CURRICULUM VITAE Erin E. Mcclure Carroll Department of Microbiology

XIAP-p47 Pairing Activates the Immune Deficiency Pathway in the Lyme Disease Tick Ixodes scapularis

Item Type dissertation

Authors McClure Carroll, Erin

Publication Date 2019

Abstract Globally, vector-borne diseases account for 17% of all infectious diseases. Most vectors are blood-feeding arthropods, which transmit bacterial, viral, and parasitic diseases to humans and animals. The tick Ixodes scapularis transmits seven pathogens...

Keywords IMD pathway; ubiquitin ligase; vector biology; Lyme Disease; Rickettsia Infections; Ticks

Download date 25/09/2021 20:55:28

Link to Item http://hdl.handle.net/10713/9604 CURRICULUM VITAE

Erin E. McClure Carroll

Department of Microbiology and Immunology 685 W. Baltimore St., HSFI Room 380 Baltimore, MD 21201

[email protected]

Doctor of Philosophy to be conferred on May 17, 2019

Education

2014 – 2019 University of Maryland Baltimore, Baltimore, MD Graduate Program in Life Sciences Molecular Microbiology and Immunology Thesis advisor: Joao Pedra, Ph.D.

2009 – 2013 Juniata College, Huntingdon, PA B.S. Biochemistry and minor in German, magna cum laude Bachelor thesis: “The temporal dysbiosis of the gut microbiota in IBD” Thesis advisor: Regina Lamendella, Ph.D.

Professional Positions Held

2019 – present Medical Writer Regulatory & Quality Solutions, LLC

2014 - 2019 Graduate Research Assistant Department of Microbiology and Immunology University of Maryland School of Medicine Thesis advisor: Joao Pedra, Ph.D.

2013 – 2014 Fulbright Fellow Department of Microbial Ecology, University of Vienna Supervised by David Berry, Ph.D. & Alexander Loy, Ph.D.

2012 – 2013 Undergraduate Research Student Biology Department, Juniata College Supervised by Regina Lamendella, Ph.D.

2011 Undergraduate Summer Research Fellow Dept. of Anesthesiology, Perioperative, and Pain Medicine, Brigham & Women's Hospital Supervised by Laura La Bonte, Ph.D. and Gregory L. Stahl, Ph.D.

Publications

1. McClure Carroll EE, Wang X, Shaw DK, O’Neal AJ, Oliva Chávez AS, Brown LJ, Boradia VM, Hammond HL, Pedra JHF. p47 licenses activation of the immune deficiency pathway in the tick Ixodes scapularis. Proc Natl Acad Sci USA. 2019; 116(1):205-210. doi: 10.1073/pnas.1808905116.

2. McClure EE, Oliva Chávez AS, Shaw DK, Carlyon JA, Ganta RR, Noh SM, Wood DO, Bavoil PM, Brayton KA, Martinez JJ, McBride JW, Valdivia RH, Munderloh UG, Pedra JH. Engineering of obligate intracellular bacteria: Progress, challenges, and paradigms. Nat Rev Microbiol. 2017; (9):544-558. doi: 10.1038/nrmicro.2017.59.

3. Shaw DK, Wang X, Brown LJ, Oliva Chávez AS, Reif KE, Smith AA, Scott AJ, McClure EE, Boradia VM, Hammond HL, Sundberg EJ, Snyder GA, Liu L, DePonte K, Villar M, Ueti MW, de la Fuente J, Ernst RK, Pal U, Fikrig E, Pedra JHF. Infection- derived lipids elicit an immune deficiency circuit in arthropods. Nat Commun. 2017; 8:14401. doi: 10.1038/ncomms14401.

4. Halfvarson J, Brislawn CJ, Lamendella R, Vazquez-Baeza Y, Walters WA, Bramer LM, D’Amato M, Bonfiglio F, McDonald D, Gonzalez A, McClure EE, Dunklebarger MJ, Knight R, Jansson JK. Dynamics of the human gut microbiome in inflammatory bowel disease. Nat Microbiol. 2017; 2:17004. doi: 10.1038/nmicrobiol.2017.4.

5. Shaw DK, McClure EE, Wang X, Pedra JHF. Deviant behavior: Tick-borne pathogens and inflammasome signaling. Vet. Sci. 2016, 3, 27.

6. Blanchette K, Shenoy A, Milner J, Gilley R, McClure E, Hinojosa C, Kumar N, Daughtery S, Tallon L, Ott S, King S, Ferriera D, Gordon S, Tettelin H, Orihuela C. Neuraminidase A exposed galactose promotes Streptococcus pneumoniae biofilm formation during colonization. Infect Immun. 2016; 84(10):2922-32.

7. Ulrich N, Rosenberger A, Brislawn C, Wright J, Kessler C, Toole D, Solomon C, Strutt S, McClure E, Lamendella R. Aquatic bacterial community is restructured by hydric dynamics associated with Superstorm Sandy. Appl Environ Microbiol. 2016; 82(12):3525-36.

8. Pavlov VI, Tan YS, McClure EE, La Bonte LR, Zou C, Gorsuch WB, Stahl GL. Human mannose-binding lectin inhibitor prevents myocardial injury and arterial thrombogenesis in a novel animal model. Am J Pathol. 2015; 185(2):347-55.

9. Trexler R, Solomon C, Brislawn CJ, Wright JR, Rosenberger A, McClure EE, Grube AM, Peterson MP, Keddache M, Mason OU, Hazen TC, Grant CJ, Lamendella R. Assessing impacts of unconventional natural gas extraction on microbial communities in headwater stream ecosystems in northwestern Pennsylvania. Front Microbiol. 2014; 5:522.

Book Chapters

1. Tick saliva and microbial effector molecules: two sides of the same coin. Shaw DK, McClure EE, Kotsyfakis M, Pedra JHF. Chapter 11 of Arthropod Vector: Controller of Disease Transmission (Volume 2): Vector Saliva-Host-Pathogen Interactions. Elsevier. 2017.

Grant Support

02/04/2018 – 02/03/2020 Principal Investigator: E. McClure (Sponsor: J. Pedra) NIH NIAID 1F31AI138440 “Characterization of an E3 Ubiquitin Ligase Substrate in the Tick Immune Deficiency Pathway”

Presentations

1. McClure EE, Wang X, & Pedra JHF. P47 restricts Anaplasma phagocytophilum acquisition by Ixodes scapularis ticks. American Society of Rickettsiology Annual Meeting, Milwaukee, WI. June 16-19, 2018. Oral presentation.

2. McClure EE & Pedra JHF. P47 restricts Borrelia burgdorferi colonization of Ixodes scapularis ticks. Biology of Spirochetes Gordon Research Seminar, Ventura Beach, CA. January 20, 2018. Oral presentation.

3. McClure EE & Pedra JHF. P47 restricts Borrelia burgdorferi colonization of Ixodes scapularis ticks. Biology of Spirochetes Gordon Research Conference, Ventura Beach, CA. January 21-26, 2018. Poster presentation.

4. McClure EE. The tick-pathogen interface: p47 restricts Borrelia burgdorferi colonization. Crosstalk. 2017 June 21. Oral presentation.

5. McClure EE, Nelson C, Felsheim R, Munderloh UG, Pedra JHF. High-Throughput Screening of Genes Associated with Host-Pathogen Interactions for an Obligate Intracellular Bacterium. Gordon Research Seminar and Conference in Tropical Infectious Diseases, Galveston, TX. March 11-17, 2017. Poster Presentation.

6. McClure E, Dunklebarger M, Li K, Halfvarsson J, Tysk C, McDonald D, Vázquez Baeza Y, Walters W, Knight R, Lamendella R, Jannson JK. Shifts in the Gut Microbiota of Inflammatory Bowel Disease Patients in a Longitudinal Study. American Society for Microbiology General Meeting, Denver, CO. May 18-21, 2013.

7. McClure E, La Bonte LR, Stahl GL. Mannose-Binding Lectin Concentrations Correlate with Enhanced Whole Blood Aggregation and Thrombin-Like Activity in Humans. Experimental Biology, San Diego, CA. April 21-25, 2012.

Awards, Honors, and Fellowships

2018 Office of Technology Transfer Graduate Translational Research Award 2018 Graduate Research Conference Outstanding Oral Presentation 2018 Ruth L. Kirschstein National Research Service Award F31 Predoctoral Fellowship 2016 T32 Infection & Immunity training grant recipient (09/01/16 – 02/03/18) 2013 Fulbright Full Research Fellowship, U.S. Department of State 2013 Distinction in Biochemistry, Juniata College 2011 German Academic Exchange (DAAD) Undergraduate Fellowship 2011 Undergraduate Summer Research Fellowship, American Physiological Society

Professional Service

 Volunteer, Baltimore Underground Science Space (BUGSS) (Sept. 2018 – present)  Co-developed rotation project and mentored rotation student (June – August 2018)  Co-organizer of 2018 Graduate Student Symposium (March 2018 – June 2018)  Co-organizer of Immunology Journal Club (July 2016 – May 2017)  Student representative for MMI program on GPILS 2016 Awards Committee  Moderated oral presentations at UM School of Medicine Summer Research Training Programs Student Research Forum (July 29, 2016 & July 28, 2017)  Member of GPILS Student Advisory Council (Sept. 2015 – May 2017)  Helped prepare students for qualifying exam twice per year (Feb. 2017 – present)

ABSTRACT

Title of Dissertation: XIAP-p47 Pairing Activates the Immune Deficiency Pathway in the Lyme

Disease Tick Ixodes scapularis

Erin E. McClure Carroll, Doctor of Philosophy, 2019

Dissertation Directed by: Joao Pedra, Ph.D., Associate Professor, Department of Microbiology

and Immunology

Globally, vector-borne diseases account for 17% of all infectious diseases. Most vectors are blood-feeding arthropods, which transmit bacterial, viral, and parasitic diseases to humans and animals. The tick Ixodes scapularis transmits seven pathogens, including Borrelia burgdorferi, the agent of Lyme disease. Lyme disease is the most important vector-borne disease in the United States and causes an estimated 329,000 infections annually. Best described in the model organism Drosophila melanogaster, the arthropod immune deficiency (IMD) pathway responds to microbial infection through activation of Relish, a nuclear factor (NF)-κB family transcription factor.

In I. scapularis ticks, the E3 ubiquitin ligase X-linked inhibitor of apoptosis (XIAP) regulates the IMD pathway through ubiquitylation. Yet, the tick genome notably lacks homologs to genes encoding key IMD pathway proteins as described in Drosophila. How XIAP activates the IMD pathway in response to microbial infection is poorly characterized and targets of XIAP- mediated ubiquitylation remain unknown. In this study, we identified the XIAP enzymatic substrate p47 as a positive regulator of the I. scapularis IMD network. XIAP polyubiquitylates p47 in a lysine (K)63-dependent manner and interacts with the ubiquitin-like (UBX) domain of p47. p47 also binds to Kenny (IKKγ/NF-κB essential modulator [NEMO]), the regulatory subunit of the inhibitor of NF-κB kinase (IKK) complex. Replacement of the amino acid lysine with arginine in the p47 linker region completely abrogated molecular interactions with Kenny.

Furthermore, reduction of p47 transcription levels through RNA interference in I. scapularis limited Kenny accumulation, reduced phosphorylation of IKKβ (IRD5), and impaired cleavage of the NF-κB molecule Relish. Accordingly, disruption of p47 expression increased microbial colonization of the tick-transmitted spirochete B. burgdorferi and the rickettsial agent Anaplasma phagocytophilum. In summary, we demonstrated that XIAP ubiquitylates p47 in a K63- dependent manner, culminating in Relish activation and antimicrobial responses. Manipulating immune signaling cascades in I. scapularis may lead to innovative approaches to reducing the burden of tick-borne diseases.

XIAP-p47 Pairing Activates the Immune Deficiency Pathway in the Lyme Disease Tick Ixodes scapularis

by Erin E. McClure Carroll

Dissertation submitted to the Faculty of the Graduate School of the University of Maryland, Baltimore in partial fulfillment of the requirements for the degree of Doctor of Philosophy 2019 © Copyright 2019 by Erin Elizabeth McClure Carroll All rights reserved

DEDICATION

To Jack, my best friend.

iii

ACKNOWLEDGEMENTS

This research was made possible by funding from the National Institutes of Health

(NIH) to Joao Pedra (P01AI138949, R01AI116523, subcontract recipient of

R01AI049424) and Erin McClure (F31AI138440). I thank Neal Silverman (University of

Massachusetts Medical School) for the Kenny antibody and the Drosophila S2* cell line,

Greg A. Snyder and Eric J. Sundberg (University of Maryland School of Medicine) for early technical assistance, Robert Bloch and Yinghua Zhang for surface plasmon resonance experimentation and expertise, David R. Goodlett and Young Ah Goo

(University of Maryland School of Pharmacy) for mass spectrometry technical services,

Jon Skare (Texas A&M Health Science Center) for the Borrelia burgdorferi B31 clone

MSK5 strain, and Ulrike G. Munderloh (University of Minnesota) for the ISE6 and

IDE12 Ixodes scapularis cell lines.

