; Yoshida, M., Erasers of histone acetylation: the histone deacetylase enzymes. Cold Spring Harb Perspect Biol 2014, 6 (4), a018713. 11. (a) Cao, J.; Sun, L.; Aramsangtienchai, P.; Spiegelman, N. A.; Zhang, X.; Huang, W.; Seto, E.; Lin, H., HDAC11 regulates type I interferon signaling through defatty acylation of SHMT2. Proc Natl Acad Sci U S A 2019, 116 (12), 5487-5492; (b) Moreno-Yruela, C.; Galleano, I.; Madsen, A. S.; Olsen, C. A., Histone Deacetylase 11 Is an epsilon-N- Myristoyllysine Hydrolase. Cell Chem Biol 2018, 25 (7), 849-856 e8; (c) Kutil, Z.; Novakova, Z.; Meleshin, M.; Mikesova, J.; Schutkowski, M.; Barinka, C., Histone Deacetylase 11 Is a Fatty-Acid Deacylase. ACS Chem Biol 2018, 13 (3), 685-693. 12. Gao, L.; Cueto, M. A.; Asselbergs, F.; Atadja, P., Cloning and functional characterization of HDAC11, a novel member of the human histone deacetylase family. J Biol Chem 2002, 277 (28), 25748-55. 13. Yanginlar, C.; Logie, C., HDAC11 is a regulator of diverse immune functions. Biochim Biophys Acta 2018, 1861 (1), 54-59. 14. Villagra, A.; Cheng, F.; Wang, H. W.; Suarez, I.; Glozak, M.; Maurin, M.; Nguyen, D.; Wright, K. L.; Atadja, P. W.; Bhalla, K.; Pinilla-Ibarz, J.; Seto, E.; Sotomayor, E. M., The histone deacetylase HDAC11 regulates the expression of interleukin 10 and immune tolerance. Nat Immunol 2009, 10 (1), 92-100. 15. (a) Huang, J.; Wang, L.; Dahiya, S.; Beier, U. H.; Han, R.; Samanta, A.; Bergman, J.; Sotomayor, E. M.; Seto, E.; Kozikowski, A. P.; Hancock, W. W., Histone/protein deacetylase 11 targeting promotes Foxp3+ Treg function. Sci Rep 2017, 7 (1), 8626; (b) Sahakian, E.; Powers, J. J.; Chen, J.; Deng, S. L.; Cheng, F.; Distler, A.; Woods, D. M.; Rock-Klotz, J.; Sodre, A. L.; Youn, J. I.; Woan, K. V.; Villagra, A.; Gabrilovich, D.; Sotomayor, E. M.; Pinilla-Ibarz, J., Histone deacetylase 11: A novel epigenetic regulator of myeloid derived suppressor cell expansion and function. Mol Immunol 2015, 63 (2), 579-85. 16. Sahakian, E.; Chen, J.; Powers, J. J.; Chen, X.; Maharaj, K.; Deng, S. L.; Achille, A. N.; Lienlaf, M.; Wang, H. W.; Cheng, F.; Sodre, A. L.; Distler, A.; Xing, L.; Perez- Villarroel, P.; Wei, S.; Villagra, A.; Seto, E.; Sotomayor, E. M.; Horna, P.; Pinilla-Ibarz, J., Essential role for histone deacetylase 11 (HDAC11) in neutrophil biology. J Leukoc Biol 2017, 102 (2), 475-486. 17. Bagchi, R. A.; Robinson, E. L.; Hu, T.; Cao, J.; Hong, J. Y.; Tharp, C. A.; Qasim, H.; Gavin, K. M.; Pires da Silva, J.; Major, J. L.; McConnell, B. K.; Seto, E.; Lin, H.; McKinsey, T. A., Reversible lysine fatty acylation of an anchoring protein mediates adipocyte adrenergic signaling. Proc Natl Acad Sci U S A 2022, 119 (7). 18. Satchell, K. J., MARTX, multifunctional autoprocessing repeats-in-toxin toxins. Infect Immun 2007, 75 (11), 5079-84. 19. Nygren, E.; Li, B. L.; Holmgren, J.; Attridge, S. R., Establishment of an adult mouse model for direct evaluation of the efficacy of vaccines against Vibrio cholerae. Infect Immun 2009, 77 (8), 3475-84. 20. (a) Bagchi, R. A.; Ferguson, B. S.; Stratton, M. S.; Hu, T.; Cavasin, M. A.; Sun, L.; Lin, Y.-H.; Liu, D.; Londono, P.; Song, K.; Pino, M. F.; Sparks, L. M.; Smith, S. R.; Scherer, P. E.; Collins, S.; Seto, E.; McKinsey, T. A., HDAC11 Suppresses the Thermogenic Program of Adipose Tissue via BRD2. bioRxiv 2018; (b) !!! INVALID CITATION !!! {}; (c) Sun, L.; Telles, E.; Karl, M.; Cheng, F.; Luetteke, N.; Sotomayor, E. M.; Miller, R. H.; Seto, E., Loss of HDAC11 ameliorates clinical symptoms in a multiple sclerosis mouse model. Life Sci Alliance 2018, 1 (5), e201800039. 21. Ahrens, S.; Geissler, B.; Satchell, K. J., Identification of a His-Asp-Cys catalytic triad essential for function of the Rho inactivation domain (RID) of Vibrio cholerae MARTX toxin. J Biol Chem 2013, 288 (2), 1397-408. 22. Satchell, K. J. F., Multifunctional-autoprocessing repeats-in-toxin (MARTX) Toxins of Vibrios. Microbiol Spectr 2015, 3 (3). 23. Woida, P. J.; Satchell, K. J. F., The Vibrio cholerae MARTX toxin silences the inflammatory response to cytoskeletal damage before inducing actin cytoskeleton collapse. Sci Signal 2020, 13 (614). 24. (a) Linhartova, I.; Bumba, L.; Masin, J.; Basler, M.; Osicka, R.; Kamanova, J.; Prochazkova, K.; Adkins, I.; Hejnova-Holubova, J.; Sadilkova, L.; Morova, J.; Sebo, P., RTX proteins: a highly diverse family secreted by a common mechanism. FEMS Microbiol Rev 2010, 34 (6), 1076-112; (b) Osickova, A.; Balashova, N.; Masin, J.; Sulc, M.; Roderova, J.; Wald, T.; Brown, A. C.; Koufos, E.; Chang, E. H.; Giannakakis, A.; Lally, E. T.; Osicka, R., Cytotoxic activity of Kingella kingae RtxA toxin depends on post-translational acylation of lysine residues and cholesterol binding. Emerg Microbes Infect 2018, 7 (1), 178. 25. Komaniecki, G.; Lin, H., Lysine Fatty Acylation: Regulatory Enzymes, Research Tools, and Biological Function. Front Cell Dev Biol 2021, 9, 717503. 26. Kosciuk, T.; Price, I. R.; Zhang, X.; Zhu, C.; Johnson, K. N.; Zhang, S.; Halaby, S. L.; Komaniecki, G. P.; Yang, M.; DeHart, C. J.; Thomas, P. M.; Kelleher, N. L.; Fromme, J. C.; Lin, H., NMT1 and NMT2 are lysine myristoyltransferases regulating the ARF6 GTPase cycle. Nat Commun 2020, 11 (1), 1067. 27. (a) Jing, H.; Zhang, X.; Wisner, S. A.; Chen, X.; Spiegelman, N. A.; Linder, M. E.; Lin, H., SIRT2 and lysine fatty acylation regulate the transforming activity of K-Ras4a. Elife 2017, 6; (b) Jiang, H.; Khan, S.; Wang, Y.; Charron, G.; He, B.; Sebastian, C.; Du, J. T.; Kim, R.; Ge, E.; Mostoslavsky, R.; Hang, H. C.; Hao, Q.; Lin, H. N., SIRT6 regulates TNF-alpha secretion through hydrolysis of long-chain fatty acyl lysine. Nature 2013, 496 (7443), 110-+; (c) Zhang, X.; Spiegelman, N. A.; Nelson, O. D.; Jing, H.; Lin, H., SIRT6 regulates Ras-related protein R-Ras2 by lysine defatty acylation. Elife 2017, 6; (d) Spiegelman, N. A.; Zhang, X.; Jing, H.; Cao, J.; Kotliar, I. B.; Aramsangtienchai, P.; Wang, M.; Tong, Z.; Rosch, K. M.; Lin, H., SIRT2 and Lysine Fatty Acylation Regulate the Activity of RalB and Cell Migration. ACS Chem Biol 2019, 14 (9), 2014-2023. 28. Ribet, D.; Cossart, P., Pathogen-mediated posttranslational modifications: A re-emerging field. Cell 2010, 143 (5), 694-702. 29. Levine, M. M.; Black, R. E.; Clements, M. L.; Cisneros, L.; Saah, A.; Nalin, D. R.; Gill, D. M.; Craig, J. P.; Young, C. R.; Ristaino, P., The pathogenicity of nonenterotoxigenic Vibrio cholerae serogroup O1 biotype El Tor isolated from sewage water in Brazil. J Infect Dis 1982, 145 (3), 296-9. 30. Ferrieres, L.; Hemery, G.; Nham, T.; Guerout, A. M.; Mazel, D.; Beloin, C.; Ghigo, J. M., Silent Mischief: Bacteriophage Mu Insertions Contaminate Products of Escherichia coli Random Mutagenesis Performed Using Suicidal Transposon Delivery Plasmids Mobilized by Broad- Host- Range RP4 Conjugative Machinery. Journal of Bacteriology 2010, 192 (24), 6418-6427. 31. Gibson, D. G.; Young, L.; Chuang, R. Y.; Venter, J. C.; Hutchison, C. A.; Smith, H. O., Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 2009, 6 (5), 343-U41. 32. (a) Horton, R. M.; Hunt, H. D.; Ho, S. N.; Pullen, J. K.; Pease, L. R., Engineering Hybrid Genes without the Use of Restriction Enzymes - Gene-Splicing by Overlap Extension. Gene 1989, 77 (1), 61-68; (b) Dalia, A. B.; Lazinski, D. W.; Camilli, A., Identification of a Membrane-Bound Transcriptional Regulator That Links Chitin and Natural Competence in Vibrio cholerae. Mbio 2014, 5 (1). 33. Dalia, A. B.; McDonough, E.; Camilli, A., Multiplex genome editing by natural transformation. Proc Natl Acad Sci U S A 2014, 111 (24), 8937-42. 34. Donnenberg, M. S.; Kaper, J. B., Construction of an eae deletion mutant of enteropathogenic Escherichia coli by using a positive-selection suicide vector. Infect Immun 1991, 59 (12), 4310-7. 35. Rui, H. P.; Ritchie, J. M.; Bronson, R. T.; Mekalanos, J. J.; Zhang, Y. X.; Waldor, M. K., Reactogenicity of live-attenuated Vibrio cholerae vaccines is dependent on flagellins. P Natl Acad Sci USA 2010, 107 (9), 4359-4364. CHAPTER 3 GOLGI STRESS INDUCES SIRT2 TO COUNTERACT SHIGELLA INFECTION VIA DEFATTY ACYLATION This is a revised version of the submitted paper: Wang, M.*, Zhang, Y.*, Komaniecki, G.*, Cao, J., Yu, T., Hou, D., Zhang, M., Lu, X., Yu, T., Spielgelman, N., Yang, M., Price, I., Lin, H. (2021) Golgi Stress Induces SIRT2 to Counteract Shigella Infection via Defatty acylation, Nat Commun (manuscript in revisions). *Authors contributed equally. This work was initiated and led by Dr. Miao Wang. I contributed many experimental data during the revision process, including verifying CREB3’s role in SIRT2 regulation through ChIP and luciferase based assays, and S. flexneri infection studies in mice and cell culture. For the purpose of this dissertation, they have been revised to highlight my contributions and the thematic connections to the work at large. ABSTRACT Enzymes from pathogens often modulate host protein post-translational modifications (PTMs), facilitating survival and proliferation of pathogens. Shigella virulence factors IpaJ and IcsB induce proteolytic cleavage and lysine fatty acylation on host proteins, which cause Golgi stress and suppress innate immunity, respectively. However, it is unknown whether host enzymes could reverse such modifications introduced by pathogens’ virulence factors to suppress pathogenesis. Herein, we report that SIRT2, a potent lysine defatty acylase, is upregulated by the transcription factor CREB3 under Golgi stress induced by Shigella infection. SIRT2 in turn removes the lysine fatty acylation introduced by Shigella virulence factor IcsB to enhance host innate immunity. SIRT2 knockout mice are more susceptible to Shigella infection than wildtype mice, demonstrating the importance of SIRT2 to counteract Shigella infection. INTRODUCTION Protein post-translational modifications (PTMs) are essential for normal physiology as well as pathogenesis. In the past decades, emerging evidence has demonstrated that pathogens utilize various PTMs as a key strategy to modulate host proteins for their optimal survival and proliferation 1,2. Such PTMs, including ADP-ribosylation 3, adenylation or AMPylation 4-6, and ubiquitination 7 target important pathways of host cells, including translation 8, signaling 9, membrane trafficking 10, and cytoskeleton arrangement 11,12. Recently, it was reported that Shigella flexneri also modulates different host protein PTMs. S. flexneri effector, IpaJ, induces Golgi fragmentation through the cleavage of N-myristoylated host proteins 13. The Golgi apparatus, apart from its role in membrane trafficking, also serves to integrate and transduce stress stimuli. When the Golgi is disassembled upon cellular stress, it signals to the nucleus to help relieve the stress 14. Another effector from S. flexneri, IcsB, alters signaling by inducing lysine fatty acylation on an array of host proteins 15. In contrast to the diverse host protein PTMs induced by pathogens, very little is known about whether host cells can reverse such modifications to fight back. Lysine fatty acylation is an emerging PTM. Recently, multiple mammalian sirtuins and histone deacetylases (HDACs) were discovered to be able to catalyze the removal of lysine fatty acylation in vitro and in cells, which suggested that this PTM might be tightly regulated with important roles in cellular function 16-19. However, only a few proteins were reported to possess lysine fatty acylation and the mammalian lysine fatty acyl transferase for most of the proteins remain unknown 18-23. With the recent identification of IcsB as a robust fatty acyltransferase from S. flexneri 15, it is intriguing to hypothesize that host sirtuins and HDACs fight Shigella infection by counteracting the function of IcsB via their defatty acylation activities. Herein, we find SIRT2, a nicotinamide adenine dinucleotide (NAD+)-dependent protein lysine deacylase, is transcriptionally induced by Golgi stress and serves as a potent defatty acylase to counteract the action of IcsB during Shigella infection. These findings reveal the novel anti-bacterial role of SIRT2 and establish reversible lysine fatty acylation as an important PTM in the war between pathogenic bacteria and mammalian hosts. RESULTS Golgi stress upregulates SIRT2 via CREB3 The project initially started when we were interested in finding out how SIRT2 is regulated as we believe knowing its regulation is crucial for understanding its biological function. Interestingly, we found that Brefeldin A (BFA) drastically increased SIRT2 protein levels in multiple cell lines (Figure 3.1A). BFA is known to induce Golgi stress 24. The ER stress inducers, tunicamycin (Tm) and thapsigargin (Tg), only slightly increased SIRT2 (Figure 3.1B), which further confirmed the specificity of the stress response. Quantitative real-time (RT)-PCR revealed that the SIRT2 transcript level increased more than 50-fold under BFA treatment (Figure 3.1C). Several sirtuins and HDACs were examined under Golgi stress and the SIRT2 increase was the most drastic. Taken together, the data suggest SIRT2 is upregulated by Golgi stress through transcriptional control. Figure 3. 1. Golgi stress upregulates SIRT2. A) Immunoblots for SIRT2 in different cell lines treated with 5 g/ml BFA for 24 hrs. HSP90 blots are shown as loading controls. Representative images from three independent experiments are shown. B) In A549 cells, BFA treatment significantly increased SIRT2 protein level, while the ER stress inducers Tunicamycin (Tm) and Thapsigargin (Tg) only weakly induced SIRT2 expression. C) RT-PCR analysis for SIRT1-7 and HDAC11 mRNA level in A549 cells treated with 5 g/ml BFA for 24 hrs. Fold change in mRNA was calculated by comparing samples with BFA treatment to control DMSO. Next, we tried to identify the transcription factor responsible for SIRT2 upregulation under Golgi stress. The Golgi apparatus is hypothesized to integrate and transduce stress stimuli to the nucleus 14. One of the mechanisms is through the CREB3 transcription factor 25. Thus, we decided to test if CREB3 is the transcription factor controlling SIRT2 expression under Golgi stress. To test if CREB3 upregulates SIRT2 under Golgi stress, CREB3 stable knockdown cells were treated with BFA and other Golgi stress inducers. As expected, the upregulation of SIRT2 was diminished in CREB3 knockdown cells at both protein and mRNA levels while overexpression of CREB3 rescued this effect (Figure 3.2A – B). Additionally, rescue of CREB3 KD by CREB3 overexpression could rescue BFA-induction of SIRT2 level (Figure 3.2C). Moreover, SIRT2 protomer contains a strong CREB3 binding site, and is responsive to the expression of the active (cleaved) form CREB3 or BFA treatment (Figure 3.3A – D). CREB3 binding of the SIRT2 promoter was further confirmed by ChIP which showed increased enrichment of the SIRT2 promoter with α-CREB3 IP after BFA treatment (Figure 3.3 E – F). Thus, the increased SIRT2 expression under Golgi stress is through CREB3. Figure 3. 2. CREB3 promotes SIRT2 transcription under Golgi stress. A) RT-PCR analysis for Sirt2 mRNA in A549 CREB3 KD cells treated with or without BFA. mRNA is normalized to DMSO treated control (shLuc) cells. Statistical evaluation was done by two-way ANOVA. (B) Immunoblots for SIRT2 protein levels in A549 control (shLuc) and CREB3 KD cells treated with or without BFA. HSP90 blot was used as the loading control. (C) Immunoblots for SIRT2 in control and CREB3 KD cells treated with BFA and rescued with CREB3 transfection. SIRT2 level relative to actin loading indicated below blots. Figure 3. 3. CREB3 binds the Sirt2 promoter to regulate transcription. A) A schematic representation of SIRT2 promoter. The annotated regions are on Chromatin 19 complement[39,389,400-39,391,600], reference: GRCh37. B – C) SIRT2-promoter driven firefly transcription in cells under CREB3 N-terminal bZIP domain overexpression (B) or BFA treatment (C). The Renilla luciferase construct was used as an internal control. D) Firefly luciferase transcription in cells treated with BFA. Cells were transfected with an empty pGL3- basic vector or pGL3-basic vector with 994 base pairs of SIRT2 5’ UTR. E) Immunoprecipitation from cells treated with BFA using an α-CREB3 antibody. Blsot shows full length and cleaved version of CREB3. F) qPCR of Sirt2 promoter from chromatin IP of cells treated with or without BFA. While trying to understand the physiological relevance of BFA-induced Golgi stress and the possible role of SIRT2 upregulation, it came to our attention that an intracellular pathogen, Shigella flexneri, is known to induce Golgi fragmentation to inhibit host cell secretion 13. Shigella utilizes a type III secretion system (TTSS) to inject multiple effectors into host cells for successful invasion. One of the effectors, IpaJ, induces Golgi stress by proteolytic cleavage of N- terminal glycine myristoylated proteins, such as ADP-ribosylation factors (ARFs) 26. To test if SIRT2 is upregulated by pathogen-induced Golgi stress, we overexpressed IpaJ in HEK293T and A549 cells. Indeed, SIRT2 protein level was upregulated (Figure 3.4A). Infecting cells or mice with wildtype S. flexneri increased SIRT2 protein and mRNA levels while infection with the IpaJ deletion strain or IpaJ/VirA double deletion strain did not significantly change SIRT2 levels (Figure 3.4B – E). The observed SIRT2 increase on cell line level was relatively small, partly because cells are killed after prolonged S. flexneri infection, while at least 12 hours of Golgi stress is required to induce robust SIRT2 upregulation. Overexpression of EspG and VirA, Golgi stress-inducing bacterial toxins from enteropathogenic E. coli and Shigella respectively, also upregulated SIRT2 consistent with a conserved mechanism of SIRT2 regulation during bacteria-induced Golgi stress (Figure 3.4F). Overall, the data is consistent with the idea that Shigella infection upregulates SIRT2 via Golgi stress and suggest a potential role of SIRT2 in pathogen-host interaction. Figure 3. 4. Shigella infection upregulates SIRT2 through Golgi stress. A) SIRT2 protein level is induced by IpaJ, showing by immunoblots for SIRT2 protein levels in HEK293T cells with or without Flag-IpaJ overexpression. HSP90 is the loading control. B) Immunoblots of SIRT2 in BMDMs infected with mock, wildtype or IpaJ deletion S. flexneri M90T from 6 to 8- week old wildtype C57BL/6J mice. C) Quantification of SIRT2 protein levels (normalized to actin) in B. Statistical evaluation was done by student t-test. D) Immunoblots of Sirt2 in bronchoalveolar lavage cells from 6 to 8-week old wildtype C57BL/6J mice intranasally infected with mock, wildtype or IpaJ deletion S. flexneri M90T. E) Quantification of SIRT2 protein levels (normalized to HSP90) in D. Statistical evaluation was done by one-way ANOVA. F) Immunoblots of HEK 293T cells transfected with empty vector or flag tagged EspG or VirA. SIRT2 counteracts the action of IcsB. To identify the role of SIRT2 during Shigella infection, we focused on the enzymatic activity of SIRT2. SIRT2 can hydrolyze long-chain fatty acyl groups on protein lysine residues. Interestingly, another effector from Shigella, IcsB, is able to fatty acylate host proteins 15. Therefore, we hypothesized that, upon Shigella infection, SIRT2 is upregulated to remove the PTM installed by IcsB as a host defense mechanism. We first tested whether SIRT2 could reverse the fatty acylation added by IcsB in vitro. An alkyne-tagged long-chain fatty acid analog, Alk14, was used to metabolically label substrate proteins 27. Several known IcsB substrate proteins were tagged with a flag tag, co-overexpressed with IcsB and immunoprecipitated from HEK293T cell that were treated with Alk14. Next, we incubated the isolated proteins with purified recombinant SIRT2 with or without NAD+, and then conjugated the substrate proteins to BODIPY-azide using click chemistry to allow visualization of fatty acylation level by in-gel fluorescence. Hydroxylamine (NH2OH) was used to remove cysteine palmitoylation. Remarkably, incubation of multiple IcsB substrate proteins with SIRT2 resulted in the removal of most of the lysine fatty acylation from the proteins in the presence of NAD+ (Figure 3.5A). This suggests that the substrate scope of IcsB and SIRT2 overlap significantly. C HDAC11 Figure 3. 5. SIRT2 can remove IcsB-catalyzed lysine fatty acylation. A) In-gel fluorescence detection of lysine fatty acylation of Flag-tagged IcsB substrate proteins treated with 5 M of SIRT2, with or without 1 mM of NAD+ in vitro. B) In-gel fluorescence detection of lysine fatty acylation of Flag-tagged IcsB substrates in HEK293T cells that were also transfected with Flag- tagged IcsB and SIRT2. Representative images from at least 3 independent experiments are shown. EV, empty vector. WT, SIRT2 WT. HY, SIRT2 HY. FL, fluorescence (indicative of lysine fatty acylation level on substrate proteins). Flag, anti-Flag immunoblot (indicative of input level of substrate proteins). C) In-gel fluorescence detection of lysine fatty acylation of Flag- tagged CHMP5 in HEK293T cells that were co-transfected with Flag-tagged IcsB and SIRT2, empty vector (EV), SIRT6, SIRT7, or HDAC11. We next tested if SIRT2 overexpression could counteract the effect of IcsB in cells. In this experiment, SIRT2-H187Y (HY), a previously reported dead deacetylase but weak defatty acylase, was used as a control 18. We co-overexpressed Flag-tagged IcsB substrate proteins, IcsB, and SIRT2 in HEK293T and examined the lysine fatty acylation on the substrate proteins by Alk14 labeling and in-gel fluorescence. As expected, co-expression of SIRT2 with multiple IcsB substrate proteins significantly decreased their lysine fatty acylation levels, whereas co- expression of SIRT2-HY had much less effect (Figure 3.5B). HDAC11, but not SIRT6 or SIRT7, could also decrease IcsB substrate fatty acylation, but is not regulated by Golgi stress suggesting the possibility of an additional alternative role during Shigella infection (Figure 3.5C). Figure 3. 6. SIRT2 -catalyzed defatty acylation promotes Rho GTPases and Rho GDI interaction. The disruption of Rho GTPase binding to RhoGDI by IcsB is rescued by SIRT2. HEK293T cells were transfected with plasmids encoding indicated proteins: IcsB, SIRT2 WT or HY mutant, and Flag-tagged Rac1 (A), Rac2 (B) or CDC42 (C). Cell lysates for each sample were subjected to GST-RhoGDI pulldown assay (shown in the first panel) and lysine fatty acylation labeling assay (shown in the last panel). The quantification of GDI enrichment of Rho GTPases, obtained by dividing Flag-Rho GTPase signal in GDI pulldown with Flag-Rho GTPase in the input (second panel), is shown at the bottom. It is reported that the fatty acylation on Rho GTPase inhibits the protein-protein interaction between Rho GTPases and RhoGDI 28, which regulates the membrane cycling and signaling of Rho GTPases 29,30. We therefore tested whether this was true and if so, whether SIRT2 could rescue this effect. As shown in Figure 3.6, lysine fatty acylation inhibits Rho GTPase and RhoGDI interaction, and this can be rescued by SIRT2 overexpression. Thus, SIRT2 works as a defatty acylase to maintain Rho GTPase signaling during Shigella infection. SIRT2 limits Shigella autophagosome escape. Host cells use autophagy to target intracellular bacteria and to restrict bacterial growth 33. IcsB is important for Shigella to escape host autophagy by modifying CHMP5 15,34. We hypothesized that SIRT2, working against IcsB, would promote the innate immunity and suppress Shigella escaping from autophagosomes. To test this hypothesis, SIRT2 knockout (Sirt2-/-) and wildtype (Sirt2+/+) mouse embryonic fibroblast (MEF) cells were infected with S. flexneri. Bacteria that were trapped in autophagosomes were visualized by staining LC3, an autophagosome-specific marker. Consistent with previous reports, more Icsb deletion (Δicsb) Shigella was trapped in autophagosomes than wildtype Shigella (Figure 3.7A-B). The percentage of Sirt2-/- MEF cells containing LC3 decorated wildtype Shigella was significantly less than those of Sirt2+/+ MEF cells (Figure 3.7B). This is further confirmed by LC3 turnover assay (Figure 3.7B). The data supports that SIRT2 limits Shigella autophagosome escape. SIRT2 restricts Shigella infection in cells and in mice. To further test if SIRT2 is able to restrict proliferation of Shigella, we performed a gentamycin killing assay to examine the intracellular Shigella growth. Equal number of Sirt2+/+ Figure 3. 7. SIRT2 suppresses Shigella autophagosome escape. A) Effect of SIRT2 knockout on S. flexneri autophagosome escape. Representative images of MEF cells infected with S. flexneri from three biological replicates are shown. Scale bar: 15 m. White arrow: LC3-positive Shigella. B) Quantitation of data shown in A. Percentage of MEF cells containing LC3-positive shigella was quantified. Data is shown as mean ± SEM with >100 infected MEF cells counted for each experiment. Statistical evaluation was done using two-way ANOVA. Data are represented as mean ± SEM. *p<0.05. ns, not significant. C) Autophagy level (measured by ratio of LC3-II to LC3-I) is lower in SIRT2 knockout and knockdown cells upon S. flexneri infection. Immunoblotting analysis of LC3 in SIRT2 knockdown A549 cells infected with wildtype or IcsB deletion S. flexneri M90Tstrain at the same MOI. The quantification of LC3-II/LC3-I is shown at the bottom. and Sirt2-/- MEF cells were infected with equal number of S. flexneri for 10 min before treated with gentamycin to kill extracellular bacteria. The intracellular bacteria number was measured by recoverable colony formation units (CFU) to give the MOI. This assay revealed that Shigella survived better without SIRT2, at both early and later time points (Figure 3.8A). Similar trends were also observed in A549 control or SIRT2 knockdown cells (Figure 3.8B). CREB3 KD cells have increased bacterial load after infection which can be partially rescued by WT but not HY SIRT2, suggesting that regulation of SIRT2 is a key function for CREB3 during Shigella infection (Figure 3.8D). The protective effect of SIRT2 is impaired against S. flexneri IpaJ deletion strain, suggesting the relevance of Golgi stress mediated SIRT2 regulation (Figure 3.8B). We also examined the effect of Sirt2 in wildtype and Sirt2 knockout C57/B6J mice. Six to eight-week old mice were intranasally administrated with S. flexneri, closely monitored, and sacrificed after 1-3 days. S. flexneri present in the lung were quantified. Consistently, there are more S. flexneri in Sirt2-/- mice, which further confirmed our model that SIRT2 protects against Shigella infection (Figure 3.8C). We also observed drastically reduced physical activity in Sirt2-/- mice comparing with Sirt2+/+ mice after bacterial infection. Overall, the data demonstrate SIRT2 restricts Shigella infection both in cells and in vivo. Δipaj Δipaj Figure 3. 