I thank my mentor, Joao Pedra, for his selfless dedication to my scientific training. He helped me cultivate writing, critical thinking, and strategic planning skills, and supported me personally and professionally. Every trainee should have a mentor like him. I also am grateful to Xiaowei (Eric) Wang, who tamed p47, helped me in all aspects of my project, and taught me everything I know about protein biochemistry. Dana Shaw mentored me during my early years in the Pedra Lab. She cheerfully passed on her laboratory know-how and served as a role model and a source of encouragement. I am also grateful to Adela Oliva Chávez for teaching me about Anaplasma, ticks, and tick cell culture. Holly Hammond taught me how to work with mice and enabled my research by managing the day-to-day lab operations. Thank you for making the Pedra Lab a wonderful place to work. I also am grateful to all of the current and former members of

iv the Pedra Lab with whom I’ve interacted over the years, including those mentioned above and Vishant Boradia, Sourabh Samaddar, Anya O’Neal, and Liron Marnin. Each of you has made a positive impact on my life, and I am thankful for you.

I am also grateful to my thesis committee: Eileen Barry, Nicholas Carbonetti,

Achsah Keegan, Ulrike Munderloh, and Joao Pedra. You provided support and guidance during my graduate training, and I’m grateful for your dedication to my success. Thank you especially to my thesis readers, Nicholas Carbonetti and Ulrike Munderloh, for taking the time to give me detailed feedback on this document. I thank Bret Hassel and

Heather Ezelle, as well as Nicholas Carbonetti and June Green, for the administrative support and scientific leadership of the MMI program. I am also grateful to the students of the MMI program for their support, positivity, and camaraderie.

Finally, I thank my friends and families for the love and encouragement you have generously given to me. Whether you are local or far-flung, human or canine, biological or in-law, I love you and am grateful for your unwavering support.

TABLE OF CONTENTS

CHAPTER 1: INTRODUCTION AND BACKGROUND ...... 1 1.1 Vector-borne infectious diseases ...... 1 1.2 Ticks as vectors ...... 1 1.3 Ixodes scapularis ...... 2 1.4 Pathogens transmitted by I. scapularis ...... 5 1.5 Arthropod immunity ...... 10 1.6 The immune deficiency pathway ...... 12 1.7 Open questions in the tick IMD pathway ...... 14 1.8 Project goals ...... 16

CHAPTER 2: MATERIALS AND METHODS ...... 20

CHAPTER 3: THE E3 UBIQUITIN LIGASE XIAP INTERACTS WITH P47 ...... 34 3.1 Introduction ...... 34 3.2 Results ...... 35 3.3 Discussion ...... 39

CHAPTER 4: P47 IS AN ENZYMATIC SUBSTRATE OF XIAP ...... 41 4.1 Introduction ...... 41 4.2 Results ...... 43 4.3 Discussion ...... 44

CHAPTER 5: P47 LICENSES ACTIVATION OF THE IMMUNE DEFICIENCY PATHWAY IN THE TICK I. SCAPULARIS ...... 46 5.1 Introduction ...... 46 5.2 Results ...... 47 5.3 Discussion ...... 54

CHAPTER 6: CONCLUSIONS AND FUTURE DIRECTIONS ...... 57 6.1 The role of the XIAP-p47 complex in IMD signaling in ticks ...... 60 6.2 Molecular details of p47-mediated IMD signaling ...... 61 6.3 Linking p47 to other functions in I. scapularis ...... 62 6.4 Conclusions ...... 63

REFERENCES ...... 65

LIST OF TABLES

Table 1: Pathogens transmitted by I. scapularis...... 6

Table 2: Antibodies used in this study...... 22

Table 3: Primers used in this study...... 27

Table 4: Putative interacting partners of I. scapularis XIAP...... 36

vii

LIST OF FIGURES

Figure 1: The life cycle of Ixodes scapularis ticks...... 3

Figure 2: Highlights of external and internal I. scapularis anatomy...... 4

Figure 3: Transmission of Borrelia burgdorferi by I. scapularis – a model for the transmission of tick-borne microbes...... 9

Figure 4: A comparison of immune deficiency (IMD) pathway architecture in Drosophila and I. scapularis...... 12

Figure 5: Graphical abstract of this project...... 17

Figure 6: The I. scapularis proteins XIAP and p47 interact...... 37

Figure 7: p47 interacts with XIAP through the ubiquitin-like (UBX) domain...... 39

Figure 8: Enzymatic reactions involved in ubiquitylation...... 41

Figure 9: The E3 ubiquitin ligase XIAP polyubiquitylates p47 in a K63-dependent manner...... 43

Figure 10: p47 knockdown impairs IMD pathway signaling...... 48

Figure 11: p47 interacts with Kenny, the regulatory subunit of the IκB kinase (IKK) complex ...... 50

Figure 12: p47 silencing increases A. phagocytophilum burden in tick cells...... 51

Figure 13: Silencing p47 increases microbial infection in vivo...... 53

Figure 14: Model of IMD pathway signaling in I. scapularis...... 59

viii

LIST OF ABBREVIATIONS

AMP antimicrobial peptide

Ap Anaplasma phagocytophilum

ATP adenosine triphosphate

BCA bicinchoninic acid

BEI Biodefense and Emerging Infections

BS3 bis(sulfosuccinimidyl)suberate

BSA bovine serum albumin

BSK-II Barbour-Stoenner-Kelly-II

CYLD Cylindromatosis

DAP-PGN diaminopimelic-type peptidoglycan

DMEM Dulbecco's modified Eagle media

DNR1 Defense repressor 1

ECL enhanced chemiluminescence

ELISA Enzyme-linked immunosorbance assay

Eyc Eyes closed

FADD Fas-associated death domain

FBS fetal bovine serum

FC fold change

FT flow-through

GNBP Gram-negative binding protein

GST Glutathione S-transferase

HA Hemagglutinin

HEK human embryonic kidney

HGA human granulocytic anaplasmosis

HRP Horseradish peroxidase

IACUC Institutional Animal Care and Use Committee

IAP2 Inhibitor of apoptosis protein 2

IBC Institutional Biosafety Committee

IKK Inhibitor of κB kinase

IMD immune deficiency

IN input

IP immunoprecipitation

IPTG isopropyl β-D-1-thiogalactopyranoside

IRD5 Immune response deficient 5

JAK-STAT Janus kinase-Signal transducer and activator of transcription

K lysine kDa kilodaltons

Key Kenny

Lys-PGN lysine-type peptidoglycan

MOI multiplicity of infection

MWCO molecular weight cut off

MyD88 Myeloid differentiation primary response 88

NEMO NF-κB essential modulator

NF-κB Nuclear factor-κB

NIH National Institutes of Health

N-Rel N-terminal portion of Relish

NSFL1 N-ethylmaleimide-sensitive factor L1

OLAW Office of Laboratory Animal Welfare

PBS phosphate buffered saline

PBS-T phosphate buffered saline pH 7.4 with 0.05% Tween-20

PCR polymerase chain reaction

PGN peptidoglycan

PGRP peptidoglycan recognition protein

PGRP-LE long peptidoglycan recognition protein E

PGRP-SA short peptidoglycan recognition protein A

PGRP-SD short peptidoglycan recognition protein D

POPG 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoglycerol

POSH Plenty of SH3s

POWV Powassan virus

PVDF Polyvinylidene difluoride qRT-PCR quantitative reverse transcription-PCR

RING Really Interesting New Gene

RIPA radioimmunoprecipitation assay

RNAi RNA interference

RPMI Roswell Park Memorial Institute

SCF complex Skp, Cullin, F-box containing complex scRNA scrambled control RNA

SDS-PAGE sodium dodecyl sulfate–polyacrylamide gel electrophoresis

SEM standard error of the mean

SEP Shp1, eyc, p47 domain

SG salivary glands

SHP1 Suppressor of high-copy phosphoprotein phosphatase 1 (PP1) siRNA silencing RNA

SLC46 Solute carrier family 46

SPR surface plasmon resonance

Spz Spätzle

TAB2 Transforming growth factor β activated kinase 1 binding protein 2

TAK1 Transforming growth factor β activated kinase 1

TBEV tick-borne encephalitis virus

TCT Tracheal cytotoxin

TMB 3,3’,5,5’-tetramethylbenzidine

TNF Tumor necrosis factor

UBA ubiquitin binding domain

Ube1 Ubiquitin activating enzyme E1

UBX ubiquitin-like domain

Uev1a Ubiquitin-conjugating enzyme E2 variant 1A

WT wild-type

XIAP X-linked inhibitor of apoptosis

xii

CHAPTER 1: INTRODUCTION AND BACKGROUND

1.1 Vector-borne infectious diseases

Infectious diseases remain a significant public health threat despite antibiotics and vaccines. According to data from the World Health Organization, 17% of all infectious diseases are vector-borne1. Blood-feeding arthropods such as mosquitoes, sandflies, and ticks transmit bacterial, viral, and parasitic infections to humans, leading to over 700,000 deaths annually as well as significant morbidity and economic costs1. Recent severe outbreaks of vector-borne infections have strained healthcare systems and highlighted the need for improved control of arthropod vectors of disease2.

1.2 Ticks as vectors

Ticks are globally distributed ectoparasites that transmit infectious diseases to humans and other animals3,4. They exclusively feed on blood and have been found on every continent except Antarctica5. As the top vector of veterinary infections, ticks cause tremendous economic losses and contribute to global food insecurity3,6. Ticks are also important vectors of human disease, especially in the Northern hemisphere4,6. Globally, the expansion of tick populations and distributions has been aided by climate change7.

Human encroachment on tick habitats8 coupled with larger tick populations and rising incidences of emerging and re-emerging infectious diseases9 mean that people are at increased risk for acquiring tick-transmitted infections4,6,10-12.

Ticks are arthropods and belong to the subphylum Chelicerata, which also contains spiders, scorpions, and mites13. Ticks can be divided into three main families:

Nuttalliellidae, Argasidae, and Ixodidae. The family Nuttalliellidae contains only a single

1 species, Nuttalliella namaqua14. All of the other tick species are categorized as either soft

(argasid) or hard (ixodid) ticks. Soft ticks tend to be larger and longer-lived, and exhibit a flexible cuticle that readily expands during blood-feeding15. Hard ticks have a tough cuticle and additional morphological and life cycle characteristics that distinguish them from soft ticks13. Both argasid and ixodid ticks transmit infectious diseases, but hard ticks contribute most to the global tick-borne disease burden16.

1.3 Ixodes scapularis

The most medically relevant arthropod vector of disease in the United States is the blacklegged tick Ixodes scapularis. I. scapularis belongs to the Ixodes ricinus complex, which consists of four species of evolutionarily related ticks with broad distributions17. I. ricinus is a widespread European tick, while Ixodes persulcatus inhabits central Asia18. In North America, Ixodes pacificus can be found on the West Coast of the

United States18. I. scapularis was primarily distributed in the Northeast and the Upper

Midwest; however, the tick’s range has expanded in recent years to include most states east of the Mississippi River18,19. This work will focus on the I. scapularis tick.

I. scapularis is a three-host tick that lives approximately two years (Figure 1)19-22.

These ticks feed once per life stage and seek progressively larger hosts as they mature.

The I. scapularis life cycle starts when larval ticks hatch from eggs in the spring to early summer. They have six legs and are barely visible to the human eye. During the late spring and summer, larvae parasitize hosts such as small rodents and birds. They molt into larger, eight-legged nymphs in the late summer. The newly emerged nymphs overwinter unfed until the spring of their second year of life. They then feed on small

2 mammals and birds during the late spring and summer and molt into sexually dimorphic,

3 mm long adults in the fall. Adult ticks mate either off host or on large mammals while the female takes a final bloodmeal either that fall or following winter dormancy. Males feed minimally, if at all, and die after mating. Female adults overwinter, deposit eggs in the spring to early summer, and die20.

Figure 1: The life cycle of Ixodes scapularis ticks. I. scapularis is a woodland-associated, three-host tick with approximately a two-year lifespan. A tick must take a single bloodmeal prior to molting and maturing into the next life stage. In the spring to early summer, eggs hatch into larvae. Larval ticks feed on small mammals and birds in the summer, and molt into nymphs in late summer to early fall. Unfed nymphs overwinter and emerge ready to feed the following spring before larvae are active. Nymphs are the most clinically relevant life stage of I. scapularis ticks as they transmit the infections acquired during as larvae to humans during their bloodmeal. Next, nymphs molt into adults in the fall. Adult ticks mate either off host or on large mammals (e.g. white-tailed deer), and the adult female must feed one final time to acquire sufficient nutrients to lay eggs. The female tick overwinters and lays eggs the following spring. Adult ticks of both sexes die following mating and oviposition. f, female; m, male. Figure adapted from the Centers for Disease Control and Prevention20.