8. SIRT2 restricts Shigella infection in cells and in mice model. A) Effect of Sirt2 knockout on S. flexneri intracellular proliferation analyzed by gentamycin killing assay. MEF Sirt2+/+ and Sirt2-/- cells were infected with equal number of S. flexneri for 10 min. Intracellular S. flexneri number and MEF cell number were counted at indicated time points to get MOI value. Data are presented as mean ± SEM with three biological replicates, and each with two technical replicates. Statistical evaluation was done by an unpaired two-tailed Student’s t test. B) The protective effect of SIRT2 was suppressed when IpaJ is deleted. A549 control and SIRT2 KD cells were infected with equal number of wildtype and IpaJ deletion S. flexneri M90T cells for 10 min, then treated with gentamycin to kill extracellular bacteria. Intracellular S. flexneri number and mammalian cell number were counted to get MOI. Data are presented as mean ± SEM with nine biological replicates. Statistical evaluation was done using two-way ANOVA. C) Recoverable CFU in lung homogenates from 6 to 8-week old Sirt2+/+ and Sirt2-/- C57BL/6J mice intranasally infected with 1 million S. flexneri M90T wildtype strain. Statistical evaluation was done using unpaired two-tail Student’s t test. Data are presented as mean ± SEM with 3 mice per group. D) CREB3 KD cells transfected with WT or HY SIRT2 and infected with S. flexneri M90T for 10 mins before washing with PBS and replacing the media with 50 g/ml gentamicin for 6 hours. Cells were collected and CFU/cell determined. To validate if the effect of SIRT2 was through counteracting the effect of IcsB, we infected the same number of Sirt2+/+ and Sirt2-/- MEF cells with same number of wildtype and Δicsb S. flexneri. Consistently, wildtype S. flexneri proliferated much better in Sirt2-/- cells than in Sirt2+/+ cells (Figure 3.9A). In contrast, Δicsb S. flexneri proliferated at similarly low levels in both Sirt2+/+ and Sirt2-/- cells, suggesting SIRT2 counteracts IcsB specifically (Figure 3.9A). Interestingly, wildtype and Δicsb S. flexneri proliferated similarly in Sirt2+/+ MEF cells, suggesting that SIRT2 is able to fully counteract IcsB (Figure 3.9A). The same trend was also observed in vivo (Figure 3.9B). In intranasally infected mice, we observed similar MOI for Δicsb S. flexneri in Sirt2+/+ and Sirt2-/- mouse lungs, but significantly higher MOI of wildtype S. flexneri in Sirt2-/- than in Sirt2+/+ mouse lungs (Figure 3.9B). To further confirm the SIRT2/IcsB axis, we generated Δicsb S. flexneri rescued by either WT or catalytic dead C306A IcsB. When infected with WT IcsB S. flexneri, Sirt2-/- mice had much more bacterial colonization. Sirt2+/+ and Sirt2-/- mice showed no statistically significant difference in colonization by C306A S. flexneri (Figure 3.9C). These results demonstrate the in vivo antibacterial role of SIRT2 as a defatty acylase to counteract the action of IcsB. Figure 3. 9. SIRT2 restricts Shigella infection in cell and in vivo by counteracting IcsB. (A) Effect of SIRT2 knockout and IcsB deletion on S. flexneri intracellular proliferation. MOI were determined after 10 min of infection and 6 hrs of incubation in the presence of 50 g/ml gentamicin. Data are represented as mean ± SEM with three biological replicates. Statistical evaluation was done by two-way ANOVA. ***p<0.001, ns, not significant. (B) CFU in lung homogenates from Sirt2+/+ and Sirt2-/- C57BL/6J mice infected with wildtype or IcsB deletion S. flexneri M90T for 3 days. Data are presented as mean ± SEM with 3 mice per group. Statistical evaluation was done using two-way ANOVA. *p<0.05, ns, not significant. (C) CFU in lung homogenates from Sirt2+/+ and Sirt2-/- C57BL/6J mice infected with wildtype or IcsB deletion S. flexneri M90T rescued with WT or C306A IcsB for 3 days. Data are presented as mean ± SEM with 3 mice per group. Statistical evaluation was done using two-way ANOVA. *p<0.05, ns, not significant. (D) Model depicting SIRT2 as a Golgi stress response protein that limits Shigella pathogenesis by counteracting Shigella-mediated host protein lysine fatty acylation. DISCUSSION Emerging studies suggest that pathogens employ PTMs to modulate host protein functions to facilitate pathogenesis 36. Such PTMs are often highlighted with novel enzymatic reactions and high reaction efficiency. One of the PTMs, protein fatty acylation, regulates the association of proteins with membranes, and is important for membrane trafficking, protein- protein interaction, and signal transduction 37. Fatty acylation on lysine residues, however, is less well studied. Multiple mammalian lysine defatty acylases were identified 16,18,19,22,38, which strongly suggested the physiological relevance of this PTM. However, very little is known on mammalian lysine fatty acyl transferases 23, which has greatly limited the study of the function of this PTM. Our findings demonstrate that reversible lysine fatty acylation plays an important role in pathogen-host interaction, highlighting the importance of lysine fatty acylation in innate immunity. SIRT2 was initially recognized as a lysine deacetylase, with its cellular function mostly attributed to its deacetylation activity 39. Many in vitro studies suggested that SIRT2 can also catalyze lysine defatty acylation 40. However, the first physiological de-fatty acylation substrate, K-Ras4a, was only recently identified 18,41. Our findings here thus provide key insights that will help understand the physiological significance of SIRT2’s defatty acylation activity and expands its substrate scope. As the hub of intracellular membrane trafficking, Golgi apparatus possess a unique structure where the stacks are interconnected into a compact ribbon. The ribbon undergoes dynamical membrane fission, fusion, and cargo transport through cisternal maturation 42. Upon certain cellular events such as mitosis and stress conditions, the ribbon can undergo distinct disassembly processes, which will lead to disruption of Golgi integrity and stress stimuli transduction to the nucleus 14. Our study revealed an important function of the Golgi stress pathway in fighting Shigella infection. Through Golgi stress, cells sense Shigella infection and activate SIRT2 via the CREB3 transcription factor to enhance innate immunity. Among all of the known lysine defatty acylases, SIRT2 is the most responsive to Golgi stress induced by pathogen. This highlights the unique role of SIRT2 in stress response. Previously, SIRT2 was reported to be hijacked by other pathogens to work as deacetylase and to promote pathogen invasion 43-45 . Here our study discovered an anti-bacterial role of SIRT2 by counteracting the fatty acyl transferase IcsB from S. flexneri (Figure 3.9D). Many other pathogen fatty acyl transferases have been identified or predicted. For example, MARTX from Vibrio cholerae shares sequence similarities and substrate proteins with IcsB 46. In addition, heat labile toxin (LTA1) that shares sequence similarities with cholera toxin, also disrupts Golgi apparatus in a pattern that is reminiscent of the actions of BFA 47. This suggests that the mechanism of SIRT2’s anti-bacterial role is potentially broader. In addition to lysine fatty acylation, pathogenic bacteria use many other PTMs to help infect host cells. As far as we know, lysine fatty acylation is the first pathogen-induced PTM that can be efficiently removed by host enzymes. This suggests that other pathogen induced PTMs may also be reversed by host enzymes and studying them may provide insights into mechanisms of innate immune responses. METHODS Reagents, antibodies and plasmids. Tunicamycin, Thapsigargin, cycloheximide were purchased from Sigma-Aldrich (St. Louis, MO); Actinomycin D and gentamicin were from Thermo Fisher. Brefeldin A was from Cell Signaling Technology (Danvers, MA). NAD+ was purchased from VWR (Radnor, PA). Alk14 and BODIPY-azide were synthesized as previously described 27. The following antibodies were purchased from Cell Signaling Technology (Danvers, MA): SIRT2 (D4050) (#12650), PARP (#9542), LC3 (#3868), and anti-rabbit IgG HRP-linked Antibody (#7074). The following antibodies were from Santa Cruz: β-actin (C-4), GRP78 (H- 129). Anti-Flag M2 antibody conjugated with horseradish peroxidase (A8592) and anti-flag M2 affinity gel (A220) were purchased from Sigma. Anti-CREB3 (ab180119) was from Abcam (Cambridge, MA). IcsB gene was synthesized by Integrated DNA Technologies (Coralville, IA) after codon optimization from Shigella flexneri 5a plasmid pWR100 193718 – 195202 with N-terminal Triple Flag-tag. The gene fragment was later cloned into pCMV4a between EcoRI and XhoI using primers 5'- AGT CAG GAA TTC ACC ATG GAT TAC AAA GAT CAC G -3' and 5'- AGT CAG CTC GAG CTA AAT ATT TGA ATG GGA GTT GTT GA -3' (the restriction sites are underlined). IpaJ was synthesis by Integrated DNA Technologies and was cloned into pEGFP-C1- FLAG-GFP-Ubl4A-C (Addgene #86923) between BglII and SalI with N-terminal Flag-tag followed by EGFP using primers 5'- AGT CAG AGA TCT ATG TCG GAA CAA CGG AAG - 3' and 5'- AGT CAG GTC GAC TTA CAA AGC CTC ATT AGT TAT AAC TAT GGA -3'. pEGFP-C1-FLAG-GFP-Ubl4A-C(90-157) was a gift from Yihong Ye (Addgene plasmid # 86923 ; http://n2t.net/addgene:86923 ; RRID:Addgene_86923) 48. For lentivirus generation, IpaJ was cloned into pCDH-SIRT2-Flag (Addgene # 102624) between EcoRI and XhoI with N- terminal Flag-tag using primers 5'- AGT CAG GAA TTC ATG TCG GAA CAA CGG AAG -3' and 5'- AGT CAG CTC GAG TTA CAA AGC CTC ATT AGT TAT AAC TAT GGA -3'. pCDH-SIRT2-Flag was previously cloned 49. Human CHMP5 was amplified from HEK293T cDNA library and cloned into pCMV4a between EcoRI and XhoI with C-terminal Flag-tag using primers 5'- AGT CAG GAA TTC ACC ATG AAC CGA CTC TTC GGG -3' and 5'- AGT CAG CTC GAG TGA AGC AGG GAT CTG TGG -3'. CHMP5 K7R was generated through site directed mutagenesis using primers 5'- AAC CGA CTC TTC GGG AGA GCG AAA CCC AAG GCT -3' and 5'- TCT CCC GAA GAG TCG GTT CAT GGT GAA TTC CTG -3' (the mutation sites are underlined). Human RhoGDI was amplified from HEK293T cDNA library and cloned into pGEX4T3 between BamHI and EcoRI using primers 5'- AGT CAG GGA TCC ATG GCT GAG CAG GAG C -3' and 5’- AGT CAG GAA TTC TCA GTC CTT CCA GTC CTT C -3’. RhoA, Rac1, Rac2, CDC42, Has, Kras, RRas, Rap1b, RalA, RalB, RheB were generated using a standard PCR cloning strategy. Cell culture, transfection and transduction. Human MCF7, Hela, MDA-MB-231, A2780, and HeyA8 were grown in DMEM media (Invitrogen) supplemented with 10% (v/v) heat- inactivated fetal bovine serum (FBS; Invitrogen, Carlsbad, CA). MEF cells were cultured in DMEM supplemented with non-essential amino acids and 15% (v/v) heat-inactivated FBS. A549 cells were cultured in RPMI-1640 (Invitrogen) with 10% (v/v) heat-inactivated FBS. HEK293T were cultured in DMEM supplemented with 10% (v/v) heat-inactivated calf serum (Sigma). For transient overexpression in HEK293T cells, mammalian expression vectors were transfected using PEI MAX 40K (PolySciences) according to the manufacturer’s protocol. Empty vector was transfected as negative control. To transduce A549 for overexpression or knockdown, lentiviral infection was performed as previously described18. The pLKO.1-puro lentiviral shRNAs constructs for Luciferase and human CREB3 were purchased from Sigma- Aldrich. Luciferase shRNA (SHC007), CREB3 shRNA1 (TRCN0000020342), CREB3 shRNA2 (TRCN0000020343) were used. Sirt2 WT and KO MEF, Sirt2 KO MEF with stable SIRT2 re-overexpression and SIRT2 stable knockdown A549 and HEK293T cells were generated as previously described 18. For ActD and CHX chase experiments, A549 cells were treated with or without BFA at t = 0 hr. During BFA treatment, ActD was added at t = 0, 12, 18, 23, 24 hrs; or CHX was added at t = 0, 12, 24 hrs. at t = 24 hrs, cells were collected and submit for western blot analysis. Bacterial strains and infection. Shigella flexneri 5a M90T is a kind gift from Prof. Neal M. Alto from UT Southwestern. IcsB M90T was generated using the lambda red recombineering system 13. Briefly, M90T containing pKD46 was transformed with KanR cassette from pKD4. Kanamycin resistant colonies were selected and cured for pKD46 to generate lcsB<>kan. The resistance was later removed with pCP20, resulting in a traceless gene deletion. The KanR cassette was amplified using primers 5'- ACA TCC CCA CAA TCA CCA AGT AAT GGA GAG TTA ATA AAG TGT GTA GGC TGG AGC TGC TTC -3' and 5'- AAA GTT TAT CAT ATA GTT TGC GAC ACA TTT CTA TGG CCT TAT GGG AAT TAG CCA TGG TCC -3' (the external overlap with IcsB was underlined). Gentamycin killing assay. S. flexneri infection was done as previously described 13. Briefly, ~1 x 106 mammalian cells were seeded into 6-well dishes. S. flexneri M90T was inoculated overnight at 30 oC in Luria-Bertani (LB) broth (Fisher, Hampton NH) with shaking. Before infection, M90T culture was back diluted 1:100 in fresh LB broth and incubated at 37 oC with shaking until the OD600 reached 0.4. The bacteria were further incubated with 0.03% Congo Red in PBS for 15 min before adding 20 L bacterial suspension to mammalian cells for an MOI of ~8. The infection was facilitated by centrifugation at 1000 g at room temperature for 10 min. After 10 min, the mammalian cells were washed with PBS 5 times and incubated in fresh media containing 50 g/ml gentamycin. At the indicated time points (30 min to 5 hrs), mammalian cells were washed with PBS 3 times, collected and counted under a light microscope. The cells were then lysed with 0.5% Triton X-100 in PBS and the cell lysates were plated on LB agar and colonies forming units (CFU) were counted after overnight incubation at 37 oC. Western blot analysis. Western Blots were performed as described previously 18. The proteins of interest were detected using enzyme-linked fluorescence (ECL Plus; Pierce Biotechnology Inc.) and visualized using the Typhoon 9400 Variable Mode Imager (GE Healthcare, Piscataway, NJ). Quantification of the western blots was done using ImageJ software. RT-PCR analysis of mRNA levels. Total mRNA was extracted using the RNeasy Mini Kit (Qiagen, CA, USA) according to the manufacturer's instructions and then reverse transcribed to cDNA using SuperScript Vilo cDNA Synthesis Kit (Thermo Fisher). Real-time PCR were performed on QuantStudio™ 7 Flex Real-Time PCR System using SYBR™ Green PCR Master Mix (Applied Biosystems) and the primers shown in Table S1. Luciferase assay for Sirt2 promoter activity. The 5’ Sirt2 promoter corresponding to the region –1 to –994 