The hallmark anatomical feature of hard ticks like I. scapularis is the scutum, a dark plate covering part or most of the dorsal surface of the tick (Figure 2)15,23. The mouthparts consist of the chelicerae and hypostome flanked by the palps24. The serrated chelicerae cut into host skin to generate the feeding lesion. The barbed hypostome anchors the tick and serves as a conduit by which blood is ingested and saliva egested24.

Like other hard ticks, I. scapularis sips slowly from the feeding lesion at first. During this initial engorgement phase, new cuticle is synthesized to accommodate the “big sip,” a colorful term which describes the final phase of tick feeding. During the big sip, the tick consumes a large volume of host blood prior to detaching5,13.

Figure 2: Highlights of external and internal I. scapularis anatomy. Externally, hard ticks can be distinguished from soft ticks by the presence of the scutum, a hard plate located on the tick dorsum. In I. scapularis, the size of the scutum depends on the life stage and sex of the tick. The tick mouthparts allow feeding, and consist of the hypostome and chelicerae, which are flanked by the palps. Internally, the hemocoel, or body cavity, of I. scapularis contains the major organs, which are bathed in a circulatory fluid known as hemolymph. Internal organs change in structure and characteristics depending on feeding status and life stage of the tick. The salivary glands and midgut are the most relevant organs in the context of tick-transmitted diseases15.

The most prominent internal anatomical features of I. scapularis are the salivary glands and midgut, which undergo morphological changes as the tick feeds and matures25. The salivary glands are responsible for synthesizing and secreting a variety of factors that enable efficient and long-term feeding26-29. Some of these factors include 4 immunomodulatory and anti-inflammatory molecules, anti-coagulants, analgesics, and cement proteins that protect the tick from host immunity and physical dislodgement.

Blood enters the midgut, the digestive organ of I. scapularis (Figure 2), while excess fluid is intermittently secreted into the host as saliva until the tick has fed to repletion and detaches. Salivary secretion during engorgement serves to concentrate the ingested blood two- to threefold. The bloodmeal remains in the midgut and is slowly digested intracellularly5,25.

1.4 Pathogens transmitted by I. scapularis

Seven pathogens, which include bacteria, viruses, and parasites, can be transmitted by I. scapularis (Table 1)19,30. Some ticks have narrowly defined host ranges; however, I. scapularis feeds promiscuously and does not require a human host to complete its life cycle22. Humans are incidental hosts for the I. scapularis-transmitted pathogens, which are maintained in reservoir populations such as small rodents19, meaning that humans are only accidentally infected19,30.

Table 1: Pathogens transmitted by I. scapularis.

The most common pathogen transmitted by I. scapularis is the Lyme disease spirochete Borrelia burgdorferi. In most cases, Lyme disease is characterized by erythema migrans (bullseye rash) and other symptoms such as fever and joint pain18,31.

While usually remedied with a course of antibiotics, about 10% of patients develop a poorly understood post-treatment syndrome characterized by persistent joint pain and neurological impairment32,33. About 30,000 cases of Lyme disease are reported annually in the United States, although the true number is likely 10 times higher, and these cases cost between $700 million and $1.3 billion in direct medical expenditures in 201534-36.

Borrelia mayonii also causes some cases of Lyme disease; however, it was only first recognized as a pathogen in 2016 and as such has not been well-studied37,38. Borrelia miyamotoi, a relative of relapsing fever spirochetes, causes a disease characterized by fever, headache and laboratory findings of leukopenia, thrombocytopenia and elevated liver transaminases, but not cyclic fevers39,40. B. miyamotoi can be cleared with antibiotics40.

In addition to the spirochetes, I. scapularis ticks transmit two other bacterial pathogens known as Anaplasma phagocytophilum and Ehrlichia muris subspecies eauclairensis. As obligate intracellular bacteria, these bacteria adopt a drastically different lifestyle than the Borrelia spirochetes. A. phagocytophilum preferentially infects neutrophils and causes human granulocytic anaplasmosis (HGA)41-44, while E. muris subsp. eauclairensis causes a mild ehrlichiosis in humans45-47. Anaplasmosis patients exhibit non-specific symptoms that often include fever, headache, chills, and occasionally liver damage. Anaplasmosis ranges in severity with older patients experiencing more severe symptoms. Doxycycline is the antibiotic of choice for treating HGA and

7 erhlichiosis41,45,46. Except for B. miyamotoi, bacterial pathogens transmitted by I. scapularis are only maintained transstadially in ticks and thus must be acquired by every new generation of ticks through feeding on reservoir hosts19.

Ticks in the I. ricinus complex transmit flaviviruses belonging to the tick-borne encephalitis (TBEV) serocomplex48. TBEV is endemic to Europe and parts of Asia, while the North American representative of the TBEV serocomplex is Powassan virus

(POWV). POWV is a rare, untreatable neurotropic viral infection against which no vaccines are available48. Initially, POWV infection presents as a fever followed by a wave of encephalitic symptoms that include paralysis and loss of coordination49. Long- term neurological side effects are evident in about half of POWV survivors48.

The seventh pathogen transmitted by I. scapularis is the apicomplexan parasite

Babesia microti, which, like other apicomplexans like Plasmodium spp., infects host erythrocytes19,50,51. Babesiosis ranges from asymptomatic illness to severe disease.

Patients with mild cases exhibit fever, flu-like illness, and anemia, while the signs of severe disease include disseminated intravascular coagulation and organ failure. Co- infection of B. microti with other tick-transmitted pathogens may lead to more severe disease50,52. A two-week regimen of anti-parasitic and anti-bacterial drugs is used to treat babesiosis50,53.

Most of our knowledge of tick-microbial interactions comes from studies of B. burgdorferi (Figure 3)54. I. scapularis larvae acquire B. burgdorferi by feeding on an infected host. The ingested spirochetes colonize the midgut and reside there for the entire life of the tick. The fed larva molts into a nymph, which is the most clinically relevant tick life stage, owing to its small size and ability to transmit pathogens18,31.

Figure 3: Transmission of Borrelia burgdorferi by I. scapularis – a model for the transmission of tick-borne microbes. I. scapularis ticks acquire B. burgdorferi by feeding on infected hosts as larvae or nymphs. Spirochetes colonize the tick midgut persistently at low levels throughout the life of the tick. (1) The incoming blood of a subsequent bloodmeal triggers the resident spirochetes to replicate rapidly. (2) Some spirochetes access the hemocoel and migrate to the salivary glands (SG), where they are (3) egested into the host along with tick saliva. Ticks must remain attached to the host for 24-48 hours for efficient spirochete transmission to occur54.

The nymph feeds on a host, triggering exponential replication of the resident spirochetes in the midgut lumen in response to the influx of blood55. A few spirochetes manage to breach the midgut epithelium and disseminate to the tick’s salivary glands54,56,57. From there, the B. burgdorferi bacteria are egested into the host with tick saliva during feeding56. I. scapularis ticks must feed for 24-48 hours for efficient bacterial transmission to occur55,58. Multiday feeding is also necessary for the transmission of B. microti58, while POWV can be passed to the host within minutes to hours of feeding59.

1.5 Arthropod immunity

Arthropods have complex immune systems that protect them from microbial infection60. Immunity is particularly relevant to vector-borne diseases, since the arthropod immune system influences vector competence, i.e., the ability to acquire, maintain, and transmit pathogens61. Arthropods engage innate cellular and humoral immunity programs to modulate microbial loads and to protect themselves from detrimental microbial invasion but lack classical adaptive immunity61,62. Our understanding of arthropod immune signaling pathways has been informed almost solely by the Drosophila melanogaster (hereafter referred to as Drosophila) model60.

Arthropods have an open circulatory system that contains hemolymph, a blood- like organ containing hemocytes63. Plasmatocytes are the most abundant type of hemocyte64,65. These professional phagocytes engulf pathogens and clear debris in a manner similar to macrophages65. Crystal cells are another class of hemocyte tasked with producing prophenoloxidase, which is cleaved to generate active phenoloxidase during

10 the melanization immune response. Collectively, hemocytes sequester and kill invading microbes through cellular immune responses66.

The hallmark of arthropod humoral immune activation is the production of antimicrobial peptides (AMPs) by plasmatocytes, enterocytes, and cells in the fat body61,64,66. Microbial ligands induce AMP production through two humoral immune signaling networks. In the Toll pathway, some members of the poorly named Gram- negative binding protein family (GNBPs) and peptidoglycan (PGN) recognition proteins

(PGRPs) detect Gram-positive bacteria and fungi67-69. GNBP1 binds to lysine-type PGN

(Lys-PGN) and forms a complex with PGRP-short A (PGRP-SA). Activated GNBP1 processes the PGN, allowing PGRP-SA-mediated recognition70,71. Another short PGRP transcript, PGRP-SD, also recognizes Lys-PGN, but without a cofactor72. Fungal β- glucans are detected by GNBP373. Following recognition of the microbial ligands, a series of protease cascades leads to the cleavage of the cytokine Spätzle (Spz). Cleaved

Spz dimerizes and activates the Toll receptor. Toll signaling culminates in MyD88- mediated nuclear factor (NF)-κB activation and subsequent AMP (e.g., drosomycin) production67.

The second arthropod humoral immune signaling pathway is the immune deficiency (IMD) pathway, which responds to Gram-negative bacteria and is functionally analogous to the tumor necrosis factor (TNF) receptor pathway in mammals60,74,75. The

Janus-activated kinase (JAK)-Signal transducer and activator of proteins (STAT) pathway participates in antiviral responses and crosstalks with the Toll and IMD networks; however, the JAK-STAT pathway technically does not constitute humoral immunity61.

1.6 The immune deficiency pathway

In Drosophila, the IMD pathway is triggered by diaminopimelic (DAP)-type PGN found primarily in Gram-negative and some Gram-positive bacteria75 (Figure 4).

Figure 4: A comparison of immune deficiency (IMD) pathway architecture in Drosophila and I. scapularis. The IMD signaling network in I. scapularis ticks contains the same core signaling relay as Drosophila and culminates in the phosphorylation and cleavage of the nuclear factor (NF)-κB molecule Relish. Similar to Drosophila, the IMD pathway is regulated by lysine (K)63-polyubiquitylation mediated by X-linked inhibitor of apoptosis (XIAP), an E3 ubiquitin ligase analogous to Drosophila Inhibitor of apoptosis 2 (IAP2). However, upstream signaling components, such as the receptor and central adaptor molecules, as well as the targets of Relish are unknown in the tick system. PGN, peptidoglycan; PGRP, Peptidoglycan recognition protein; DAP, diaminopimelic; FADD, Fas-associated death domain; Uev1a; Ubiquitin-conjugating enzyme E2 variant 1A; TAK1, Transforming growth factor β-activated kinase 1; TAB2, TAK1-binding protein 2; Key, Kenny; POSH, Plenty of SH3s; CYLD, Cylindromatosis; DNR1, Defense repressor 1; SCF complex, Skp, Cullin, F-box containing complex; N-Rel; N-terminal Relish; AMPs, antimicrobial peptides.

As in the Toll pathway, PGRPs detect PGN76. Transmembrane PGRP-LC binds

DAP-PGN in concert with the two secreted co-receptors PGRP-SD77 and truncated

PGRP-long E (PGRP-LE)75. Full-length cytosolic PGRP-LE surveys the intracellular space and detects cytoplasmic monomeric DAP-PGN (also known as tracheal cytotoxin

[TCT])75, which is transported into the cytosol by solute carrier family (SLC) 46 transporters78. Other PGRPs dampen IMD responses by degrading free DAP-PGN or by antagonizing PGRP-LC79.

The binding of DAP-PGN to PGRPs recruits a protein complex consisting of the death domain-containing adaptor proteins Imd80 and Fas-associated death domain protein

(FADD)81, as well as the caspase-8 homolog Dredd in Drosophila82,83. Dredd cleaves

Imd, exposing a polyubiquitylation site, allowing the E3 ubiquitin ligase Inhibitor of apoptosis 2 (IAP2) to ubiquitylate Imd in a lysine (K)63-dependent manner84,85. The resulting ubiquitin scaffold functions as a docking platform for a downstream kinase cascade86-89, which culminates in the phosphorylation and cleavage of the NF-κB molecule Relish90,91. The N-terminal Relish fragment translocates to the nucleus and activates genes encoding proteins involved in the immune response, such as AMPs79,91-93.

For decades the insect model organism Drosophila has been used to study arthropod-pathogen interactions and immunity61. However, increasing evidence suggests that Drosophila may not faithfully represent non-insect arthropods, such as ticks, whose ancestral lineage diverged from insects and crustaceans over 500 million years ago61,62,94-

96. For example, the IMD pathway exists as two distinct immune circuits in Drosophila and I. scapularis97. While both pathways are triggered by microbial ligands and result ultimately in the cleavage of the NF-κB molecule Relish to initiate antimicrobial

13 transcriptional programs, key differences exist between the upstream signaling cascades

(Figure 4).