nucleotides upstream of the Sirt2 transcription start site was cloned into the pGL3-basic vector (Promega) to create pGL3-994. Firefly/renilla luciferase ratio was determined by co-transfecting A549 cells with pGL3-994 and pGL4.73 (Renilla luciferase) for 24 hours. Cells were either co-transfected with N-terminal flag tagged CREB3 (1-220) or treated with 5 g/mL BFA for an additional 24 hours. SIRT2 promoter was cloned into pGL3-basic using Gibson cloning using the primers 5’- GTG CTA GCC CGG GCT CGA GAT CTG CGA TCT AAG TAC CAC AGT TCT AAC AGA AGT CTC AGG -3’ and 5’- AGA ATG GCG CCG GGC CTT TCT TTA TGT TTT TGG CGT CTT CCA TGG GCG CGG TG -3’. CREB3 (1-220) was cloned into pCMV5 between EcoRI and XhoI using primers 5’- AGT CAG GAA TTC ATG GAC TAC AAA GAC GAT GAC GAC AAG ATG GAG CTG GAA TTG GAT G -3’ and 5’- AGT CAG CTC GAG CTA TGA TAT CTC AAT CAC CAT GGC -3’. Dual luciferase activity was then measured using the Promega Dual-Luciferase® Reporter Assay (E1910) according to manufacture specifications. To determine BFA-induced luciferase signal for SIRT2 promoter, A549 cells were transfected with pGL3-basic or pGL3-994 for 24 hours then treated with 5 g/mL BFA for an additional 24 hours. Cells were suspended in PBS to determine cell density and then processed according to manufacturer recommendations (Promega E1910). Firefly luciferase signal was normalized to cell number. Chromatin Immunoprecipitation. Approximately 1.5 × 107 A549 cells in a 15 cm dish were treated with 5 g/mL BFA or DMSO for 24 hours. Cells were crosslinked with 1% formaldehyde for 10 minutes and processed according to ChIP-IT® Express Enzymatic Magnetic Chromatin Immunoprecipitation Kit (Active Motif, 53009) manufacturer recommendations using an α-CREB3 antibody (Proteintech, 11275-1-AP). Chromatin inputs and eluted ChIP samples were analyzed via qPCR using the primers 5’- AGA CTC TAG ACC CCT GGT GG -3’ and 5’- TTG GAG GGG GAG AGA AGA ACA -3’ which flank the putative CREB3 binding site and result in a 472 bp amplification product. qPCR was carried out on a QuantStudio 7 Flex (Applied Biosystems) using SYBR green (Abclonal, RK21203). Detection of fatty acylation on protein of interest using Alk14. HEK293T cells were transfected with IcsB and the gene of interest for 24 hrs, and then treated with Alk14 for 6 hrs before harvest. Alternatively, HEK293T cells transfected with gene of interest overnight were infected with M90T in the presence of Alk14 for 3 hrs. The cell pellets were processed as previously described 18. Defatty acylation assay by sirtuins in vitro. HEK293T cells were transfected with IcsB and substrate genes for 24 hrs and treated with Alk14 for 6 hrs before harvest. The immunoprecipitated substrate proteins with Alk14 labeling were pulled down using anti-Flag affinity gel. The affinity gel containing the pulled down proteins was suspended in 25 l of assay buffer (50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 2 mM MgCl2, 1 mM DTT) with 5 M of SIRT2, with or without 1 mM NAD+. The defatty acylation reaction was allowed to proceed for 30 min at 37 oC and quenched by washing the affinity gel with IP wash buffer (25 mM Tris- HCl pH 8.0, 150 mM NaCl, 0.2% Nonidet P-40). On-bead click chemistry and in-gel fluorescence was carried out as described previously. SIRT2 purification was done as previously described 18. GDI pulldown. GST-RhoGDI were purified from E. coli strain BL21(DE3) cells using glutathione resin. For the pulldown assay, 500 g lysates from transfected cells were incubated with 60 g GST-RhoGDI pre-coupled to glutathione resin. After three washes in IP wash buffer, the bound proteins were eluted and analyzed by immunoblotting. Immunofluorescence. MEF cells were infected with M90T at MOI of 100:1 in glass bottom dishes (MatTek, Ashland MA). After 3 hrs, cells were washed with PBS three times and fixed with pre-chilled methanol for 10 min. Then the cells were permeabilized with 0.25% Triton X-100 for 10 min, blocked with 1% (w/v) BSA/0.1% Tween-20 in PBS for 1 hr and incubated with LC3 antibody overnight. The cells were washed with 0.1% Tween-20 in PBS three times and incubated with Cy3-conjugated goat anti-rabbit IgG (H + L) secondary antibody for 1 hr. The cells were washed with 0.1% Tween-20 in PBS three times and mounted with Fluoromount- G (SouthernBiotech, Homewood AL) containing DAPI before imaged on Cytation 5 (BioTek, Winooski VT). The number of MEF cells that contained Shigella and the number of MEF cells that contained LC3-postive Shigella were counted manually. Intraperitoneal shigellosis mice model study. The S. flexneri intraperitoneal infection mouse study was done as previously described 50. Briefly, 6-week old wildtype and SIRT2 knockout C57/B6J mice (Jackson Lab, Sirt2tm1.1Fwa) were intraperitoneally administrated with 150 𝑚𝑖𝑙𝑙𝑖𝑜𝑛 Shigella per 20 grams of mouse body weight. After 17 hrs, mice were sacrificed, and tissue were harvested. The peritoneal wash was also collected. Tissues were washed in PBS with gentamycin (50 g/ml) to remove superficial bacteria, rinsed twice in antibiotic-free PBS, and then mechanically homogenized in PBS with 2.5% Triton-X100 (200 mg of tissue per ml of PBS). The tissue extracts were diluted and plated onto Hektoen Enteric Agar plate. Tissue extracts and peritoneal wash from mice without Shigella infection were used as blank control. Colonies were counted after 18 hrs of culture at 37 oC. All animal experiments were approved by Cornell University’s Institutional Animal Care and Use Committee. Intranasal shigellosis mice model study. The S. flexneri intranasal infection mouse study was done as previously described 13. Briefly, 6- to 8-week-old wildtype and SIRT2 knockout C57/B6J mice were intranasally administrated with 1 𝑚𝑖𝑙𝑙𝑖𝑜𝑛 Shigella. After 3 days (or at indicated timepoints), mice were sacrificed, and lung tissue were harvested. The bronchoalveolar lavage (BAL) fluid was also collected. Tissues were mechanically homogenized in PBS with 2.5% Triton-X100. The tissue extracts were diluted and plated onto Hektoen Enteric Agar plate. Tissue extracts and BAL fluid from mice without Shigella infection were used as blank control. Colonies were counted after 18 hrs of culture at 37 oC. All animal experiments were approved by Cornell University’s Institutional Animal Care and Use Committee. ACKNOWLEDGEMENTS We thank Prof. Neal M. Alto for providing S. flexneri M90T strains and Dr. Toren Finkel for providing the SIRT2+/+ and SIRT2-/- MEF cells. pKD46 and pKD4 plasmids are kind gifts from Dr. Tobias Doerr. REFERENCES 1. Ribet, D. & Cossart, P. Pathogen-mediated posttranslational modifications: A re- emerging field. Cell 143, 694-702, doi:10.1016/j.cell.2010.11.019 (2010). 2. Cui, J. & Shao, F. Biochemistry and cell signaling taught by bacterial effectors. Trends in biochemical sciences 36, 532-540, doi:10.1016/j.tibs.2011.07.003 (2011). 3. Aktories, K. et al. Botulinum C2 toxin ADP-ribosylates actin. Nature 322, 390-392, doi:10.1038/322390a0 (1986). 4. Yarbrough, M. L. et al. AMPylation of Rho GTPases by Vibrio VopS disrupts effector binding and downstream signaling. Science 323, 269-272, doi:10.1126/science.1166382 (2009). 5. Worby, C. A. et al. The fic domain: regulation of cell signaling by adenylylation. Molecular cell 34, 93-103, doi:10.1016/j.molcel.2009.03.008 (2009). 6. Muller, M. P. et al. The Legionella effector protein DrrA AMPylates the membrane traffic regulator Rab1b. Science 329, 946-949, doi:10.1126/science.1192276 (2010). 7. Ashida, H. et al. A bacterial E3 ubiquitin ligase IpaH9.8 targets NEMO/IKKgamma to dampen the host NF-kappaB-mediated inflammatory response. Nature cell biology 12, 66-73; sup pp 61-69, doi:10.1038/ncb2006 (2010). 8. Su, X., Lin, Z. & Lin, H. The biosynthesis and biological function of diphthamide. Critical reviews in biochemistry and molecular biology 48, 515-521, doi:10.3109/10409238.2013.831023 (2013). 9. Li, H. et al. The phosphothreonine lyase activity of a bacterial type III effector family. Science 315, 1000-1003, doi:10.1126/science.1138960 (2007). 10. Huang, J. & Brumell, J. H. Bacteria-autophagy interplay: a battle for survival. Nat Rev Microbiol 12, 101-114, doi:10.1038/nrmicro3160 (2014). 11. Jimenez, A., Chen, D. & Alto, N. M. How Bacteria Subvert Animal Cell Structure and Function. Annu Rev Cell Dev Biol 32, 373-397, doi:10.1146/annurev-cellbio-100814- 125227 (2016). 12. Aktories, K. & Barbieri, J. T. Bacterial cytotoxins: targeting eukaryotic switches. Nat Rev Microbiol 3, 397-410, doi:10.1038/nrmicro1150 (2005). 13. Burnaevskiy, N. et al. Proteolytic elimination of N-myristoyl modifications by the Shigella virulence factor IpaJ. Nature 496, 106-109, doi:10.1038/nature12004 (2013). 14. Machamer, C. E. The Golgi complex in stress and death. Frontiers in neuroscience 9, 421, doi:10.3389/fnins.2015.00421 (2015). 15. Liu, W. et al. N(epsilon)-fatty acylation of multiple membrane-associated proteins by Shigella IcsB effector to modulate host function. Nature microbiology 3, 996-1009, doi:10.1038/s41564-018-0215-6 (2018). 16. Jiang, H. et al. SIRT6 regulates TNF-alpha secretion through hydrolysis of long-chain fatty acyl lysine. Nature 496, 110-113, doi:10.1038/nature12038 (2013). 17. Tong, Z. et al. SIRT7 Is an RNA-Activated Protein Lysine Deacylase. ACS Chem Biol 12, 300-310, doi:10.1021/acschembio.6b00954 (2017). 18. Jing, H. et al. SIRT2 and lysine fatty acylation regulate the transforming activity of K- Ras4a. Elife 6, doi:10.7554/eLife.32436 (2017). 19. Cao, J. et al. HDAC11 regulates type I interferon signaling through defatty acylation of SHMT2. Proceedings of the National Academy of Sciences of the United States of America 116, 5487-5492, doi:10.1073/pnas.1815365116 (2019). 20. Bursten, S. L., Locksley, R. M., Ryan, J. L. & Lovett, D. H. Acylation of monocyte and glomerular mesangial cell proteins. Myristyl acylation of the interleukin 1 precursors. The Journal of Clinical Investigation 82, 1479-1488, doi:10.1172/JCI113755 (1988). 21. Jiang, H. et al. SIRT6 regulates TNF-α secretion through hydrolysis of long-chain fatty acyl lysine. Nature 496, 110-113, doi:10.1038/nature12038 (2013). 22. Zhang, X., Spiegelman, N. A., Nelson, O. D., Jing, H. & Lin, H. SIRT6 regulates Ras- related protein R-Ras2 by lysine defatty acylation. Elife 6, doi:10.7554/eLife.25158 (2017). 23. Kosciuk, T. et al. NMT1 and NMT2 are lysine myristoyltransferases regulating the ARF6 GTPase cycle. Nature Communications 11, 1067, doi:10.1038/s41467-020-14893-x (2020). 24. Klausner, R. D., Donaldson, J. G. & Lippincott-Schwartz, J. Brefeldin A: insights into the control of membrane traffic and organelle structure. The Journal of cell biology 116, 1071-1080, doi:10.1083/jcb.116.5.1071 (1992). 25. Reiling, J. H. et al. A CREB3-ARF4 signalling pathway mediates the response to Golgi stress and susceptibility to pathogens. Nature cell biology 15, 1473-1485, doi:10.1038/ncb2865 (2013). 26. Burnaevskiy, N., Peng, T., Reddick, L. E., Hang, H. C. & Alto, N. M. Myristoylome profiling reveals a concerted mechanism of ARF GTPase deacylation by the bacterial protease IpaJ. Molecular cell 58, 110-122, doi:10.1016/j.molcel.2015.01.040 (2015). 27. Charron, G. et al. Robust fluorescent detection of protein fatty acylation with chemical reporters. Journal of the American Chemical Society 131, 4967-4975, doi:10.1021/ja810122f (2009). 28. Liu, W. et al. Nε-fatty acylation of multiple membrane-associated proteins by Shigella IcsB effector to modulate host function. Nat Microbiol 3, 996-1009, doi:10.1038/s41564- 018-0215-6 (2018). 29. Boulter, E. et al. Regulation of Rho GTPase crosstalk, degradation and activity by RhoGDI1. Nature Cell Biology 12, 477-483, doi:10.1038/ncb2049 (2010). 30. Garcia-Mata, R., Boulter, E. & Burridge, K. The 'invisible hand': regulation of RHO GTPases by RHOGDIs. Nature Reviews Molecular Cell Biology 12, 493-504, doi:10.1038/nrm3153 (2011). 31. Teng, Y. B. et al. Efficient demyristoylase activity of SIRT2 revealed by kinetic and structural studies. Scientific Reports 5, doi:10.1038/srep08529 (2015). 32. Jiang, H., Zhang, X. & Lin, H. Lysine fatty acylation promotes lysosomal targeting of TNF-α. Sci Rep 6, 24371, doi:10.1038/srep24371 (2016). 33. Xu, Y. et al. A Bacterial Effector Reveals the V-ATPase-ATG16L1 Axis that Initiates Xenophagy. Cell 178, 552-566 e520, doi:10.1016/j.cell.2019.06.007 (2019). 34. Ogawa, M. et al. Escape of intracellular Shigella from autophagy. Science 307, 727-731, doi:10.1126/science.1106036 (2005). 35. Sharma, D. et al. Shigellosis murine model established by intraperitoneal and intranasal route of administration: a comparative comprehension overview. Microbes and infection 19, 47-54, doi:10.1016/j.micinf.2016.09.002 (2017). 36. Salomon, D. & Orth, K. What pathogens have taught us about posttranslational modifications. Cell host & microbe 14, 269-279, doi:10.1016/j.chom.2013.07.008 (2013). 37. Jiang, H. et al. Protein Lipidation: Occurrence, Mechanisms, Biological Functions, and Enabling Technologies. Chemical reviews 118, 919-988, doi:10.1021/acs.chemrev.6b00750 (2018). 38. Aramsangtienchai, P. et al. HDAC8 Catalyzes the Hydrolysis of Long Chain Fatty Acyl Lysine. ACS Chem Biol 11, 2685-2692, doi:10.1021/acschembio.6b00396 (2016). 39. Kosciuk, T., Wang, M., Hong, J. Y. & Lin, H. Updates on the epigenetic roles of sirtuins. Curr Opin Chem Biol 51, 18-29, doi:10.1016/j.cbpa.2019.01.023 (2019). 40. Wang, Y. et al. Deacylation Mechanism by SIRT2 Revealed in the 1'-SH-2'-O-Myristoyl Intermediate Structure. Cell Chem Biol 24, 339-345, doi:10.1016/j.chembiol.2017.02.007 (2017). 41. Spiegelman, N. A. et al. A Small-Molecule SIRT2 Inhibitor That Promotes K-Ras4a Lysine Fatty acylation. ChemMedChem 14, 744-748, doi:10.1002/cmdc.201800715 (2019). 42. Klumperman, J. Architecture of the mammalian Golgi. Cold Spring Harbor perspectives in biology 3, doi:10.1101/cshperspect.a005181 (2011). 43. Eskandarian, H. A. et al. A role for SIRT2-dependent histone H3K18 deacetylation in bacterial infection. Science 341, 525-+, doi:10.1126/science.1238858 (2013). 44. Gogoi, M. et al. Salmonella escapes adaptive immune response via SIRT2 mediated modulation of innate immune response in dendritic cells. Plos Pathogens 14, doi:10.1371/journal.ppat.1007437 (2018). 45. Bhaskar, A. et al. Host Sirtuin 2 as an immunotherapeutic target against tuberculosis. eLife 9, e55415, doi:10.7554/eLife.55415 (2020). 46. Zhou, Y. et al. N(epsilon)-Fatty acylation of Rho GTPases by a MARTX toxin effector. Science 358, 528-531, doi:10.1126/science.aam8659 (2017). 47. Zhu, X. & Kahn, R. A. The Escherichia coli Heat Labile Toxin Binds to Golgi Membranes and Alters Golgi and Cell Morphologies Using ADP-ribosylation Factor- dependent Processes. Journal of Biological Chemistry 276, 25014-25021 (2001). 48. Xu, Y., Anderson, D. E. & Ye, Y. The HECT domain ubiquitin ligase HUWE1 targets unassembled soluble proteins for degradation. Cell discovery 2, 16040, doi:10.1038/celldisc.2016.40 (2016). 49. Jing, H. et al. A SIRT2-Selective Inhibitor Promotes c-Myc Oncoprotein Degradation and Exhibits Broad Anticancer Activity. Cancer cell 29, 297-310, doi:10.1016/j.ccell.2016.02.007 (2016). 50. Yang, J. Y. et al. A mouse model of shigellosis by intraperitoneal infection. The Journal of infectious diseases 209, 203-215, doi:10.1093/infdis/jit399 (2014). CHAPTER 4 LYSINE FATTY ACYLATION CATALYZED BY A FAMILY OF LEGIONELLA PNUEMOPHILA TOXINS This project is a collaboration initiated by Wenjie Zeng and Professor Yuxin Mao. Wenjie initially identified the sequence homology to RID, generated the Alphafold structure files, and generated L. pnuemophila strains. I contributed to all of other data presented in this chapter. ABSTRACT Bacterial pathogens secrete toxins to promote replication and survival during infection. Modifying host proteins is a common mechanism by which bacterial toxins modulate host physiology. Lysine fatty acylation has recently been found to be regulated by two secreted bacterial toxins from Vibrio cholerae and Shigella flexneri. Here, we identify three proteins from Legionella pnuemophila, lpg1096, lpg1387, and lpg1797, that can also catalyze lysine fatty acylation on host proteins. These proteins are predicted to have a highly similar core structure, including the active sites, with unique structural extensions. Lpg1387 has the highest acyltransferase activity. Proteomics screening for lpg1387 substrates revealed multiple proteins whose modification could be reversed by HDAC11 and SIRT2, including the mTORC1 activator RheB. Infection by WT and lpg1387 knockout strains verified lysine fatty acylation is modulated by lpg1387 and confirmed that RheB is a substrate. This work reveals a novel mode of host protein modification during L. pnuemophila infection and further cements lysine fatty acylation as a relevant protein modification during bacterial infection. INTRODUCTION Lysine acylation is a form of protein post-translation modification (PTM) that is important for the regulation of biological processes in every kingdom of life.1, 2 Acetylation, the best understood form of lysine acylation, is added to a protein by lysine acetyl transferases (KATs) and removed by lysine deacetylases (KDACs).3 In addition to acetylation, several other acyl groups are known to modify lysines by the same enzymes that catalyze acetylation, by unique enzymes, or through non-enzymatic mechanisms.1 Lysine fatty acylation (KFA) is the modification of lysines by long chain fatty acyl groups, typically of 14-18 carbons long. KFA is an understudied form of lysine acylation but has garnered increased attention lately due to the characterization of enzymes with lysine fatty acyl transferase (KFAT) or lysine defatty acylase (KDFA) activity.4 Many bacterial pathogens have been found to promote pathogenesis through modulating host PTMs.5 This is done in part by protein toxins that are secreted from the bacteria. Bacterial toxins have been shown to catalyze a diverse set of PTMs including ADP-ribosylation, ubiquitination, and polyglutamylation.6-8 Recently, the Rho-inactivating domain (RID) present in several Vibrio species and the IcsB toxin from Shigella flexneri were identified to catalyze KFA on host proteins.9, 10 These toxins function to inhibit actin polymerization and suppress inflammatory signaling for Vibrio and to inhibit autophagic destruction of Shigella flexneri. KFAT enzymes also make up a key component of RTX toxin operons, which form a pore in mammalian cell membranes and are present in several human pathogens.11 The abundance of KFAT enzymes in bacterial pathogens suggests that KFA may be a conserved mechanism of pathogenicity and warrants exploration for more such enzymes. CLUSTAL O(1.2.4) multiple sequence alignment lpg1387 --------------MIEVNIWLSTAFLFKKRIKHKFFGPLLASEDKGENVGHVNFTIEID 46 lpg0196 ------MLLGDFIMSLEIYTWKN-------GSKGLGFTKPVKSMLLGGNVGHAAVELTFP 47 lpg1797 MFKFIYCINQEHIMSITIHTWKG-------GTKGLGFKTPVKSTLFGGNVGHAALELTWP 53 : : * . * * : * * ****. . : lpg1387 ERTKTKN-SFDFIEQHGSELRAKKTLRVVPTKATNTSPPNIESSHLIPTIVRSDIVSHSF 105 lpg0196 ADAKGDELAKKYQDVPG----------LSISKRTEIVPEKQENGSYKPKEQVVYFVYFSW 97 lpg1797 ANKQGDELAKKYAQAHG----------VIISKRTEIVGETQEEGRIMPKERVTYFAYFSW 103 : .: : .: : * : :* *: . *.. *. :. .*: lpg1387 WPDARPTKSETIKGKAVKPKFKTHEEDMISEDSVTAMTIIHRKSALDEISREKMEDLDFL 165 lpg0196 WPGHTNGHHINSHREDLESEWRHEPAPKIKPEIQE------HLYGEN-IPTANKTN---- 146 lpg1797 WPGEQNGHYINSFNDDKDAEWRHEPDKKMHADWKN------QIYGPQGMSEENTTD---- 153 **. : . . . ::: . : : : . : : : : lpg1387 VEISDLEVNLEQRKLFMDDLKKLQLEKDELIKQQKQLTNSYQTQLKDLKGQLAKLELHLN 225 lpg0196 -------------------VTGLLI-SDKTITKVKEISHQSIKETTTIE----------- 175 lpg1797 -------------------VKGILI-KNKNIQKLKTIQHSTLTA--DIA----------- 180 :. : : .:: * : * : :. . : lpg1387 KTNGQITATERTINYLNKLKNPDPKTKTQLIELTEKLIKLKQEFETTINSQHEISALISK 285 lpg0196 ----------NDPA-YQQLKAEEAALQTELQQLIDKRKIYEQELARSGLEQRQPNK---- 220 lpg1797 ----------QDKA-YNELLQHKEKLDEEYKSLHAKAVAYQKEKENAIREEREINP---- 225 . ::* . . : .* * ::* : .::: . lpg1387 SELTYHEDLQQVEKKLQQIELAMNHRIEDLKKLDIKINGRDENDLKELQEEARRRKEYTS 345 lpg0196 ----------------------------ELCLTDAEI-----ARGYEISKELKRVAEQIK 247 lpg1797 ----------------------------NFELLDEDI-----SRMGDLPREFSILEAQLN 252 :: * .* :: .* . lpg1387 RKEQFIKSRDFTEGRQPDYSVTLPTAE--SGLTYYVDEIKILKAMQE--ERSQNYSFLFN 401 lpg0196 LCQIDFEERYLSVGEPPSSVIRIPTTLDHNAPTFALDAAGILDKMASLAQSTTAYSFYKF 307 lpg1797 LCIADFNERHLSSGKKPDSSVILPTSMDETSLNHTLDTESILREMITLANSEKAYNFTKF 312 ::.* :: *. *. : :**: .. .. :* ** * : *.* lpg1387 NCASSAKRCLLAGIDEKLRAKLKETGLGSKFFEISKVETCKSLRDWARTLDSKLTELNLR 461 lpg0196 NCSTSATQVVKAGISDELKNIMKTDGFNVEKASQTTIATPTSFRHFSQQVQTELILLNTN 367 lpg1797 NCSTSVCHIVKAGVDEKLKENLSDNGFNINRYVTSYITTPTNVNALGLKLQDALLKLELA 372 **::*. : : **:.::*: :. *:. : : : * .... . :: * *: lpg1387 SPAPTRGA--------------------------------------------------- 469 lpg0196 QALVEQQRAKTENTSEAPTTSTQSFKQKFQDTVKEKSEKAVREQDESEENRIGLTIS-- 424 lpg1797 EHTKEAEQVQKP-VANTISNRFNEFKSRLQAVVSERNAH----KDDTEEVTINNHNSLR 426 Figure 4. 1. Sequence alignment of L. pnuemophila toxins lpg1387, lpg0196, and lpg1797. Putative catalytic residues are colored in red. Legionella pnuemophila is a facultative intracellular pathogen that causes a pneumonia known as Legionnaire’s disease.12 Once internalized through phagocytosis, L. pnuemophila secretes over 300 effectors into the host cytoplasm that rewire host machinery to establish a replicative niche known as the Legionella-containing vacuole.13, 14 Here, we describe the identification of three L. pnuemophila effectors (lpg0196, lpg1387, lpg1797) with KFAT activity. We use SILAC proteomics to identify substrates and verify our findings with L. pnuemophila infection. RESULTS Identification of potential Rho-inactivating domain homologs in L. pnuemophila KFA is increasingly recognized as an important PTM catalyzed by bacterial toxins.9, 10 Toxins secreted by L. pnuemophila are known to modulate many PTMs so we wondered if any of them could catalyze KFA. We carried out a bioinformatics BLAST (NCBI) search using the V. vulnificus RID protein sequence to identify potential L. pnuemophila KFAT toxins. We identified three proteins (lpg0196, lpg1387, and lpg1797) with limited characterization which share sequence similarity with the V. vulnificus RID protein. Except for an apparent insertion in the interior of the lpg1387 sequence and an elongated C-terminal tail in lpg0196 and lpg1797, the three encoding proteins are quite similar and the encoding genes have been annotated as paralogs (Figure 4.1).15 RID utilizes a His-Asp-Cys catalytic triad.16 A putative catalytic histidine and cysteine align perfectly between the three proteins. While a catalytic aspartate does not align for lpg1387 in the primary sequence, subsequent structural studies identify aspartate residues positioned similarly in three dimensions. Little is known about these proteins, but Table 4. 1. Top five hits from an HHpred structural search for lpg1387 from residues 152- 355. The HDA1 coiled coil is formed from a single chain. The coiled coil structure in the other hits is formed from at least two chains. Protein Structural Feature PDB TSC1 Coiled coil helix 7DL2 HDA1 Coiled coil helix 6Z6O BICDR-1 Coiled coil helix 6F1T T1L Coiled coil helix 6GAO NudEL Coiled coil helix 2V71 Figure 4. 2. Structures of lpg0196, lpg1387, and 1797 predicted by Alphafold. Structures are overlapped to demonstrate structural similarities. The putative catalytic site and residues are shown below. lpg1387 knockout L. pnuemophila has recently been found to have enhanced biofilm formation, decreased mobility, and enhanced uptake my amoeba.17 We sought to biochemically characterize these proteins and determine whether they have KFAT activity. Structural prediction identifies unique lpg1387 coiled coil motif The protein sequences for lpg0196, lpg1387, and lpg1797 were initially submitted to the HHpred bioinformatic search tool to identify potential structural motifs.18 Unsurprisingly, all three have potential structural homology to the V. vulnificus RID. The interior insertion of lpg1387 also aligned with several proteins which contained an elongated α-helix coiled coil motif including a segment of the TSC complex, the GTPase-activating complex for RheB. We then used AlphaFold to predict the structure of lpg0196, lpg1797, and lpg0196 (Figure 4.2).19 Based on these predictions, all three proteins share a core structure which houses the putative catalytic residues in appropriate proximity to participate in a biochemical reaction. The structures also contain some unique aspects. lpg0196 and lpg1797 are predicted to manifest their elongated C-terminal tail as an α-helix connected to the core structure with a flexible linker. Additionally, the interior sequence of lpg1387 is predicted to manifest as an elongated α-helix coiled coil, just as predicted from the HHpred search. Together, these structural predictions suggest a shared or similar biochemical function for all three proteins, with potentially unique regulation or substrate preference. lpg1387, lpg1797, and lpg0196 have lysine fatty acyl transferase activity To determine if these proteins have KFAT activity, we utilized alkyne probe labeling. Alk14, a palmitic acid analog containing a terminal alkyne modification, was fed to cells co- transfected with one of the identified toxins and a previously identified RID substrate, Rac3. Rac3 protein was then immunoprecipitated and alkyne probe incorporation was measured by attaching an azide-conjugated fluorophore via click chemistry. Rac3 displayed strong fluorescence when overexpressed with any of the toxins with lpg1387 demonstrating the strongest activity (Figure 4.3). This signal was resistant to hydroxylamine treatment, which can hydrolyze thioester groups, demonstrating it is not cysteine palmitoylation. Modification of Rac3 resulted in a downward shift on SDS- PAGE which matches similar assays using RID.9 Additionally, mutation of the putative Figure 4. 3. Fluorescence gels and catalytic cysteine or histidine residues immunoblots of Rac3 cotransfected with the abolishes the activity of lpg1387. lpg1387 was indicated toxins to assay for fatty acylation also identified to have hydroxylamine-resistant using Alk14 labeling. HA = hydroxylamine. alkyne probe labeling while lpg0196 and HA lpg1387 = H38A mutant. CA lpg1387 = lpg1797 did not (Figure 4.4 A). This signal C403A mutant. was also abolished with catalytic dead mutants of lpg1387 demonstrating self-modification or modification by another lpg1387 molecule (Figure 4B). lpg1387 fatty acylation could be reduced by mutating a patch of lysine residues to arginine suggesting this modification is also on a lysine (Figure 4C). Mammalian lysine fatty acyl hydrolase enzymes were co-transfected with lpg1387 to see if they could remove lpg1387 KFA. HDAC11 and, to a lesser extent, SIRT2 could remove lpg1387 KFA (Figure 4D). However, mutant lpg1387 lacking fatty acylation had no decrease in activity suggesting this modification is not important for enzymatic activity (Figure 4E). Figure 4. 4. lpg1387 has KFA. A) In-gel fluorescence detection for lysine fatty acylation of indicated L. pneumophila proteins treated with or without hydroxylamine. Coomasie brilliant blue (CBB) stained gel was used to check protein loading. B) In-gel fluorescence detection of WT or catalytic dead mutants of lpg1387, H38A and C403A. C) In-gel fluorescence detection of WT or indicated lysine to arginine mutants of lpg1387 and with all four lysines mutated to arginine (4KR). D) In-gel fluorescence and immunoblots of lpg1387 co-transfected with indicated mammalian lysine fatty acyl hydrolase enzymes. E) In-gel fluorescence detection of flag-RheB lysine fatty acylation co-transfected with WT or 4KR lpg1387. HA = hydroxylamine. Figure 4. 