Homologs to the key Drosophila IMD pathway genes, such as imd, fadd, and the transmembrane PGRP ird7 have not been identified in the tick genome62,94-96,98.

However, we discovered that I. scapularis employs an alternative IMD signaling network to limit B. burgdorferi and A. phagocytophilum acquisition97. While the tick IMD pathway retains most of the core IMD pathway components found in insects, some key differences remain. We showed that microbial lipids (e.g., 1-palmitoyl-2-oleyl-sn- glycerol-3-phosphoglycerol (POPG)) activate the IMD pathway, although the receptor to which they bind is currently unknown97. In the Drosophila IMD pathway, IAP2 positively regulates signal transduction through K63-ubiquitylation of the adaptor Imd. In

I. scapularis, the Really Interesting New Gene (RING)-type ubiquitin ligase XIAP is responsible for positively regulating the IMD pathway (Figure 4)97,99. Together with the

Ubiquitin activating enzyme E1 (Ube1) and the heterodimeric E2 conjugating enzyme complex Ubiquitin-conjugating enzyme E2 variant 1A (Uev1a) and Bendless, XIAP performs K63-linked ubiquitylation and restricts B. burgdorferi and A. phagocytophilum infection97,99.

1.7 Open questions in the tick IMD pathway

The absence of genes encoding key classical IMD pathway components suggests several open questions. For instance, the targets of I. scapularis Relish are unknown, as homologs to AMPs specifically regulated by Drosophila Relish are absent in I. scapularis98, and attempts to find Relish-regulated genes in ticks have proven fruitless.

The receptor of the tick IMD pathway is also unknown. Despite lacking transmembrane

PGRPs, the I. scapularis IMD signaling network responds to DAP-PGN and two lipid agonists97. The diversity of agonists suggests that there may be several receptors or co- receptors that lead to IMD pathway-regulated Relish cleavage. The molecular mechanism linking XIAP-mediated polyubiquitylation and Relish activation is also poorly understood. In the Drosophila system, IAP2 polyubiquitylates the adaptor molecule Imd

(Figure 4)80,85. However; in I. scapularis, no homolog to imd or substrates of XIAP have been identified. Whether ubiquitylated substrates of XIAP impact anti-pathogen defenses in I. scapularis also remains elusive.

To find candidate substrates of XIAP, A. phagocytophilum-infected I. scapularis cell line (ISE6) lysates were incubated with recombinant I. scapularis XIAP tagged with glutathione-S transferase (GST). We then co-immunoprecipitated XIAP-GST and identified the protein p47 as a potential binding molecule through mass spectrometry. p47 is a highly conserved protein found in diverse eukaryotes ranging from Saccharomyces cerevisiae (Suppressor of high-copy PP1 protein [Shp1])100-102, Drosophila (Eyes closed

[Eyc])103, and to mammals (N-ethylmaleimide-sensitive factor [NSF]-like 1 [NSFL1] cofactor p47)104. It contains two ubiquitin-associated domains: an N-terminal ubiquitin binding domain (UBA)105 and a C-terminal ubiquitin-like domain (UBX)105,106. First described as a cofactor of the ATPase p97, p47 binds p97 and targets this molecule to various subcellular locations100,102,104,107,108. The molecular scaffold composed of p47 and p97 is involved in cell cycle regulation and organelle membrane biogenesis100,102,104,107,108. In yeast, the Shp1 (p47)/Cdc48 (p97) complex has also been implicated in membrane formation in autophagy. Shp1 interacted directly with ubiquitin-

15 like Atg8 to promote autophagosome biogenesis through an unknown mechanism109. p47 also acts independently of p97 in NF-κB regulation. In humans, p47 blunted NF-κB signaling in vitro by promoting the degradation of the polyubiquitylated NF-κB essential modulator (NEMO, also known as IKKγ)110.

1.8 Project goals

Given the evolutionary conservation of p47 and the link to regulation of NF-κB signaling pathways, we investigated the role of p47 in tick IMD antimicrobial responses.

We hypothesized that XIAP binds and polyubiquitylates p47 in a K63-dependent manner during IMD pathway activation in I. scapularis ticks. I investigated this central hypothesis in three specific aims (Figure 5).

Figure 5: Graphical abstract of this project. The protein p47 was identified as a putative substrate of XIAP in I. scapularis. Aims 1-3 are designed to investigate the central hypothesis that XIAP binds and polyubiquitylates p47 in a K63-dependent manner during IMD pathway activation in ticks.

Aim 1: Examine the molecular and biophysical interactions between XIAP and p47. In this aim, I used standard protein biochemistry techniques as well as surface plasmon resonance (SPR) to validate the interaction between XIAP and p47, measure the binding affinity between the two molecules, and determine which domain of p47 was responsible for binding to XIAP. We demonstrated that p47 and XIAP interact with nanomolar affinity, and that the UBX domain of p47 is required for binding to XIAP.

Aim 2: Examine the biochemical interactions between XIAP and p47. The goal of Aim 2 was to determine if p47 was an enzymatic substrate of the E3 ubiquitin ligase XIAP. I used cell-free ubiquitylation assays to demonstrate that XIAP polyubiquitylates p47 in a K63-dependent manner.

Aim 3: Determine if p47 is member of the IMD pathway in I. scapularis.

While Aims 1 and 2 focused on the interactions between XIAP and p47, the goal of Aim

3 was to define the functional role of p47 in I. scapularis immunity. I demonstrated impaired IMD pathway signaling as well as enhanced microbial colonization upon p47 silencing in I. scapularis cell lines and nymphs. I also showed that p47 binds to Kenny, another member the IMD signaling network.

Our findings answered two open questions in the tick humoral immune signaling field. First, we demonstrated that p47 is a substrate of XIAP, that is, p47 is polyubiquitylated by XIAP in a K63-dependent manner. Second, we also showed that p47, as a substrate of XIAP, positively regulates IMD pathway-mediated antimicrobial responses in I. scapularis. Collectively, these findings enhance our knowledge of immune signaling in ticks, which are critically understudied when compared to other medically relevant hematophagous arthropods (e.g. mosquitoes). Furthermore, our data highlight

18 the diversity of arthropod immune systems, and remind us that the simple transposition of immune signaling pathways from model organisms to other arthropods may lead to poorly characterized immune signaling paradigms and erroneous conclusions. In the future, our enhanced knowledge of the I. scapularis IMD signaling network may be leveraged to design novel strategies to reduce the global burden of vector-borne diseases.

CHAPTER 2: MATERIALS AND METHODS1

The experimental procedures, antibodies, and primers in this chapter have been previously published111.

Bacteria. E. coli BL21 (DE3) cultures were grown overnight at 37°C in lysogeny broth supplemented with 100 μg/ml ampicillin. Anaplasma phagocytophilum HZ was cultured in the human HL-60 cell line (ATCC, CCL-240)112. Host-free A. phagocytophilum was obtained by mechanical lysis using a syringe. Briefly, infected HL-

60 cells were collected at 2300 x g for 10 minutes at 4°C. The pellet was resuspended in cold phosphate buffered saline (PBS) pH 7.4 and lysed by passing four times through a

25-gauge needle. Cellular debris was collected by centrifuging at 750 x g for 5 minutes at

4°C. The supernatant was kept and subjected to syringe lysis as above. Host-free A. phagocytophilum were collected by centrifuging at 2300 x g for 10 minutes at 4°C. Host- free bacteria were enumerated by this formula: number of infected HL-60s x 5 morulae/cell x 19 bacteria/morula x 0.5 (50% recovery)113. Low passage B. burgdorferi

B31 clone MSK5 was grown in Barbour-Stoenner-Kelly (BSK)-II medium supplemented with 6% normal rabbit serum at 37°C, as described elsewhere114,115. Profiling to ensure the presence of necessary virulence plasmids was performed, as described elsewhere114.

Enzyme-linked immunosorbent assay (ELISA). A high-binding 96-well

ELISA plate (Costar) was coated with either no protein, 11.4 pmol bovine serum albumin

(BSA), 11.4 pmol p47-His, 50 ng recombinant wild-type (WT), Δ UBA, or Δ UBX p47-

His (for recombinant protein binding experiments), or 200 ng pooled flat tick lysate (for in vivo binding experiments) in 500 mM carbonate-bicarbonate buffer (pH 9.5) at 4°C

1 McClure Carroll EE, et al. (2019). p47 licenses activation of the immune deficiency pathway in the tick Ixodes scapularis. Proc Natl Acad Sci U S A 116(1):205-210. 20 overnight. Plates were washed with PBS pH 7.4 with 0.05% Tween-20 (PBS-T) three times and blocked with 10% heat-inactivated fetal bovine serum (FBS) for 1 hour at room temperature, after which they were incubated with the indicated concentrations of GST

(Thermo Scientific, 20237), XIAP-GST, or Kenny-GST for two hours at room temperature. Plates were washed five times with PBS-T and subsequently incubated with

α-GST (Table 2) followed by an additional 5 washes and incubation with α-mouse IgG conjugated to horseradish peroxidase (HRP) for one hour at room temperature (Abcam,

1:10,000). Plates were washed seven times in PBS-T and developed with 3,3’,5,5’- tetramethylbenzidine (TMB) (BD Biosciences) for 10-30 minutes. The reaction was stopped with 1 M H2SO4 and the absorbance was read at 450 nm with a 595 nm correction (Bio-Rad iMark). Background signal (BSA or no protein) was subtracted from the p47-His-coated well signal prior to graphing and statistical analysis.

Table 2: Antibodies used in this study.

*No longer commercially available, **From Rhipicephalus microplus

Eukaryotic cell culture. All cell cultures were verified by polymerase chain reaction (PCR) to be Mycoplasma-free (Southern Biotech, 13100-01). The human cell line HL-60 (ATCC, CCL-240) was cultured in Roswell Park Memorial Institute (RPMI) medium supplemented with 10% heat-inactivated FBS (Gemini Bio-Products) and 1X

GlutaMAX (Gibco). HL-60s were maintained in suspension cultures at densities ranging from 1 x 105 to 1 x 106 cells/ml. Human embryonic kidney (HEK) 293T cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with

GlutaMAX, 10% heat-inactivated FBS, and 1% penicillin and streptomycin antibiotics

(Corning). Mammalian cell cultures were maintained at 37°C and 5% CO2. The I. scapularis ISE6 and IDE12 cell lines were cultured in L15C300 medium supplemented with 10% heat-inactivated FBS (Sigma), 10% tryptose phosphate broth (BD, 260300), and 0.1% bovine lipoprotein concentrate (MP Biomedicals,191476) at 34°C. ISE6 and

IDE12 cells were grown to confluence and either seeded at 3 x 106 cells/well in 6-well plates (IDE12) (Sigma) or 1 x 106 cells/well in 24-well plates (ISE6) (Celltreat), or subcultured at 1:1-1:20 in capped T25 flasks (Cellstar)116. The S2* cell line was cultured in Schneider’s Drosophila medium supplemented with 10% heat-inactivated FBS and 1% penicillin and streptomycin antibiotics117. The loosely adherent culture was maintained at a density between 1 x 106 and 1 x 107 cells/ml at 28°C.

HEK 293T transfection and co-immunoprecipitation. HEK 293T cells were transfected, as described elsewhere97. Briefly, 4 μg pCMV-xiap- hemagglutinin (HA), pCMV-kenny-HA, pCMV-p47-FLAG, or pCMV-*p47-FLAG plasmids were transfected into 1 x 106 HEK 293T cells with Lipofectamine 2000 (Invitrogen) using Opti-MEM I

Reduced Serum Medium (Invitrogen). After five hours, the transfection reagent was

23 removed, and cells were left in complete medium for 48 hours. Cells were collected, washed once with PBS, and lysed in immunoprecipitation lysis buffer (Thermo

Scientific). The cell lysate (250 μg to 1 mg) was incubated with 30-60 μl α-FLAG-M2

(Sigma, A2220) or α-HA (Pierce, 26181) cross-linked agarose beads overnight at 4°C.

Beads were washed three times (XIAP/p47 and Kenny/p47 co-immunoprecipitations) or six times (head-to-head comparison of Kenny/p47 and Kenny/*p47 co- immunoprecipitations) with 50 mM Tris, 150 mM NaCl, pH 7.5 and boiled for 5 minutes in 2X Laemmli buffer to elute proteins. Boiled beads and input samples were analyzed by western blot for XIAP-HA, Kenny-HA, or p47-FLAG expression.