5. SILAC proteomic screen for lpg1387 substrates. A) Experimental setup for processing proteomic samples. B) Flow chart for selecting high confidence substrate hits from the SILAC screen. Chemical proteomics reveals several host protein substrates of lpg1387 A SILAC (stable isotope labelling with amino acids in cell culture) proteomics approach was utilized to identify endogenous substrates for lpg1387-catalyzed lysine fatty acylation. HEK Table 4. 2. Proteins identified in SILAC proteomics screen for lpg1387 lysine fatty acylation substrates. Heavy to light ratio for both the forward are reverse experiment is shown along with a molecular detail for each protein. 293T cells transfected with either wild type or catalytically dead lpg1387 were fed Alk14 and lysates were reacted with biotin azide via click chemistry. Streptavidin enrichment was carried out and tryptic digests were analyzed via mass spectrometry (Figure 4.5A). Our analysis (Figure 4.5B) revealed 18 proteins to have increased modification with wild type lpg1387 overexpression (Figure 5B, Table 2). The list notably includes several C-terminally prenylated small GTPases which is in line with previously identified substrates for KFAT enzymes (RheB, Rap1B, Rab13, Rab8A, Rab10, RhoG, RalB, RalA). Indeed, several proteins were previously identified in a SILAC screen of IcsB substrates, a KFAT enzyme from S. flexneri.10 Also identified were proteins associated with proteasomal degradation (CUL4B, PSMC5), Na/K transport (ATP1B3, Table 4. 3. Selected gene ontology biological process identified to be enriched in proteomic hits from SILAC screen for lpg1387 substrates. ATP1A1), and septins (SEPT7, SEPT11). Gene ontology analysis revealed an enrichment of proteins involved in membrane trafficking, Na/K homeostasis, and Ras protein signal transduction (Table 3).20, 21 Validation of the proteomic data was carried out with fluorophore click chemistry on several substrates (Figure 4.6A). Furthermore, mutation of K178, the lysine closest to the prenylated cysteine on RheB, to arginine blocked modification by lpg1387 whereas K169R is still modified (Figure 4.6B - C). These data suggest a broad substrate potential for lpg1387 with preference for membrane-proximal lysines. Infection with L. pnuemophila affects lysine fatty acylation levels We next sought to confirm the activity of these KFAT enzymes during L. pnuemophila infection. HEK 293T cells were first transfected with the antibody receptor FcγRII and indicated flag- tagged substrates or FcγRII alone. The next day, transfected cells were fed Alk14 and infected with the indicated L. pnuemophila strains, opsonized by anti – L. pnuemohila antibody. Fatty acylation levels were then assayed using click chemistry and in-gel fluorescence. Examining global fatty acylation levels following infection revealed several fluorescence bands Figure 4. 6. KFA of lpg1387 substrates. A) In-gel fluorescence and immunoblots of flag- tagged SILAC hits co-transfected with lpg1387. Coomasie brilliant blue (CBB) stained gel was used to check protein loading. B) In-gel fluorescence of WT and lysine to arginine mutant flag- RheB co-transfected with lpg1387. C) Diagram of RheB C-terminal sequence showing position of mutated lysines and terminal farnesylation. D) In-gel fluorescence and immunoblots of flag- RheB co-transfected with lpg1387 and indicated lysine fatty acyl hydrolases. E) In-gel fluorescence and immunoblots of flag-Rac3 co-transfected with lpg1387 and indicated lysine fatty acyl hydrolases. that increased in intensity with either WT Lp02 or an Lp02 strain where lpg1387 was knocked out (red arrows). These bands were resistant to hydroxylamine treatment demonstrating they are not due to cysteine palmitoylation and could be due to the activity of lpg1797 or lpg0196 (Figure 4.7A). There were also fluorescence bands that increased in intensity with infection by WT Lp02, but not with Δlpg1387 Lp02 (blue arrows). This includes an intriguing band of approximately 25 kDa that could correspond to one or more of the small GTPases identified in the SILAC screen. We then looked at fatty acylation levels of RheB during infection. Overexpressed flag- RheB was immunoprecipitated and showed higher levels of fatty acylation in cells infected with WT but not Δlpg1387 Lp02 with or without hydroxylamine treatment demonstrating that lpg1387 can modify RheB with KFA during infection (Figure 4.7B). Moreover, Lp02 infection could not increase fatty acylation of RheB K178R, consistent with co-transfection results (Figure 4.7C). Together, this data demonstrates that L. pnuemophila can increase lysine fatty acylation during infection and that RheB is a substrate of the endogenous lpg1387. DISCUSSION Bacterial pathogens often modulate host molecular machinery to promote survival. The mechanisms used to do so are diverse. As shown in the above chapters, the catalysis of lysine fatty acylation is emerging as a new battleground in the fight between bacterial pathogen and host. In this work, we identified three additional KFAT toxins secreted by L. pnuemophila. The three proteins are similar in both amino acid sequence and predicted three-dimensional structure. However, they all demonstrate different activity with lpg1387 being the most active. It is possible the extended coiled coil motif in lpg1387 can participate in unique protein binding that enhances its activity or that the C-terminal helices in lpg0196 and lpg1797 play an inhibitory Figure 4. 7. L. pnuemophila infection modulates KFA levels. A) In-gel fluorescence of whole cell lysates from HEK 293T cells incubated with Alk14 and infected with WT or lpg1387 KO (Δ) Lp02. Coomasie brilliant blue (CBB) stained gel was used to check protein loading. Red arrows indicate bands that increase in intensity with infection of both strains. Blue arrows indicate bands that increase in intensity only with infection by WT Lp02. B) In-gel fluorescence and immunoblots of flag-RheB transfected into HEK 293T cells infected with WT or lpg1387 KO (Δ) Lp02 strains. C) In-gel fluorescence and immunoblots of WT or K178R flag-RheB transfected into HEK 293T cells infected with WT Lp02 strain. “+HA” samples were incubated at 95 oC for five minutes with 0.66 M hydroxylamine. role. We focused on lpg1387 due to its higher activity, but further exploration of lpg0196 and lpg1797 could prove enlightening. Do these enzymes have unique biological functions or are they functionally redundant? Screens to identify substrates of lpg0196 and lpg1797 would help answer this. A previous study showed that knockout of lpg1387 does not affect intracellular growth of L. pnuemophila. Could lpg0196 and lpg1797 compensate for loss of lpg1387? Double or triple knockout strains could be used to more comprehensively understand what role KFA plays during L. pneumphila infection. Unlike many other PTMs catalyzed by bacterial toxins, KFA can be reversed by host enzymes. SIRT2 is able to hydrolyze KFA catalyzed by IcsB from S. flexneri (Chapter 3) and HDAC11 can counteract RID from V. cholera (Chapter 2). Correspondingly, SIRT2 and HDAC11 promote the defense of S. flexneri and V. cholerae infections. Both SIRT2 and HDAC11 can remove lpg1387 catalyzed KFA though HDAC11 appears to be more effective. Infection studies with SIRT2 or HDAC11 knockout cells or mice could demonstrate if this is physiologically relevant. Many of the substrates of lpg1387 identified by SILAC proteomics are shared with other bacterial KFAT toxins, especially IcsB. IcsB functions to prevent destruction of S. flexneri through autophagy by modifying CHMP5. CHMP5 was not detected when screening for lpg1387 substrates, but modulating autophagy signaling could be beneficial for L. pnuemophila growth and survival. To this end, RheB, the top SILAC hit, is a key activator of the mTORC1 signaling complex which controls levels of autophagy along with protein synthesis and cell proliferation. L. pnuemophila has been shown to modulate mTORC1 and autophagy signals so RheB KFA could be a mechanism by which this occurs.22, 23 Further study of lpg1387 along with lpg0196 and lpg1797 would help to answer this question and could further reveal the biochemical, molecular, and physiological roles of KFA. METHODS Detection of individual protein fatty acylation by in-gel fluorescence. Indicated plasmids were transfected into HEK 293T cells using PEI transfection reagent following the manufacturer's protocol. The pCMV-Tag 4a or pEGFP C1 empty vector was used as the negative control. After overnight transfection, cells were treated with 50 μM Alk14 for 6 hours. The cells were washed 2x with ice cold PBS and collected by centrifugation at 1000 g for 5 min. Cells were then lysed in 1 mL 1% NP-40 lysis buffer (25 mM Tris-HCl, pH 7.8, 150 mM NaCl, 10% glycerol and 1% NP-40) with protease inhibitor cocktail (1:100 dilution) at 4 °C for 30 min. After centrifugation at 17,000 g for 30 min at 4 ˚C, the supernatant was collected and protein concentration was determined with Bradford assay (23200, Thermo Fisher). Lysates at a concentration of 0.5-2 mg/mL were incubated with 20 μL of anti-Flag affinity gel per mg of protein at 4 °C for 2 h. The affinity gel was washed three times with washing buffer (25 mM Tris-HCl, pH 7.8, 150 mM NaCl, 0.2% NP-40). Beads were dried with gel loading tips and resuspended in 20 μL of washing buffer. TAMRA-N3 (47130, Lumiprobe, 1 μL of 2 mM solution in DMF), Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (T2993, TCI chemicals, 1 μL of 10 mM solution in DMF), CuSO4 (1 μL of 40 mM solution in H2O) and Tris(2-carboxyethyl)phosphine (TCEP) (580560, Millipore, 1 μL of 40 mM solution in H2O) were added into the reaction mixture in the order listed. The click chemistry reaction was allowed to proceed at room temperature for 30 min. The reaction was quenched by adding 10 μL of 6x SDS loading dye to achieve a final concentration of ~2x and then boiled at 95 ˚C for 5 min. Where indicated, samples were treated with hydroxylamine to remove cysteine fatty acylation, 10 μL of the quenched reaction was treated with 2 uL of 4 M hydroxylamine (438227, Sigma, pH 7.4) and boiled at 95 ˚C for another 5 min. The samples were then resolved by 12% SDS- PAGE. The gel was then incubated in destaining buffer (50% CH3OH, 40% water and 10% acetic acid) by shaking 2-8 hours at 4˚C and then incubated in water for 2 hours. The gel was scanned to record fluorescence signal using a ChemiDoc MP (RioRad). Protein loading was checked by staining the gel with Coomassie Brilliant Blue (CBB) (B7920, Sigma) or carrying out a western blot on the samples. Detection of global protein fatty acylation by in-gel fluorescence After indicated treatments, cells were collected and lysed in 1% NP40 as above. Lysates at a concentration of 0.5-2 mg/mL were brought up to 50 μL and mixed with 2.5 μL of TAMRA-N3, TBTA, CuSO4, and TCEP at the above concentrations. The click chemistry reaction was carried out at room temperature for 30 min and then quenched by precipitating the proteins. 200 μL cold MeOH was added to the reaction mixture followed by 75 μL cold CHCl3 and 150 μL cold water. The sample was vortexed then centrifuged at 17,000 g for 15 min at 4 ˚C. The upper aqueous phase was removed and the pellet washed by adding 1 mL cold MeOH and centrifuging at 17,000 g for 10 min at 4 ˚C. After a second identical wash, the supernatant was removed and the pellet was allowed to dry for 30 min. Proteins were resolubilized by adding 50 μL 4% SDS buffer (4% w/v SDS, 150 mM NaCL, 25 mM TEA pH 7.5) and 10 μL 6x load dye followed by boiling at 95 ˚C for 5 min. Two 10 μL aliquots were placed in new tubes and treated with 2 μL water or 2 μL 4M NH2OH followed by boiling at 95 ˚C for 5 min. Samples were then resolved on SDS-PAGE and visualized as above. Detection of endogenous protein fatty acylation by biotin click chemistry Cells were treated as indicated and fed 50 μM Alk14 was added for 6 hours. Cells were washed, collected, and lysed in 1% NP-40 lysis buffer as above. Lysates were normalized to equal protein level at no more than 2 mg/mL protein concentration. Lysates were incubated with Biotin-N3 (final concentration of 25 μM), Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (final concentration of 500 μM), CuSO4 (final concentration of 2.5 mM), and Tris(2- carboxyethyl)phosphine (final concentration of 2.5 mM) for 60 min at room temperature. Proteins were precipitated by adding 4 volumes of cold MeOH, 1 volume cold CHCl3, then 3 volumes cold water followed by a 30 min centrifuge at 4696 g at 4 ˚C. The upper aqueous phase was removed, and the protein pellet was washed with 8 volumes of cold MeOH and centrifuged for 15 min at 4696 g at 4 ˚C. The MeOH was removed, and the pellet allowed to dry at room temperature for 20 min. Samples were then redissolved in 100 mM Tris–HCl pH 7.2, 150 mM NaCl, 5 mM EDTA, 2.5% SDS, 8 M urea. Solubilized protein solution was then diluted 10x with PBS and 20 μL of streptavidin-agarose beads (ThermoFisher) were added. Streptavidin was incubated for 2 hours at room temperature with rocking. Streptavidin beads were collected by centrifugation at 2000 g then washed two times with 1 mL 1% SDS in PBS and 1 time with PBS. The beads were dried with a gel loading tip and 50 μL of 2x loading dye was added followed by boiling at 95 ˚C for 5 min. Levels of protein fatty acylation were measured via western blot. SILAC labeling for lpg1387 lysine fatty acylation substrates HEK 293T cells were cultured in SILAC DMEM supplemented with 10% dialyzed FBS and either normal lysine and arginine or heavy ([13C6, 15N2]-L-lysine, and [ 13C , 156 N4]-L-arginine) lysine and arginine. WT or H38A mutant lpg1387 was transfected for 24 hours and 50 μM Alk14 was added for 6 hours. Biotin click chemistry was carried out as above. Proteins were precipitated by adding 4 volumes of cold MeOH, 1 volume cold CHCl3, then 3 volumes cold water followed by a 30 min centrifuge at 4696 g at 4 ˚C. The upper aqueous phase was removed, and the protein pellet was washed with 8 volumes of cold MeOH and centrifuged for 15 min at 4696 g at 4 ˚C. The MeOH was removed, and the pellet allowed to dry at room temperature for 20 min. Samples were then redissolved in 100 mM Tris–HCl pH 7.2, 150 mM NaCl, 5 mM EDTA, 2.5% SDS, 8 M urea. Solubilized protein solution was then diluted 10x with PBS and 100 μL of streptavidin-agarose beads were added. Streptavidin was incubated for 2 hours at room temperature with rocking. Streptavidin beads were collected by centrifugation at 2000 g then washed three times with 1 mL of 0.2% SDS in PBS, followed by three washes with 1 mL PBS, three washes with 1 mL of 20 mM Tris–HCl, 500 mM KCl, pH 7.4, and finally three washes with 1 mL of 20 mM Tris–HCl, pH 7.4. The beads were dried with a gel loading tip and proteins were reduced by adding 400 μL of a PBS solution containing 6 M urea and 9.5 mM TCEP and incubating for 20 min at 37 ˚C with gentle rotation. For protein alkylation, 20 μL of a fresh solution of 400 mM iodoacetamide (in water) was added to the bead suspension and incubated for 20 min at 37 ˚C with gentle rotation. The supernatant was removed and the beads were washed 1 mL of 2 M urea in PBS, and then incubated with 2 μg of trypsin in 200 μL of 2 M urea in PBS with 1 mM CaCl2 at 37 ˚C overnight with gentle rotation. Following trypsin digestion, beads were centrifuged at 1000 x g, room temperature and the supernatant was transferred to a 1.5-mL tube. The beads were washed twice with 300 μL of water. Washes were combined with the supernatant and the solution diluted to 1 mL by adding water. pH was adjusted to ~2 by adding 15 μL of 10% trifluoroacetic acid (TFA, Sigma). A Sep-Pak Vac C18 cartridge (Waters) was conditioned by passing 1 mL of 90% methanol–0.1% TFA through the cartridge three times. The cartridge was equilibrated by passing 1 mL of 0.1% TFA through three times. The sample was added to the cartridge slowly, at a rate of approximately 1 drop/s. The loaded cartridge was washed with 1 mL of 0.1% TFA three times. The peptides were then eluted by passing 1 mL of 80% acetonitrile–0.1% TFA through the cartridge once. The cartridge was dried with nitrogen to ensure all peptides were eluted. Eluted peptides were flash frozen in liquid nitrogen and lyophilized. Protein Identification by nano LC/MS/MS Analysis: The tryptic digests of SILAC samples were reconstituted in 2% acetonitrile with 0.5% formic acid (FA) to a final concentration of 0.02 µg/µl for nanoLC-ESI-MS/MS analysis. The analysis was carried out using an Orbitrap FusionTM TribridTM (Thermo-Fisher Scientific, San Jose, CA) mass spectrometer equipped with a nanospray Flex Ion Source, and coupled with a Dionex UltiMate 3000 RSLCnano system (Thermo, Sunnyvale, CA)24, 25. Peptide samples (10 µl) were injected onto a PepMap C-18 RP nano trapping column (5 µm, 100 µm i.d. x 20 mm) with nanoViper Fittings at 20 µL/min flow rate for rapid sample loading and then separated on a PepMap C-18 RP nano column (2 µm, 75 µm i.d. x 25 cm) at 35 °C. The tryptic peptides were eluted in a 90 min gradient of 5% to 35% ACN in 0.1% formic acid at 300 nL/min, followed by a 7 min ramping to 90% ACN-0.1% FA and an 8 min hold at 90% ACN-0.1% FA. The column was re-equilibrated with 0.1% FA for 25 min prior to the next run. The Orbitrap Fusion was operated in positive ion mode with spray voltage set at 1.1 kV and source temperature at 275°C. External calibration for FT, IT and quadrupole mass analyzers was performed. In data-dependent acquisition (DDA) analysis, the instrument was operated using FT mass analyzer in MS scan to select precursor ions followed by 3 second “Top Speed” data-dependent CID ion trap MS/MS scans at 1.6 m/z quadrupole isolation for precursor peptides with multiple charged ions above a threshold ion count of 10,000 and normalized collision energy of 30%. MS survey scans at a resolving power of 120,000 (fwhm at m/z 200), for the mass range of m/z 375-1575. Dynamic exclusion parameters were set at 50 s of exclusion duration with ±10 ppm exclusion mass width. All data were acquired under Xcalibur 4.3 operation software (Thermo-Fisher Scientific). MS data analysis: The DDA raw files with MS and MS/MS spectra were subjected to database searches using Proteome Discoverer (PD) 2.4 software (Thermo Fisher Scientific, Bremen, Germany) with the Sequest HT algorithm. The PD 2.4 processing workflow containing an additional node of Minora Feature Detector for precursor ion-based quantification was used for protein identification and relative quantitation of identified peptides and their modified forms. The database search was conducted against a Homo sapiens Uniprot database. The peptide precursor tolerance was set to 10 ppm and fragment ion tolerance was set to 0.6 Da. Variable modifications of methionine oxidation; deamidation of asparagine/glutamine; SILAC heavy: R10 (10.008 Da) and K8 (8.014 Da) and light labeling on R and K; palmitoylation plus biotin on K, protein N-terminus and fixed modification of cysteine carbamidomethylation were set for the database search. Only high confidence peptides defined by Sequest HT with a 1% FDR by Percolator were considered for confident peptide identification. Relative quantitation of identified proteins of heavy versus light labeled was determined by the Precursor Ions Quantifier node for SILAC 2plex (Arg10, Lys8) within Precursor Quan workflow in PD 2.4. The precursor abundance intensity for each peptide identified by MS/MS in each replicate was automatically determined and their unique plus razor peptides for each protein in each replicate were used for calculating the heavy/light ratios by pairwise ratio based for all identified proteins without normalization. The final protein group list was further filtered with 5ppm for identified peptides and two peptides per protein in which only #1-ranked peptides within top scored proteins were used. Generation of lpg1387 knockout Construction of the lpg1387 knockout plasmid was done by amplifying two regions of DNA flanking lpg1387, with 45 overlapping bases using the following primers: lpg1387KO_Upstream_SalI_Fwd (5’- GGCGGTCGACGAAGCCTAATGATTATCTCAATCAAGC-3’), lpg1387KO_Upstream_BamHI_Rvs (5’- GATGGATCCCTTAAACAAAAAAGCAGTTGATAACCAAATATTTACTTCAATCAT-3’), lpg1387KO_Downstream_BamHI_Fwd (5’- CGGGGGATCCTTGACAGAACTTAATCTTCGATCTCCTGCTCCTACCCGAGGTGCT-3’), and lpg1387KO_Downstream_SacI_Rvs (5’- CCGGGAGCTCCAAAACCATTTTTCCAATAGGTGATTTTAC-3’). The upstream amplicon was digested with SalI and BamHI and the downstream amplicon was digested with BamHI and SacI according to manufacturer recommendations. Restriction enzymes were purchased from NEB. The vector pSR47S was digested with SalI and SacI. Then, the upstream, downstream, and vector digests were incubated together and triple-ligated using T4 DNA ligase (NEB) at room temperature for 1 hour. The ligated product was transformed into E. coli DH5α λpir, and colonies were screened for positives. The lpg1387 knockout plasmid was introduced into Legionella using triparental mating. Legionella pneumophila WT strain (Lp02) was streaked to single colony on CYET (Charcoal- buffered Yeast Extract with Thymidine) agar plate at 37oC for 3-4 days. A single colony was then used to patched onto a fresh CYET plate. At the same time, E. coli DH5α strain transformed with the pHelper plasmid was patched onto LB-chloramphenicol plate at 37oC for 1 day. E. coli DH5α λpir transformed with the correct lpg1387 knockout plasmid (pLpg1387KO) was inoculated in LB-Kan medium overnight at 37oC. On the day of triparental mating, cells from Lp02 and pHelper patches were scraped with a sterile wooden toothpick and added to 200uL of overnight culture of DH5α λpir-pLpg1387KO. This mixture was then centrifuged at 16,000 RPM at room temperature for 1 minute using a microcentrifuge, and about 150uL of liquid was removed. The remaining cell pellet was resuspended using residual liquid and spotted onto CYET plate and allowed the triparental mating to occur for 4 hours at 37oC. Following triparental mating, the entirety of the cells were streaked to single colonies onto CYET + 50ug/mL of kanamycin + 50ug/mL of streptomycin. And the plate was incubated at 37oC for 4-6 days until single colonies were formed. Single colonies from this plate were then streaked onto CYET + 50ug/mL of kanamycin and grown at 37oC for 4-6 days until single colonies formed. Single colonies from this plate were then streaked onto CYET (with no antibiotics) and grown at 37oC for 4-6 days until single colonies formed. Finally, single colonies from this plate were streaked onto CYET + 5% sucrose to select for individual bacterium that have the correct double-crossover lpg1387 deletion event. To screen for positive lpg1387 knockout Lp02 clones, about 40 colonies from the CYET + 5% sucrose plate were patched onto a fresh CYET plate. Colony PCR was performed to check for positive clones using the following forward and reverse primer pairs: 1. lpg1387KO_Upstream_SalI_F and lpg1387KO_Downstream_SacI_R a. WT PCR product = 3800bp b. KO plasmid and KO strain PCR product = 2400bp 2. lpg1387_BamHI_F and lpg1387_XhoI_R a. WT PCR product = 1400bp b. KO plasmid and KO strain PCR product = 100bp 3. lpg1387_BamHI_F and lpg1387KO_Downstream_SacI_R a. WT PCR product = 2600bp b. KO plasmid and KO strain PCR product = 1500bp 4. lpg1387KO_Upstream_SalI_F and lpg1387_XhoI_R a. WT PCR product = 2600bp b. KO plasmid and KO strain PCR product = 1500bp Infection of HEK 293T cells by L. pnuemophila L. pnuemophila colonies were patched onto CYET plates and incubated at 37oC for 2-3 days until a thick bacterial lawn was formed. Meanwhile, HEK 293T cells were seeded into 6-well plates. At 70% confluence HEK 293T cells were transfected with FcγRII and additional KFA substrates where indicated for 24 hours. The next day, L. pnuemophila lawns were resuspended in AYET to OD600 = 2.8 and incubated with shaking at 37 oC for 1.5 hours. During incubation, HEK 293T media was changed to include 50 uM Alk14. L. pnuemophila cultures, now at OD600 ~ 3.0 were incubated 1:500 with anti-L. pnuemophila (Abcam) for 20 min at 37oC. 15 uL of opsonized bacteria (MOI ~ 30) were added to each well of HEK 293T cells and infection was initiated by centrifuging for 5 min at 3000 rpm. Infection was carried out for 6 hours and samples were analyzed for fatty acylation levels as outlined above. ACKNOWLEDGEMENTS We thank the Proteomics and Metabolomics Facility of Cornell University for providing the mass spectrometry data and NIH SIG grant 1S10 OD017992-01 support for the Orbitrap Fusion mass spectrometer. REFERENCES 1. Baldensperger, T., and Glomb, M. A. (2021) Pathways of Non-enzymatic Lysine Acylation, Front Cell Dev Biol 9, 664553. 2. Wang, M., and Lin, H. (2021) Understanding the Function of Mammalian Sirtuins and Protein Lysine Acylation, Annu Rev Biochem. 3. Sabari, B. R., Zhang, D., Allis, C. D., and Zhao, Y. (2017) Metabolic regulation of gene expression through histone acylations, Nat Rev Mol Cell Biol 18, 90-101. 4. Komaniecki, G., and Lin, H. (2021) Lysine Fatty Acylation: Regulatory Enzymes, Research Tools, and Biological Function, Front Cell Dev Biol 9, 717503. 5. Galan, J. E. (2009) Common themes in the design and function of bacterial effectors, Cell Host Microbe 5, 571-579. 6. Barbieri, J. T. (2000) Pseudomonas aeruginosa exoenzyme S, a bifunctional type-III secreted cytotoxin, Int J Med Microbiol 290, 381-387. 7. Zhu, Y., Li, H., Hu, L., Wang, J., Zhou, Y., Pang, Z., Liu, L., and Shao, F. (2008) Structure of a Shigella effector reveals a new class of ubiquitin ligases, Nat Struct Mol Biol 15, 1302- 1308. 8. Sulpizio, A., Minelli, M. E., Wan, M., Burrowes, P. D., Wu, X., Sanford, E. J., Shin, J. H., Williams, B. C., Goldberg, M. L., Smolka, M. B., and Mao, Y. (2019) Protein polyglutamylation catalyzed by the bacterial calmodulin-dependent pseudokinase SidJ, Elife 8. 9. Zhou, Y., Huang, C., Yin, L., Wan, M., Wang, X., Li, L., Liu, Y., Wang, Z., Fu, P., Zhang, N., Chen, S., Liu, X., Shao, F., and Zhu, Y. (2017) N(epsilon)-Fatty acylation of Rho GTPases by a MARTX toxin effector, Science 358, 528-531. 10. Liu, W., Zhou, Y., Peng, T., Zhou, P., Ding, X., Li, Z., Zhong, H., Xu, Y., Chen, S., Hang, H. C., and Shao, F. (2018) N(epsilon)-fatty acylation of multiple membrane-associated proteins by Shigella IcsB effector to modulate host function, Nat Microbiol. 11. Benz, R. (2016) Channel formation by RTX-toxins of pathogenic bacteria: Basis of their biological activity, Biochim Biophys Acta 1858, 526-537. 12. McDade, J. E., Shepard, C. C., Fraser, D. W., Tsai, T. R., Redus, M. A., and Dowdle, W. R. (1977) Legionnaires' disease: isolation of a bacterium and demonstration of its role in other respiratory disease, N Engl J Med 297, 1197-1203. 13. Zhu, W., Banga, S., Tan, Y., Zheng, C., Stephenson, R., Gately, J., and Luo, Z. Q. (2011) Comprehensive identification of protein substrates of the Dot/Icm type IV transporter of Legionella pneumophila, PLoS One 6, e17638. 14. Isberg, R. R., O'Connor, T. J., and Heidtman, M. (2009) The Legionella pneumophila replication vacuole: making a cosy niche inside host cells, Nat Rev Microbiol 7, 13-24. 15. Huang, L., Boyd, D., Amyot, W. M., Hempstead, A. D., Luo, Z. Q., O'Connor, T. J., Chen, C., Machner, M., Montminy, T., and Isberg, R. R. (2011) The E Block motif is associated with Legionella pneumophila translocated substrates, Cell Microbiol 13, 227-245. 