Mouse infections. All mouse experiments were approved by the Institutional

Biosafety (IBC, 00002247) and Animal Care and Use (IACUC, 0216015) committees at the University of Maryland School of Medicine and complied with National Institutes of

Health (NIH) guidelines (Office of Laboratory Animal Welfare [OLAW] assurance numbers A3200-01, A323-01, A3270-1). Female eight- to 10-week old mice were used for tick experiments. C57BL/6 mice were infected intraperitoneally with 1 x 107 host-free

A. phagocytophilum HZ. C3H/HeJ mice (Jackson Laboratories, 000659) were infected intradermally with 1 x 105 low-passage B. burgdorferi B31 MSK5.

p47 identification by mass spectrometry. The XIAP-GST co- immunoprecipitation was performed by crosslinking recombinant XIAP-GST to glutathione agarose beads with bis(sulfosuccinimidyl)suberate (BS3, ThermoFisher,

21580). ISE6 cells were infected with A. phagocytophilum at a multiplicity of infection

(MOI) of 50 for 30 minutes and then sonicated in 20 mM Tris pH 8.9, 150 mM NaCl,

0.01% Triton X-100 with protease inhibitors. The lysate was incubated with crosslinked

XIAP-GST for 1 hour at 4°C. Beads were washed four times and eluted in 20 mM Tris pH 8.9, 150 mM NaCl, 10 mM DTT, 5 mM EDTA, 0.01% Triton X-100 with

PreScission Protease (GE Healthcare, 27-0843-01). Eluted proteins were precipitated, trypsinized overnight, desalted, and analyzed by the University of Maryland School of

Pharmacy Mass Spectrometry Facility. Identified proteins were filtered to remove contaminants (e.g. keratin), XIAP, GST, and other associated proteins, as well as proteins involved in metabolism, DNA replication, RNA transcription, and protein synthesis.

Plasmids. The plasmids pCMV-HA (Christopher A. Walsh, plasmid #32530), pCMV-FLAG (Joan Massague, plasmid #15738), pMT/V5/His (Cyril Gueydan, plasmid

#63752), and pCoPURO (Francis Castellino, plasmid #17533) were obtained from

Addgene and were received as gifts from the depositing scientists. All constructs were amplified from tick or IDE12 cDNA using high-fidelity polymerase. Bendless and xiap were cloned into pGEX-6P-2 as previously reported97,99. Xiap was cloned into pCMV-

HA as previously reported for expression in HEK 293T97. Kenny cDNA cloned into pCMV-HA using the EcoRI and NotI restriction enzyme sites for expression in HEK

293T cells. GenScript performed codon optimization of tick kenny for expression in E. coli. The codon-optimized insert was digested with BamHI and EcoRI and subcloned into the pGEX-6P-2 vector.

GenScript performed codon optimization of p47 cDNA for expression in

Drosophila S2* cells. Codon-optimized p47 was cloned into the pCMV-FLAG vector using the EcoRI and SalI. For expression in S2* cells, WT and mutant p47 genes were cloned into the pMT/V5/His vector using the EcoRI and AgeI sites. Domain deletion p47 mutant constructs were generated from the full-length template using the primers

25 indicated in Table 3 and cloned into the pCMV-FLAG and pMT/V5/His vectors. PCMV-

*p47-FLAG was generated by mutating codons encoding lysine to arginine residues at positions 109, 139, 141, 154, and 300 (New England Biolabs, E0554). All plasmids were verified by sequencing.

Protein extraction and subcellular fractionation. Cell pellets or pooled flat ticks were lysed with radioimmunoprecipitation (RIPA) buffer (Thermo Scientific,

89900) buffer with protease inhibitors (Roche, 11836170001), and incubated at 4°C for

20 minutes prior to centrifugation at 12,000 x g for 15 minutes at 4°C. The supernatant was kept at -80°C until western blotting. Cytoplasmic and nuclear protein extracts were prepared from 3 x 106 IDE12 cells using the NE-PER Nuclear and Cytoplasmic

Extraction Kit (Thermo Scientific, 78833) and stored at -80°C. Concentrated protein extracts were obtained by following the kit protocol for 10 μl packed cell volume.

Quantitative reverse transcription PCR (qRT-PCR). The PureLink RNA Mini kit (Ambion, 12183025) was used to extract RNA from cells or engorged ticks preserved in Trizol. 500 ng RNA was used to synthesize cDNA with the Verso cDNA Synthesis Kit

(ThermoFisher, AB-1453). P47 silencing and bacterial burdens were quantified by qRT-

PCR using the primers listed in Table 3.

Table 3: Primers used in this study.

Table 3 Continued

*These sequences were codon- optimized for expression in Escherichia coli, HEK 293T, and Drosophila melanogaster hosts. Kenny-HA was not codon- optimized; however, the kenny sequence subcloned into the pGEX-6p-2 vector was codon- optimized for E. coli expression.

** Primers were designed based on the relish sequence derived from an I. scapularis transcriptome (NCBI project number: PRJNA230499).

***These primers were used to generate the *pCMV-p47-FLAG construct, where codons encoding lysines 109, 139, 141, 154, and 300 were mutated to arginine.

Recombinant proteins. Recombinant XIAP and Bendless were produced as described previously97. E. coli BL21 (DE3) was transformed with the pGEX-6P-2-xiap, pGEX-6P-2-bendless, or pGEX-6P-2-kenny plasmids and induced with 0.1 mM isopropyl

β-D-1-thiogalactopyranoside (IPTG) for 18 hours at 18-22°C. Cells were collected by centrifugation, resuspended in 20 mM Tris pH 8.9 (XIAP) or pH 8.1 (Bendless and

Kenny), 300 mM NaCl, 5% glycerol, and lysed three times with the Microfluidics LV2 homogenizer. Lysates were clarified by centrifugation, and the resulting supernatant was incubated with glutathione agarose beads (Thermo Scientific, 16100) for 1 hour at room temperature. Beads were washed three times (50 mM Tris pH 8.0 and 150 mM NaCl) prior to elution with 50 mM Tris pH 8.0, 150 mM NaCl, and 10 mM reduced L- glutathione. Eluates were concentrated on 3 kDa (Bendless-GST), 30 kDa (XIAP-GST), or 50 kDa molecular weight cut off (MWCO) (Kenny-GST) spin filter concentrators

(Amicon EMD Millipore). Concentrated eluates were dialyzed against 1X PBS on 7 kDa

MWCO desalting columns (Thermo Scientific, 89889).

Recombinant full-length and deletion mutant p47-His proteins were produced by inducing stably transfected pMT-p47-His S2* cells with 500 μM CuSO4 for 48 hours.

Cells were collected by centrifugation, resuspended in RIPA buffer with protease inhibitors and incubated in ice for 30 minutes with periodic brief agitation. The lysate was clarified by centrifugation at 3200 x g, 4°C, 30 minutes, and the supernatant was incubated with prepared Hi60 Ni Superflow resin (Clontech, 635659) for 1 hour at room temperature or overnight at 4°C. The resin was washed once with binding buffer (50 mM

Tris pH 8.0, 100 mM NaCl, 5 mM imidazole, 0.1 mM EDTA) followed by three buffer washes (50 mM Tris pH 8.0, 300 mM NaCl, 10 mM imidazole, 0.1 mM EDTA).

Recombinant p47-His was eluted by incubating the resin for 5 minutes in 50 mM Tris pH

8.0, 50 mM NaCl, 300 mM imidazole, 0.1 mM EDTA with protease inhibitors. All eluates were concentrated and dialyzed against 1X PBS.

RNA interference (RNAi) silencing and tick experiments. P47 silencing RNA

(siRNA) and the scrambled control RNA (scRNA) were synthesized with the Silencer siRNA construction kit (Ambion, AM1620) using the primers listed in this section. To silence p47, 3 μg or 10 μg siRNA or scRNA was transfected into 1 x 106 ISE6 or 3 x 106

IDE12 cells using Lipofectamine 2000, respectively. After 24 hours, the transfection reagents were removed, and cells were infected with A. phagocytophilum at an MOI of 50

(ISE6) or 150 (IDE12) for 18 hours (ISE6), or times indicated (IDE12). IDE12 cells were harvested, washed twice with PBS, and stored at -80°C. ISE6 cells were harvested in

Trizol (Ambion, 15596018) and stored at -80°C. I. scapularis ticks were microinjected with 10 ng siRNA or scRNA, as described elsewhere97. After resting overnight, ticks were placed on infected, anesthetized mice seven (A. phagocytophilum) or 10 (B. burgdorferi) days post-infection and allowed to attach for 30 minutes. The anesthetic was subsequently removed, and ticks were allowed to feed until repletion. Fully engorged ticks were collected, weighed, and frozen in single tubes at -80°C until analysis. Ticks were snap frozen in liquid nitrogen and homogenized prior to RNA or protein extraction.

S2* transfection. 3 x 106 S2* cells were seeded in 6-well plates and grown overnight at 28°C. CaCl2 (240 mM final concentration) was mixed with 19 μg plasmid

DNA. Sterile H2O was added to bring the solution volume to 300 μl. The DNA/CaCl2 solution was added dropwise to HEPES-buffered saline (50 mM HEPES, 1.5 mM

Na2HPO4, 280 mM NaCl, pH 7.1). The resulting solution was incubated at room

31 temperature for 30 minutes, vortexed briefly, and added dropwise to the S2* cells. After incubating for 24h, cells were washed twice followed by the addition of fresh complete medium. Stably transfected S2* cell lines were generated by co-transfecting the pMT-His plasmid of interest and 1 μg pCoPURO. After 24 hours, cells were washed twice and grown in fresh complete medium for 48 hours. Successfully transfected cells were selected with 5 μg/ml puromycin.

SPR. The Biacore 2000 instrument (GE Healthcare) was used to perform SPR.

XIAP-GST or GST alone was captured to a Biacore CM5 chip (GE Healthcare) coated in

α-GST antibodies. To determine the association and dissociation constants (KD = Kd/Ka) of the protein-protein interaction, increasing concentrations of p47-His diluted in HBS-

EP buffer (10 mM HEPES pH 7.4, 150 mM NaCl, 3 mM EDTA, 0.005% v/v Surfactant

P20) were flowed over the chip. Non-specific binding was corrected for by subtracting the GST: p47-His signal from the XIAP-GST: p47-His signal. Data were fitted with a 1:1 binding mode with mass transfer using Biacore T200 Evaluation software to determine the kinetic rate constants and the KD for the XIAP: p47 interactions.

Ticks. I. scapularis nymphs were supplied by the NIH Biodefense and Emerging

Infectious Diseases (BEI) Research Resources Repository. Ticks were housed upon arrival at 23°C with 85% relative humidity and a 12/10-hour light/dark photoperiod regimen.

Western blotting. The full western blotting procedure was described elsewhere97.

Briefly, protein lysates were quantified by bicinchoninic acid (BCA) assay (Thermo

Scientific, 23227). Equal amounts of protein were boiled in 6X Laemmli sample buffer

(VWR) containing 5% β-mercaptoethanol. Proteins were separated by sodium dodecyl

32 sulfate polyacrylamide gel electrophoresis (SDS-PAGE) (Biorad mini-PROTEAN gels) and transferred onto polyvinylidene fluoride (PVDF) membranes (Biorad). Membranes were blocked for 1 hour with 5% skim milk (Biorad) in PBS-T, followed by overnight primary antibody incubation at 4°C in 0.25% skim milk PBS-T at the dilutions indicated in Table 2. Blots were washed three times in PBS-T and incubated with secondary antibodies for 1 hour at room temperature. Blots were next washed three times in PBS-T before 1-2 minutes incubation in fresh enhanced chemiluminescence (ECL) substrate solution prior to visualization (Thermo Scientific, 32106). If needed, blots were incubated in stripping buffer (Thermo Scientific, 21059) for 15 minutes at room temperature, and then washed three times in PBS-T prior to blocking and re-probing with the primary antibody.

Ubiquitylation assays. The in vitro ubiquitylation reactions contained reaction buffer (500 mM Tris pH 7.4, 10 mM DTT), 1.5 mM Mg-ATP (Boston Biochemical, B-

20), 155 nM Ube1 (Boston Biochemical, E-305), 250 nM Uev1a (Boston Biochemical,

E2-662), 375 nM Bendless-GST, 2.4 mM XIAP-GST, 10 mM p47-His, and 50 mM ubiquitin (Boston Biochemical, U-100H) and were incubated at 37°C for 1 hour prior to stop buffer addition (Boston Biochemical, SK-10). Reactions were dialyzed against PBS.

P47-His was isolated from the reactions using the protocol for p47-His purification from

S2* cells. The eluate and flow-through were dialyzed against PBS to remove imidazole and subjected to western blotting to assess K63- and K48-specific ubiquitylation (Table

2).

CHAPTER 3: THE E3 UBIQUITIN LIGASE XIAP INTERACTS

WITH P471

3.1 Introduction

The spirochete B. burgdorferi, which causes an estimated 329,000 cases of Lyme disease each year36, is a zoonotic disease agent transmitted from woodland rodents to humans through the bite of infected I. scapularis ticks31. I. scapularis is a competent vector for six additional bacterial, viral, and protozoan pathogens, many of which cause emerging infectious diseases (Table 1)3-6,19,30. Despite the increasing public health relevance of I. scapularis, relatively little is known about the tick-microbe interface in comparison to other arthropod-borne infectious diseases.

The tick immune system impacts the acquisition, maintenance, and transmission of tick-borne pathogens61,118. Arthropods lack adaptive immunity, and instead rely exclusively on cellular and humoral innate responses60-62. Drosophila has served as a valuable model for elucidating the principles of arthropod humoral immunity; however, flies and ticks have evolved under different evolutionary pressures62,94. Therefore, it is unlikely that simply transposing the signaling paradigms discovered in Drosophila onto I. scapularis will yield accurate information about tick humoral immunity. Indeed, we discovered that the IMD relay in I. scapularis functions differently than the IMD pathway in Drosophila (Figure 4)97. Furthermore, B. burgdorferi and A. phagocytophilum trigger

I. scapularis IMD pathway activation despite lacking the PGN agonists described in the

Drosophila model79,97.

1 McClure Carroll EE, et al. (2019). p47 licenses activation of the immune deficiency pathway in the tick Ixodes scapularis. Proc Natl Acad Sci U S A 116(1):205-210. 34

The I. scapularis IMD signaling network is regulated by XIAP, an E3 ubiquitin ligase that promotes signal transduction through K63-linked ubiquitylation97,99. How

XIAP activates the IMD pathway in response to microbial infection is poorly characterized in ticks. To find candidate substrates of XIAP, we co-immunoprecipitated recombinant I. scapularis XIAP incubated with infected tick cell lysate and identified the proteins by mass spectrometry. The evolutionarily conserved protein p47 was found in the pulldown fraction. However, co-immunoprecipitations can sometimes lead to spurious results, as non-physiological interactions may occur.

Therefore, we sought to validate the XIAP-p47 interaction using a variety of different approaches, including standard protein biochemistry techniques and SPR.

Specifically, we hypothesized that XIAP binds to p47. Ticks have been understudied despite their considerable global disease burden and increasing public health threat3-6,19,30.

Our findings will improve our understanding of XIAP enzymology as well as define additional functions of p47 in arthropods. By characterizing the interaction between

XIAP and its candidate substrate p47, we will close a critical gap in our knowledge of

IMD signaling in I. scapularis.

3.2 Results

Mass spectrometry was used to identify the potential binding partners of XIAP.

Common “sticky” proteins and contaminants (e.g. keratin) were removed from the list.

The algorithm Ubpred was used to predict whether candidate proteins were likely to be ubiquitylated119. The top hit is most similar to the evolutionarily conserved protein p47

35 and was likely misannotated as “protein tyrosine phosphatase” as this tick protein lacks genetic evidence of phosphatase activity (Table 4).

Table 4: Putative interacting partners of I. scapularis XIAP.

Protein ID* FASTA name Ubiquitylation Prediction B7PEL3 Protein tyrosine phosphatase (p47) Yes B7QF96 Putative uncharacterized protein No B7Q4P0 Putative uncharacterized protein Yes B7Q0Q1 Putative uncharacterized protein Yes B7PIP9 Ankyrin 2,3/unc44, putative Yes B7PJK3 Putative uncharacterized protein No B7PSX9 Tropomyosin invertebrate, putative No B7PU84 Putative uncharacterized protein Yes B7Q121 Putative uncharacterized protein No B7Q2T7 Putative uncharacterized protein No B7Q740 Putative uncharacterized protein No B7Q815 Secreted protein Yes B7QD48 Putative uncharacterized protein No B7QGQ3 Putative uncharacterized protein No *After removal of contaminant proteins, XIAP, and GST.

We validated the XIAP-GST co-immunoprecipitation by interrogating the molecular interactions between XIAP and p47 in a physiological environment.

Recombinant plasmids encoding I. scapularis XIAP tagged with HA and p47-FLAG were generated and transfected into HEK 293T cells. We co-immunoprecipitated XIAP-

HA and detected the FLAG tag, and vice versa, indicating that I. scapularis XIAP and p47 interact under physiological conditions (Figure 6A). Next, we expressed p47-His in

Drosophila S2* cells and purified the recombinant protein for binding experiments.

Using ELISA, we showed that p47-His and XIAP-GST interacted in vitro (Figure 6B).

We then employed SPR to calculate the dissociation constant (KD) of the XIAP-p47 interaction and measured the binding affinity to be in the nanomolar range (Figure 6C).

Figure 6: The I. scapularis proteins XIAP and p47 interact. The binding of XIAP and p47 was validated using three different approaches. A) p47-FLAG and XIAP-hemagglutinin (HA) were co-expressed in human embryonic kidney (HEK) 293T cells and co- immunoprecipitation was assessed by western blotting. Recombinant p47-His and XIAP tagged with glutathione S-transferase (GST) were shown to interact through B) enzyme-linked immunosorbance assay (ELISA). Equimolar amounts of p47-His or BSA were incubated with increasing amounts of XIAP-GST to assess binding by ELISA. C) Surface plasmon resonance (SPR) was performed by coating anti-GST antibodies onto a Biacore chip, followed by capture of XIAP-GST or GST alone. P47-His was diluted to the indicated concentrations in Biacore buffer and was flowed over the chip. Means +/- standard of error (SEM) are graphed. Statistical significance was evaluated by a one-way analysis of variance (ANOVA) followed by Tukey post-hoc testing to determine which conditions were different from baseline. Experiments were performed at least twice. IB, immunoblot; IP, immunoprecipitation; KD, dissociation constant; #, P < 0.0001. Adapted from McClure Carroll, et al 111.

Our next research question focused on using p47 deletion mutants to determine which domain of p47 was required for binding to XIAP (Figure 7A). We cloned truncated p47 constructs into the pCMV-FLAG vector and co-transfected these plasmids one-by-one with xiap-HA into HEK 293T cells. Only p47-FLAG mutants with UBX domains were able to co-immunoprecipitate XIAP-HA, strongly suggesting that the UBX domain is responsible for mediating the interaction between p47 and XIAP in I. scapularis (Figure 7B). To validate this result, we expressed the His-tagged ΔUBA and the ΔUBX p47 truncation mutants in S2* cells and used these recombinant proteins in an

ELISA-based binding assay. We showed that the binding between ΔUBX p47-His and

XIAP-GST was approximately 50% less than the interaction between XIAP-GST and the

WT and ΔUBA p47-His proteins (Figure 7C). Collectively, our results demonstrated that

I. scapularis XIAP and p47 interact and suggest that the UBX domain of p47 is required for the binding of these two tick proteins.

Figure 7: p47 interacts with XIAP through the ubiquitin-like (UBX) domain. A) p47-FLAG deletion constructs were transfected into HEK 293T cells with xiap-HA and B) subjected to co-immunoprecipitation and western blotting. C) 50 ng p47-His or domain deletion mutants were incubated with 5 μM XIAP-GST in triplicate (ELISA). The binding was expressed as a percentage of wild-type (WT) p47: XIAP-GST binding. Statistical significance was evaluated by a one-way ANOVA followed by a Tukey post-hoc test. Experiments were performed at least twice. (-), mock transfection; ns, not significant; UBA, ubiquitin-binding domain; SEP, Shp1, eyes closed, p47 domain; * P < 0.05. Adapted from McClure Carroll, et al 111.

3.3 Discussion

Homologs to several key IMD pathway genes described in Drosophila remain unidentified in the I. scapularis genome62,94-96,98, leaving the underlying regulatory mechanisms of the IMD pathway unknown in ticks. XIAP has been well characterized as the central regulator of the I. scapularis IMD pathway97,99; however, no substrates of

XIAP had been identified prior to the discovery of p47 as a putative binding partner. To our knowledge, the interaction between p47 and a ubiquitin ligase has never been reported. Ubiquitin ligases typically bind transiently and weakly to substrates when

39 compared to other protein-protein interactions120,121. In this case, however; XIAP and p47 bound to each other more strongly than expected (Figure 6C).

Canonically, p47 interacts with the ATPase p97, and targets this enzyme to various subcellular locations based on monoubiquitin recognition conferred by the N- terminal UBA domain104,105,122. The C-terminal UBX domain structurally resembles ubiquitin; however, it lacks the ability to be conjugated to substrates123. The Shp1, EYC, p47 (SEP) domain located between the UBA and UBX domains has no known function, though one study suggested that the structure resembled a cathepsin L inhibitor105,124.

Finally, the UBX domain and a contact point within SEP domain allow p47 to interact with p97105,107,123. Consistent with this literature, we observed that p47 and XIAP interacted through the UBX domain of p47 (Figure 7).

CHAPTER 4: P47 IS AN ENZYMATIC SUBSTRATE OF XIAP1

4.1 Introduction

Fine-tuned regulatory programs govern the magnitude and duration of inflammation125. Immune signaling networks are controlled in part through ubiquitylation126, a process which results in the post-translational conjugation of ubiquitin proteins onto substrates127. A series of ATP-dependent reactions covalently links the C- terminus of a ubiquitin monomer to a lysine or methionine residue encoded in a substrate protein (Figure 8)126-129. Each enzyme in the ubiquitylation cascade confers increasing specificity, enabling the cell to exert tight control over ubiquitylation dynamics130.

Figure 8: Enzymatic reactions involved in ubiquitylation. An E1 activating enzyme activates ubiquitin (U) and transfers the activated protein to an E2 conjugating enzyme. The Really Interesting New Gene (RING)-type E3 ubiquitin ligase coordinates the transfer of the ubiquitin monomer from the E2 conjugating enzyme to a lysine or the N-terminal methionine in the substrate protein. Mono- and polyubiquitylation is possible, with diverse ubiquitin chain topology leading to different fates for the substrate protein. Deubiquitinases (DUBs) cleave and remodel ubiquitin chains. All reactions require ATP. Adapted from Metzger, et al129.

1 McClure Carroll EE, et al. (2019). p47 licenses activation of the immune deficiency pathway in the tick Ixodes scapularis. Proc Natl Acad Sci U S A 116(1):205-210. 41

Ubiquitin is an 8.5 kDa protein that contains seven lysine residues (K6, K11, K27,

K29, K33, K48, K63) and an N-terminal methionine residue127. Ubiquitin monomers can be conjugated to substrate proteins singly (e.g., monoubiquitylation) or in complex polyubiquitin chains consisting of various linkages and chain topologies126. The type of ubiquitin linkage also has implications for the fate of the modified substrate protein.

Ubiquitins linked at the K48 position typically tag the substrate for proteasomal degradation, while K63-linked ubiquitin modifications promote signal transduction or regulate protein-protein interactions127. Deubiquitinases (DUBs) also confer an additional layer of regulation by stripping ubiquitin from substrates and remodeling chain linkages126-128.

Regulation of immune signaling through ubiquitylation has been understudied in arthropod vectors of disease, especially in the tick I. scapularis128,130. Previously, we demonstrated that the E3 ubiquitin ligase XIAP positively regulates tick IMD signaling97,99. However, the link between XIAP-mediated ubiquitylation and IMD signal transduction was poorly understood, as no substrate of XIAP had been identified. In Aim

1, I proposed to investigate the molecular and biophysical interactions between XIAP and the putative substrate p47. However, whether XIAP and p47 interacted functionally as a ubiquitin ligase and substrate pair was unknown.

To our knowledge, there has been only one detailed characterization of post- translational modifications of p47. Phosphorylation of rat p47 at residue 140 blocked its ability to bind Golgi stacks and target p97 to the Golgi membranes for repair after mitosis107, suggesting that post-translational modifications may regulate p47 function. In this section, we investigated the specific hypothesis that XIAP polyubiquitylates p47 in a

K63-dependent manner using a cell-free ubiquitylation approach. Ultimately, these findings could lead to the discovery of a novel function for p47 as a participant in IMD signaling in arthropods.

4.2 Results

Having demonstrated that XIAP and p47 interact (Figures 6 & 7), we next aimed to determine whether p47 was an enzymatic substrate of XIAP. We employed a cell-free ubiquitylation assay to assess whether p47 could be polyubiquitylated by XIAP in vitro.

The reaction consisted of the recombinant proteins human Ube1 (E1), human Uev1a

(E2), I. scapularis Bendless-GST (E2), I. scapularis XIAP-GST (E3), human ubiquitin monomers, I. scapularis p47-His, and ATP (Figure 9A).

Figure 9: The E3 ubiquitin ligase XIAP polyubiquitylates p47 in a K63-dependent manner. A) A cell-free ubiquitylation reaction containing the components necessary for ubiquitylation was incubated for 1h at 37°C prior to isolating p47-His with nickel resin. The isolated p47-His protein was divided into three aliquots and used for western blots to detect B) K63-specific ubiquitylation and the His tag as well as C) K48-specific ubiquitylation. Western blots panels are from separate but representative experiments due to limited material. Experiments were repeated at least twice. Numeric labels on the left side of the western blot panels indicate molecular weight (kDa). Ub, ubiquitin; Bend, Bendless; IN, input. Adapted from McClure Carroll, et al111.