16. Ahrens, S., Geissler, B., and Satchell, K. J. (2013) Identification of a His-Asp-Cys catalytic triad essential for function of the Rho inactivation domain (RID) of Vibrio cholerae MARTX toxin, J Biol Chem 288, 1397-1408. 17. Marin, C., Kumova, O. K., and Ninio, S. (2022) Characterization of a Novel Regulator of Biofilm Formation in the Pathogen Legionella pneumophila, Biomolecules 12. 18. Zimmermann, L., Stephens, A., Nam, S. Z., Rau, D., Kubler, J., Lozajic, M., Gabler, F., Soding, J., Lupas, A. N., and Alva, V. (2018) A Completely Reimplemented MPI Bioinformatics Toolkit with a New HHpred Server at its Core, J Mol Biol 430, 2237-2243. 19. Jumper, J., Evans, R., Pritzel, A., Green, T., Figurnov, M., Ronneberger, O., Tunyasuvunakool, K., Bates, R., Zidek, A., Potapenko, A., Bridgland, A., Meyer, C., Kohl, S. A. A., Ballard, A. J., Cowie, A., Romera-Paredes, B., Nikolov, S., Jain, R., Adler, J., Back, T., Petersen, S., Reiman, D., Clancy, E., Zielinski, M., Steinegger, M., Pacholska, M., Berghammer, T., Bodenstein, S., Silver, D., Vinyals, O., Senior, A. W., Kavukcuoglu, K., Kohli, P., and Hassabis, D. (2021) Highly accurate protein structure prediction with AlphaFold, Nature 596, 583-589. 20. Ashburner, M., Ball, C. A., Blake, J. A., Botstein, D., Butler, H., Cherry, J. M., Davis, A. P., Dolinski, K., Dwight, S. S., Eppig, J. T., Harris, M. A., Hill, D. P., Issel-Tarver, L., Kasarskis, A., Lewis, S., Matese, J. C., Richardson, J. E., Ringwald, M., Rubin, G. M., and Sherlock, G. (2000) Gene ontology: tool for the unification of biology. The Gene Ontology Consortium, Nat Genet 25, 25-29. 21. Gene Ontology, C. (2021) The Gene Ontology resource: enriching a GOld mine, Nucleic Acids Res 49, D325-D334. 22. De Leon, J. A., Qiu, J. Z., Nicolai, C. J., Counihan, J. L., Barry, K. C., Xu, L., Lawrence, R. E., Castellano, B. M., Zoncu, R., Nomura, D. K., Luo, Z. Q., and Vance, R. E. (2017) Positive and Negative Regulation of the Master Metabolic Regulator mTORC1 by Two Families of Legionella pneumophila Effectors, Cell Reports 21, 2031-2038. 23. Choy, A., Dancourt, J., Mugo, B., O'Connor, T. J., Isberg, R. R., Melia, T. J., and Roy, C. R. (2012) The Legionella Effector RavZ Inhibits Host Autophagy Through Irreversible Atg8 Deconjugation, Science 338, 1072-1076. 24. Yang, Y., Anderson, E., and Zhang, S. (2018) Evaluation of six sample preparation procedures for qualitative and quantitative proteomics analysis of milk fat globule membrane, Electrophoresis 39, 2332-2339. 25. Harman, R. M., He, M. K., Zhang, S., and GR, V. D. W. (2018) Plasminogen activator inhibitor-1 and tenascin-C secreted by equine mesenchymal stromal cells stimulate dermal fibroblast migration in vitro and contribute to wound healing in vivo, Cytotherapy 20, 1061- 1076. CHAPTER 5 SUMMARY AND FUTURE DIRECTIONS SUMMARY This thesis describes work on characterizing the importance of KFA during infection by three unique bacterial pathogens (Figure 5.1). Bacteria can utilize an incredibly diverse armory of toxin proteins to promote replication in the host and to prevent elimination by the immune system.1 V. cholerae replicate extracellularly in the crypts between the intestinal villi where they secrete a large, multifunctional toxin that includes a KFAT domain, RID.2 S. flexneri replicates intracellularly in a variety of cell types in the digestive track.3 The KFAT toxin IcsB is secreted directly into the mammalian cell cytoplasm to carry out its function. L. pnuemophila infect lung alveolar macrophages where they establish a replicative niche in a vacuole inside the macrophage.4 The L. pnuemophila KFAT toxins are secreted across the vacuole membrane into the host cytoplasm. Despite these diverse mechanisms of infection, all these bacteria have evolved to catalyze KFA, emphasizing its functional utility. The presence of mammalian enzymes that can hydrolyze this moiety thus provides a compelling toxin-antitoxin battleground. We found that RID, the V. cholerae KFAT, has lysine fatty acylation. HDAC11 can hydrolyze KFA on RID, subsequently decreasing its activity towards its substrates. In this way, HDAC11 promotes bacterial phagocytosis and clearance of V. cholerae infection in mice. We found that SIRT2 can serve a similar function during S. flexneri infection. SIRT2 is upregulated by Golgi stress during S. flexneri infection and can then reverse KFA catalyzed by IcsB. This promotes the elimination of S. flexneri by autophagic machinery and decreases bacterial load during infection. Finally, we discovered three new toxins from L. pnuemophila that can catalyze KFA. HDAC11 can reverse KFA catalyzed lpg1387 so further work is warranted in order to determine if HDAC11 can be beneficial during L. pnuemophila infection. Figure 5. 1. Summary of KFA regulation reactions discussed in this work. FUTURE DIRECTIONS Further characterization of V. cholerae and L. pnuemophila KFAT toxins The molecular function of RID was largely determined before the discovery of its enzymatic activity.5 When RID was found to catalyze KFA on RhoA-family, this provided a compelling mechanism for its molecular function. However, a systematic identification of RID substrates has not been done. While RhoA-family GTPases may indeed be the primary substrate, overexpressed RID is able to modify CHMP5, a IcsB substrate, so it is worth identifying any additional substrates (Figure 5.2A). To do this, WT and RID catalytic dead mutant V. cholerae should be used to infect mammalian cells incubated with Alk14. Then, a proteomics screen like that in Chapter 4 can be carried out. Identified substrates can then be further characterized for physiological relevance. Increased RID substrate fatty acylation following V. cholerae infection is clear using in-gel fluorescence following IP. However, when carrying out biotin-azide click chemistry followed by streptavidin enrichment, substrate fatty acylation apparently decreases (Figure 5.2B). This assay-specific contradiction would confound screen for endogenous substrates and thus must be understood. Some MARTX toxins contain a Ras/Rap1-specific endopeptidase (RRSP) domain. Cleavage of Ras by this domain results in two peptides that stay associated until denatured. This could explain the discrepancy in Figure 5.2B as proteins are denatured during a biotin click chemistry experiment, but not for in-gel fluoresence. The O1 El Tor E7946 V. cholerae strain used in this study does not have an MARTX RRSP domain, but it is possible a similar mechanism is taking place. Understanding this discrepancy would be crucial before identifying additional RID substrates. Figure 5. 2. A) In-gel fluorescence and immunoblots of CHMP5 co-transfected with RID. B) In- gel fluorescence of Rac3 in cells infected with V. cholerae. Panels at left correspond to Alk14- modified Rac3 reacted with TAMRA-N3 for in-gel fluorescence. Panels at right correspond to Alk14-modified Rac3 reacted with biotin-N3 then precipitated, resolubilized and purified with streptavidin. In Chapter 4, three toxins were identified to have KFAT activity. However, much additional work needs to be done to fully understand the role of these proteins. lpg1387 was identified to be able to catalyze KFA on several proteins, but it is not yet clear if any of the substrates are physiologically significant. RheB was the top SILAC hit and it is modified during infection. RheB is known to activate mTORC1, an essential regulator of cell growth in response to nutrients.6 How does lpg1387 affect RheB function? KFA is known to decrease GTP loading of other small GTPases.7 Is this true for RheB as well? How does this affect RheB function during L. pnuemophila infection? Detailed mechanistic studies are necessary to understand the functional impact of lpg1387 modification of RheB. lpg1387 was screened for substates via plasmid transfection. L. pnuemophila secretes over 300 different effectors into the host cytoplasm, many of which interact or modify each other.8 To more accurately identify physiological substrates, proteomics screens should be done following infection by WT and Δlpg1387 strains. Moreover, substrates of lpg0196 and lpg1797 should be similarly identified. With substrate identities for all three enzymes, the true physiological role of KFA during L. pnuemophila infection can then be determined. lpg1387 was the most active in an overexpression system. Is this also true during a bacterial infection? Are each of the KFATs functional unique or are they redundant? How do each of these enzymes affect intracellular proliferation? These questions and more can be answered by generating knockout strains and identifying proteins substrates. Further exploration of HDAC11’s role during bacterial infection HDAC11 has the highest activity for any known KDFA enzyme.9 In Chapter 2, HDAC11 was identified to hydrolyze lysine fatty acylation on RID to decrease its activity towards mammalian substrates. In Chapter 4, HDAC11 was identified to efficiently decrease lpg1387 substrate KFA as well as KFA on lpg1387 itself. Unlike RID, KFA on lpg1387 does not appear to affect its activity. Not mentioned in Chapter 3, but in a similar vein, HDAC11 can decrease KFA on both IcsB and CHMP5 (Figure 5.3). HDAC11 was found to be beneficial during V. cholerae infection. Might it also help to fight S. flexneri and L. pnuemophila infection? Interestingly, HDAC11 knockout increased phagocytosis of MARTX knockout V. cholerae. This is the opposite trend from the WT bacteria. Does MARTX somehow regulate HDAC11 activity? What role does HDAC11 play during infection by a bacteria that does not have a KFAT toxin? Infection of HDAC11 knockout mice by a variety of different bacteria can help answer these important questions and understand the role of HDAC11 in bacterial infections. Discovery of additional KFAT toxins and characterization of Burkholderia BopA Recent literature and the work outlined above has demonstrated the physiological significance of KFA during infection by multiple species of pathogenic bacteria. Identified bacterial KFAT toxins have been found to reduce Figure 5. 3. In-gel fluorescence and inflammatory signaling and prevent phagocytic immunoblots of CHMP5 and IcsB co- destruction, two crucial steps in fighting a bacterial transfected with HDAC11. infection.10 It is feasible that many other bacteria may take advantage of this mechanism as part of their pathogenesis. Indeed several proteins, some from human pathogens, have been found to share sequence similarity.11 To this end, a Burkholderia spp. toxin, BopA, has homology both to RID and especially to IcsB, the KFAT toxins from V. cholerae and S. flexneri, respectively. Homology to these proteins appears to be reliable predictor for identifying additional KFAT enzymes suggesting that BopA could be a KFAT toxin. BopA is present in many Burkholderia species including B. pseudomallei, a dangerous pathogen classified as a Category B bioterrorism threat by the CDC. B. pseudomallei infection proceeds in a similar manner to S. flexneri. B. pseudomallei can invade a variety of different cells type through endocytosis where it escapes the endosome to proliferate intracellularly. Stimulating autophagy has been found to suppress B. pseudomallei intracellular growth.12 Like IcsB and RID, BopA has been found to have an anti-autophagy effect, thereby promoting intracellular proliferation.13 Given the shared homology and function of BopA and previously identified KFATs, it is likely that BopA has KFAT activity. REFERENCES 1. Galan, J. E. (2009) Common themes in the design and function of bacterial effectors, Cell Host Microbe 5, 571-579. 2. Muanprasat, C., and Chatsudthipong, V. (2013) Cholera: pathophysiology and emerging therapeutic targets, Future Med Chem 5, 781-798. 3. Jennison, A. V., and Verma, N. K. (2004) Shigella flexneri infection: pathogenesis and vaccine development, FEMS Microbiol Rev 28, 43-58. 4. Cianciotto, N. P. (2001) Pathogenicity of Legionella pneumophila, Int J Med Microbiol 291, 331-343. 5. Sheahan, K. L., and Satchell, K. J. (2007) Inactivation of small Rho GTPases by the multifunctional RTX toxin from Vibrio cholerae, Cell Microbiol 9, 1324-1335. 6. Valvezan, A. J., and Manning, B. D. (2019) Molecular logic of mTORC1 signalling as a metabolic rheostat, Nat Metab 1, 321-333. 7. Zhou, Y., Huang, C., Yin, L., Wan, M., Wang, X., Li, L., Liu, Y., Wang, Z., Fu, P., Zhang, N., Chen, S., Liu, X., Shao, F., and Zhu, Y. (2017) N(epsilon)-Fatty acylation of Rho GTPases by a MARTX toxin effector, Science 358, 528-531. 8. Zhu, W., Banga, S., Tan, Y., Zheng, C., Stephenson, R., Gately, J., and Luo, Z. Q. (2011) Comprehensive identification of protein substrates of the Dot/Icm type IV transporter of Legionella pneumophila, PLoS One 6, e17638. 9. Cao, J., Sun, L., Aramsangtienchai, P., Spiegelman, N. A., Zhang, X., Huang, W., Seto, E., and Lin, H. (2019) HDAC11 regulates type I interferon signaling through defatty acylation of SHMT2, Proc Natl Acad Sci U S A 116, 5487-5492. 10. Komaniecki, G., and Lin, H. (2021) Lysine Fatty Acylation: Regulatory Enzymes, Research Tools, and Biological Function, Front Cell Dev Biol 9, 717503. 11. Pei, J., and Grishin, N. V. (2009) The Rho GTPase inactivation domain in Vibrio cholerae MARTX toxin has a circularly permuted papain-like thiol protease fold, Proteins 77, 413- 419. 12. Cullinane, M., Gong, L., Li, X., Lazar-Adler, N., Tra, T., Wolvetang, E., Prescott, M., Boyce, J. D., Devenish, R. J., and Adler, B. (2008) Stimulation of autophagy suppresses the intracellular survival of Burkholderia pseudomallei in mammalian cell lines, Autophagy 4, 744-753. 13. Gong, L., Cullinane, M., Treerat, P., Ramm, G., Prescott, M., Adler, B., Boyce, J. D., and Devenish, R. J. (2011) The Burkholderia pseudomallei type III secretion system and BopA are required for evasion of LC3-associated phagocytosis, PLoS One 6, e17852.