The ubiquitylation assay was performed with and without p47, the candidate substrate, and both reactions were incubated with nickel resin. We observed that XIAP generated free polyubiquitin chains in the absence of substrate as previously reported

(Figure 9B, K63 blot, flow-through [FT], right-most lane)97,99, indicating our enzymes were functional. We next probed the isolated p47-His (IP) for the His tag and K63- dependent polyubiquitylation signals (Figure 9B). We detected higher molecular weight species after the reaction (Figure 9B, His blot, compare input (IN) and IP p47-His), as well as a smear that overlapped with p47-His (Figure 9B, K63 blot, left-most lane). XIAP did not ubiquitylate p47-His in a K48-dependent manner (Figure 9C). Altogether, these results demonstrated that I. scapularis p47 was ubiquitylated by XIAP in a K63- dependent manner, and strongly suggest that p47 is an enzymatic substrate of XIAP.

4.3 Discussion

In the previous chapter we validated that XIAP binds to p47 and determined that the interaction between the two proteins is dependent on the UBX domain of p47

(Figures 6 & 7). However; it was unknown if the interaction between XIAP and p47 was a bona fide ubiquitin ligase-substrate pairing. Here, we demonstrated that XIAP, together with human Ube1 (E1) and the heterodimeric E2 conjugating enzymes Bendless and

Uev1a, ubiquitylated p47 in a K63-dependent manner (Figure 9B). These results are consistent with our previous work showing that XIAP performs K63-linked autoubiquitylation in the presence of Ube1, Bendless/Uev1a, ATP, and ubiquitin97,99.

Due to technical limitations, ectopically expressing proteins in tick cells is currently not feasible for routine laboratory applications. Therefore, we employed an in

44 vitro reaction system, which offered the distinct advantage of avoiding the off-target ubiquitylation that could occur in a surrogate cell line (e.g., HEK 293T). The in vitro assay demonstrated specifically that XIAP is the E3 ubiquitin ligase responsible for ubiquitylating p47, since all the inputs were known and controlled. These assays require large amounts of protein. Unfortunately, p47 can only be expressed in a eukaryotic system, likely due to its requirement for post-translational modifications or toxicity to bacterial cells. We purified recombinant His-tagged p47 from Drosophila S2* cells; however, the yields were poor. Therefore, the ubiquitylation reactions were performed on a small scale, and it was challenging to isolate sufficient amounts of ubiquitylated p47-

His from the reaction to perform more sophisticated experiments.

XIAP is a RING-type E3 ubiquitin ligase99. Members of this class of ubiquitin ligases physically bridge the E2 conjugating enzyme and the substrate protein, but do not contain catalytic domains (Figure 8)129. That is, RING-type ubiquitin ligases serve to coordinate the transfer of the activated ubiquitin monomer from the E2 conjugating enzyme to the substrate protein. Accordingly, the function of the E3 ubiquitin ligase is linked to the specific E2 enzyme it binds, so the pairing of these two enzymes impacts the ubiquitylation pattern on the substrate protein129. Previous work has demonstrated that the

K63 ubiquitin linkage promotes signal transduction by generating scaffolds for protein recruitment in many immune signaling pathways, including IMD79,85,131,132. Our finding that XIAP polyubiquitylates p47 in a K63-dependent manner suggests that p47 may belong to the I. scapularis IMD signaling relay.

CHAPTER 5: P47 LICENSES ACTIVATION OF THE IMMUNE

DEFICIENCY PATHWAY IN THE TICK I. SCAPULARIS1

5.1 Introduction

In the Drosophila model, ubiquitylated substrates of IAP2 positively regulate the

IMD pathway, leading to the cleavage of the NF-κB molecule Relish and the production of AMPs (Figure 4)79,85. However, the link between XIAP-mediated ubiquitylation and

IMD pathway signaling regulation remains poorly understood in ticks, as homologs to core Drosophila IMD pathway genes are absent in the I. scapularis genome62,94-96,98. We identified the protein p47 as a substrate of XIAP (Figure 6) and demonstrated that XIAP polyubiquitylates p47 in a K63-dependent manner (Figure 9)111. Whether p47 was involved in I. scapularis IMD pathway signaling remained unproven.

We employed several approaches to investigate the specific hypothesis that p47, the substrate of XIAP, is a member of the IMD pathway in I. scapularis. First, we examined biochemical evidence for the role of p47 in tick IMD pathway signaling. p47 was silenced in vitro prior to stimulation with A. phagocytophilum. We then determined whether p47 expression impacted signal transduction by observing key IMD pathway signaling events. We also checked if p47 interacted with any other canonical members of the IMD pathway through co-immunoprecipitation. Finally, we demonstrated that p47 impacts pathogen infection in I. scapularis ticks. These data improve our understanding of the IMD pathway in I. scapularis ticks, which serve as a model for non-insect

1 McClure Carroll EE, et al. (2019). p47 licenses activation of the immune deficiency pathway in the tick Ixodes scapularis. Proc Natl Acad Sci U S A 116(1):205-210. 46 arthropods. Our findings may inform future strategies for preventing vector-borne diseases.

5.2 Results

We demonstrated that p47 interacts with (Figures 6 & 7) and is polyubiquitylated by XIAP (Figure 9), the E3 ubiquitin ligase responsible for centrally regulating the IMD signaling pathway in I. scapularis in response to microbial infection97. Next, we investigated whether p47 was a member of the IMD pathway signaling network by silencing p47 expression in tick cells and stimulating the pathway with A. phagocytophilum. We monitored uninfected cells (“(-)”) and early (one, five, and 60 minutes) and late (48 hours) activation of the I. scapularis IMD pathway. These timepoints were chosen because independent laboratories have demonstrated Relish cleavage within minutes of microbial stimulation85,91,97. The activation events, which included Kenny (IKKγ) accumulation, IKKβ phosphorylation, and Relish cleavage, were assessed by western blot. Our findings revealed that when p47 was silenced, we observed reduced Kenny accumulation after five minutes (Figure 10A) and no IKKβ phosphorylation above baseline (Figure 10B), suggesting that in the absence of p47, the

IKK complex is impaired. Furthermore, Relish cleavage (indicated by arrowheads) was barely detectable in the p47-silenced cells, indicating that defects in IMD pathway signaling arise when p47 is absent (Figure 10C).

Figure 10: p47 knockdown impairs IMD pathway signaling. IDE12 cells were transfected with silencing p47 (si-p47 and si) or control constructs (sc-p47 and sc) and stimulated with Anaplasma phagocytophilum multiplicity of infection (MOI) 150. Cytoplasmic and nuclear proteins were extracted and the IMD pathway signaling events A) Kenny (IKKγ) accumulation, B) IKKβ phosphorylation, and C) Relish cleavage were assessed by western blot. Arrowheads indicate bands of interest. Densitometry analysis was performed to normalize the target to the loading control. The normalized value for each target and time point was divided by the corresponding scrambled control and multiplied by 100. Three experiments were performed. Means± SEM are graphed. Two- way ANOVA followed by Sidak post-hoc testing was performed to determine statistical significance compared to unstimulated samples. (-), unstimulated; E, early; L, late; p, phosphorylated; * P < 0.05; ** P < 0.005; **** P < 0.0001. Adapted from McClure Carroll, et al111.

p47 was shown to bind polyubiquitylated NEMO to negatively regulate NF-κB signaling in the human system110. In I. scapularis and other arthropods, NEMO is known as Kenny. Given that disrupting p47 levels in vitro impaired IMD pathway activation

(Figure 10), we next sought to determine whether p47 interacted with Kenny in the tick system. Recombinant I. scapularis Kenny-GST and p47-His interacted in vitro, as demonstrated by ELISA (Figure 11A). We also co-transfected kenny-HA and p47-FLAG constructs into HEK 293T cells to investigate whether the proteins interacted in a cellular environment. As expected, we detected the HA tag in the p47-FLAG pulldown and vice versa, indicating that I. scapularis p47 and Kenny interacted in cells (Figure 11B).

We predicted that if the ubiquitylation of p47 were to be prevented, p47 would no longer bind to Kenny. We selected five lysine residues in the p47-FLAG construct that were either candidates for ubiquitin modifications as predicted by Ubpred119 or located within the putative regulatory region between the UBA and SEP domains105,107. Point mutations were made to replace the lysines at positions 109, 139, 141, 154, and 300 with arginine residues, generating a mutant p47-FLAG construct (*p47-FLAG) (Figure 11C).

Lysine to arginine replacements are typically made in ubiquitin biology to prevent ubiquitylation while retaining the positively charged amino acid to maintain proper protein folding133. Using a co-immunoprecipitation approach, we demonstrated that the

*p47-FLAG mutant protein was unable to interact with Kenny-HA (Figure 11D).

Figure 11: p47 interacts with Kenny, the regulatory subunit of the IκB kinase (IKK) complex A) Recombinant p47-His and Kenny-GST were shown to interact through ELISA. B) Kenny-HA and p47-FLAG were co-expressed in HEK 293T cells and co-immunoprecipitation was assessed by western blotting. C) Five lysines (indicated by arrows) were mutated to arginine residues by site-directed mutagenesis, generating a mutant *p47-FLAG construct. D) Kenny-HA, p47-FLAG, and *p47-FLAG were co-transfected into HEK 293T cells followed by co-immunoprecipitation to evaluate Kenny-p47 binding. Statistical significance was evaluated by one-way ANOVA followed by Tukey post-hoc testing to determine which conditions were different than baseline. Experiments were performed twice. ns, not significant; * P < 0.05; ** P < 0.01. Published in McClure Carroll, et al111.

Our findings in Figures 10 and 11 suggested that p47 is a member of the IMD pathway signaling network in I. scapularis ticks. We next sought to determine whether p47 affected microbial infection in vitro by silencing p47 in ISE6 cells and infecting with

A. phagocytophilum (Figure 12A). The expression of p47 was reduced by 31% in the cells transfected with siRNA targeting p47 (si-p47) compared to control cells (sc-p47)

(Figure 12B), which corresponded to a threefold increase in A. phagocytophilum burden

(Figure 12C).

Figure 12: p47 silencing increases A. phagocytophilum burden in tick cells. A) p47 in ISE6 cells was silenced followed by A. phagocytophilum (Ap) MOI 50 infection. B) P47 expression and C) A. phagocytophilum transcripts were quantified by quantitative reverse transcription polymerase chain reaction (qRT-PCR) and normalized to I. scapularis β-actin prior to calculating fold- change (FC). Results are graphed as means ± SEM and were assessed by Student’s T-tests. ** P < 0.01; *** P <0.001. Published in McClure Carroll, et al111.

We next investigated the role of p47 during microbial colonization of I. scapularis ticks. Accordingly, I. scapularis nymphs were microinjected with si-p47 or sc- p47 siRNA. A subset of microinjected ticks was pooled and lysed to obtain protein extracts, while the rest of the ticks fed to repletion on B. burgdorferi- or A. phagocytophilum-infected mice (Figure 13A). No difference in weight was observed in

51 p47-silenced versus the control treatment, indicating that p47 did not impact tick feeding on A. phagocytophilum- (Figure 13B) or B. burgdorferi-infected mice (Figure 13C). The unfed tick lysate was incubated with either XIAP-GST or GST alone to show successful p47 knockdown by ELISA (Figure 13D), as single ticks did not yield enough material to detect p47 levels by western blot. Engorged ticks were subjected to qRT-PCR to determine p47 expression and bacterial burden. Consistently, despite modest silencing

(Figure 13E), p47-deficient ticks acquired more A. phagocytophilum (Figure 13F) and B. burgdorferi (Figure 13G) when compared to the control ticks. Collectively, the data presented this chapter strongly suggest that p47 participates in antimicrobial defenses by promoting IMD pathway signal transduction in I. scapularis.

Figure 13: Silencing p47 increases microbial infection in vivo. A) I. scapularis nymphs were microinjected with si-p47 or sc-p47 siRNA and fed to repletion on infected mice. P47 silencing did not impact tick feeding success on B) A. phagocytophilum- or (C) B. burgdorferi-infected mice as assessed by tick weight. D) A subset of unfed, p47-silenced ticks was pooled, lysed, and incubated with XIAP-GST or GST in an ELISA binding assay to assess p47 knockdown. Engorged ticks were subjected to qRT-PCR to quantify (E) p47, (F) A. phagocytophilum 16S rRNA, and (G) B. burgdorferi recA transcripts. All qRT-PCR values were normalized to I. scapularis β-actin prior to calculating FC. Three independent experiments were plotted and analyzed together. Means ± SEM are graphed. Statistical significance was assessed by two-way Student’s T-tests. * P < 0.05; *** P < 0.001; recA, recombinase A. Adapted from McClure Carroll, et al111.

5.3 Discussion

Genetic tools for arthropod research outside of the Drosophila community are extremely limited. It is currently not feasible to ablate genes or to routinely ectopically express proteins in tick cell lines or animals. The relatively long life cycle and the inconvenient developmental characteristics of hard ticks have contributed to the genetic intractability of I. scapularis134. Therefore, since genetic knockouts are unavailable, we rely on RNA interference (RNAi) to manipulate gene expression in I. scapularis ticks.

SiRNA constructs can be delivered into the tick midgut, salivary glands, or hemocoel to modulate gene expression135-137. Changes in bacterial load are typically modest97,99.

Nevertheless, despite incomplete knockdowns and the technical challenges of working with ticks in the laboratory setting, we demonstrated that p47 promotes signal transduction in the I. scapularis IMD pathway (Figure 10), interacts with Kenny, another member of the canonical IMD signaling network (Figure 11), and restricts microbial infection in cells (Figure 12) and ticks (Figure 13).

Silencing p47 expression resulted in persistent defects in the biochemical relay of the IMD pathway (Figure 10). However, we noticed that some Relish cleavage still occurred at one and five minutes post-stimulation in p47-silenced cells (Figure 10C).

Additional regulation may explain the residual signaling in the I. scapularis IMD signaling network. For example, the Drosophila HOIL-1L interacting protein (HOIP) ortholog linear ubiquitin E3 ligase (LUBEL) works in concert with IAP2 to activate

Relish through methionine 1-dependent ubiquitylation of Kenny in flies138. This is expected, as the transduction of NF-κB signals is likely tightly modulated through redundancy to prevent aberrant inflammation and limit fitness costs110,125.

We showed that p47 binds to Kenny, the regulatory component of the IKK complex (Figures 11A & B). In humans, p47 negatively regulates NF-κB signaling by interacting with the mammalian homolog of Kenny, NEMO110. In that report, p47, independent of p97, bound to polyubiquitylated NEMO and led to its degradation through an unknown mechanism. In I. scapularis, the interaction between Kenny and p47 has the opposite effect and seems to promote signal transduction, as silencing p47 led to defects in IMD signaling (Figure 10) and increased microbial infection of cells (Figure 12) and ticks (Figure 13).

In our model of the tick IMD network, XIAP-mediated K63-dependent polyubiquitylation of p47 leads to IMD pathway activation. Therefore, we predicted that if ubiquitylation of p47 could be prevented, binding to Kenny would be lost. Indeed, our data showed that the mutant *p47-FLAG protein could not bind Kenny-HA in the HEK

293T system (Figure 11D). Perhaps the abrogation of binding we observed is due to the inability of the *p47-FLAG mutant to be ubiquitylated at those critical residues. Our data suggested that XIAP activates p47 via polyubiquitylation, perhaps forming a scaffold consisting of K63-linked ubiquitin monomers that enables the recruitment of other members of the I. scapularis IMD signaling pathway, as has been described for other immune signaling cascades79,85,131,132.

We demonstrated that silencing p47 led to enhanced microbial infection of ticks by two clinically relevant I. scapularis-borne pathogens (Figures 12 & 13), strongly suggesting that p47 is involved in the antimicrobial defenses of ticks. We observed approximately two- to fourfold increases in microbial burden in nymphs that were treated with si-p47, despite reducing p47 expression by only about 25%. The factors that

55 influence siRNA silencing efficiency are not well understood in ticks, so the siRNA constructs may benefit from experimental optimization134. An alternative explanation for the discrepancy between modest p47 knockdown and increased microbial burdens may be attributed to tick biology. In a typical experiment, the I. scapularis nymphs are microinjected and placed on the mice after resting overnight. The ticks feed for up to five days on the infected mice, after which they detach and are collected for qRT-PCR.

Therefore, the p47 expression that we measured almost one week post-microinjection likely did not reflect the true expression of p47 during feeding and acquisition of A. phagocytophilum and B. burgdorferi. Furthermore, in Drosophila, the expression of some

IMD pathway genes is controlled by Relish93,139-141. If p47 expression is induced by

Relish in I. scapularis, the true extent of p47 silencing may be obscured in our experimental model.

Nevertheless, despite the limitations of tick genetic tools, we demonstrated that p47 participates in I. scapularis IMD pathway signal transduction. Our findings provide evidence for a molecular mechanism that links XIAP polyubiquitylation to Relish activation and IMD pathway-mediated antimicrobial responses in ticks. Collectively, these data extend our knowledge of tick humoral immune signaling pathways in the context of vector-borne diseases and highlight the danger of relying solely on model organisms for characterizing vector immunity.

CHAPTER 6: CONCLUSIONS AND FUTURE DIRECTIONS1

Drosophila has served as a powerful model for innate immunity for decades60.

Yet, even though many arthropods serve as disease vectors, immune signaling pathways in non-model organisms like the Lyme disease tick I. scapularis remain poorly defined62.

For example, the Drosophila model was used to discover and characterize the IMD pathway, a humoral immune signaling network that, upon activation, results in the production of AMPs that kill microbes74,79. The details of IMD signaling in ticks were unknown, as the I. scapularis genome lacks homologs to key IMD pathway genes characterized in Drosophila61,62,79,95,96,98.We previously discovered that the I. scapularis

IMD signaling network is regulated by XIAP-mediated K63 ubiquitylation and limits the infection of clinically relevant tick-borne microbes97,99. However, the link between

XIAP-regulated IMD pathway signaling and Relish activation was unclear, because no substrates of XIAP had been identified.

We recently uncovered p47 as potential substrate of XIAP and hypothesized that

XIAP binds and polyubiquitylates the protein p47 in a K63-dependent manner during

IMD pathway activation in I. scapularis ticks. This central hypothesis was investigated in three specific aims. In Aim 1, we validated the interaction between XIAP and p47, and determined that the UBX domain of p47 was responsible for mediating protein-protein interactions. In Aim 2, we demonstrated that p47 is a substrate of XIAP, and that the ubiquitin linkage conjugated to p47 by XIAP is K63-linked ubiquitin. In Aim 3, we assessed whether p47 was a member of the IMD signaling relay in ticks. We demonstrated that silencing p47 led to defects in IMD signal transduction and increased

1 McClure Carroll EE, et al. (2019). p47 licenses activation of the immune deficiency pathway in the tick Ixodes scapularis. Proc Natl Acad Sci U S A 116(1):205-210. 57 microbial infection of ticks. We also showed that p47 interacted with Kenny, another member of the canonical IMD signaling pathway.

Our model suggests that, upon microbial infection, p47 is ubiquitylated by XIAP in a K63-dependent manner, generating a ubiquitin scaffold essential for recruiting the additional proteins required for signal transduction (Figure 14). Ubiquitylated p47 binds to Kenny, the regulatory subunit of the IKK complex, which leads to the phosphorylation of Relish by IKKβ. As in Drosophila, IMD signaling culminates in the cleavage and nuclear translocation of Relish, and ultimately the activation of immune response genes.

Our findings provide strong evidence that the pairing of XIAP and p47 licenses IMD pathway activation in I. scapularis (Figure 14) and suggest several future research directions.

Figure 14: Model of IMD pathway signaling in I. scapularis. IMD signaling in I. scapularis ticks is triggered by A. phagocytophilum and B. burgdorferi and leads to the cleavage and nuclear translocation of Relish (N-Rel) to promote antibacterial responses. The E3 ubiquitin ligase XIAP, together with the heterodimer Bendless: Uev1a, binds to p47 and regulates the signal transduction through K63-dependent polyubiquitylation. Ubiquitylated p47 is responsible for recruiting Kenny in the tick IMD network. R, unidentified receptor; C, cytoplasm; N, nucleus. Adapted from McClure Carroll, et al111.

6.1 The role of the XIAP-p47 complex in IMD signaling in ticks

p47 canonically interacts with the ATPase p97 in a large protein complex, which consists of a p97 hexamer assembled in a wheel-like structure with individual p47 molecules bound to each p97 protein104,106. Another study concluded that only three p47 molecules bound to one p97 hexamer104, suggesting that the stoichiometry of the p47 and p97 complex may change depending on the cellular context. To our knowledge, no reports of p47 binding to a ubiquitin ligase have been published prior to our study111.

Therefore, determining whether the stoichiometry and the binding parameters of the p47-XIAP interaction differ from that of the p47-p97 complex may impact our understanding of p47 biology. The structural arrangement of p47 and XIAP may also have implications for ubiquitylation and IMD signal transduction. For instance, in the

Drosophila IMD pathway, Imd proteins assemble into large amyloid fibrils upon activation of the pathway. Without Imd aggregation, efficient signal transduction is prevented and IMD responses are blunted142. Large protein complexes, or

“signalosomes,” are also important for productive signal transduction in NF-κB and other inflammatory cascades143,144. Perhaps XIAP and p47 nucleate a signalosome required for

IMD pathway-mediated Relish activation in I. scapularis.

In Chapter 4, we demonstrated that p47 is the enzymatic substrate of XIAP

(Figure 9). While we know that p47 is K63-polyubiquitylated by XIAP, the identity of the target lysine remains unknown. Diglycine remnant profiling could be performed in the future to ascertain which lysine is ubiquitylated133,145. In diglycine remnant profiling, the ubiquitylated protein is digested with trypsin, which cleaves the protein as well as the ubiquitin and leaves a G-G adduct on the modified residue. Mass spectrometry can be

60 used to detect this diglycine remnant, and given enough coverage, the ubiquitylation sites can be mapped133,145. We would validate the results by using site-directed mutagenesis to replace the candidate lysine with arginine. The resulting p47 mutant would likely adopt the correct folding, as the charge and size of the amino acid side chains are similar; however, the protein would lack the capability to be ubiquitylated at that site. Next, we would express the protein and perform the cell-free ubiquitylation assay to show that ubiquitylation is abrogated.

6.2 Molecular details of p47-mediated IMD signaling

Although we determined that p47 interacts with both XIAP and Kenny to promote

IMD responses (Figure 14), the detailed molecular mechanism of IMD pathway signal transduction remains unknown. Our data suggest that the K63-dependent polyubiquitylation of p47 by XIAP is an activating signal that enables p47 to bind to

Kenny. However; there may be other proteins involved and almost certainly additional regulatory mechanisms in place to exert control over Relish activation. In Drosophila,

IAP2, the analog of XIAP, polyubiquitylates several proteins in the IMD pathway, including Imd (Figure 4). We showed that p47 is the substrate of XIAP (Figures 6 & 9); however, p47 and Imd have little in common structurally, and p47 lacks the death and cryptic receptor-interacting protein (RIP) homotypic interaction motif (cRHIM) homology domains that are required for downstream Imd signaling in flies142. These data suggest that p47 is likely not the central adaptor molecule that replaces Imd one-to-one in the I. scapularis IMD pathway. Given that insects and non-insect arthropods diverged over 500 million years ago62,146, and that ticks and flies have evolved under different

61 selection pressures, it is possible that the IMD pathway exists as two distinct signaling circuits in each taxonomic group.

Another hypothesis is that the tick mode of IMD signaling exists in insects as well; however, it is only activated under specific conditions that have not yet been explored in Drosophila. In our previous work, Drosophila S2* cells responded to A. phagocytophilum-stimulation via the IMD pathway, even though this bacterium lacks characterized IMD agonists97,147. Furthermore, a series of recent papers implicated other proteins in the IMD-dependent response to viral and bacterial infections, suggesting that

IMD signaling may be more complex than previously characterized148-151. Our data, together with these recent findings, suggest that IMD signaling relays may adopt complex regulatory circuits to contextually tune inflammatory responses. Future experiments should determine if p47 also plays a role in the IMD pathway in other arthropods and attempt to characterize additional modes of IMD signaling in ticks.

6.3 Linking p47 to other functions in I. scapularis

p47 was first discovered as a cofactor of the ATPase p97 and participates with p97 in the remodeling of organelle membranes104. Like p47, p97 is highly conserved and is encoded in the genome of I. scapularis152,153. However, we did not find p97 in the list of potential XIAP binding partners identified through our initial XIAP-GST co- immunoprecipitation experiment (Table 4). In other contexts, p47 was shown to act independently of p97 in the negative regulation of NF-κB110, so we did not address in this study whether p97 was also involved in IMD pathway signaling. Future experiments should determine the role, if any, for p97 in I. scapularis immunity.

The body of published literature on p47 mostly focuses on the role of p47 in targeting p97 to various subcellular locations100,102,104,107,108,152. p97 hydrolyzes ATP to provide energy for membrane remodeling152. In yeast, p47 was shown to bind the ubiquitin-like molecule Atg8, thereby targeting p97 to the nascent autophagosome and perhaps regulating the growth and closure of autophagic membranes109. The potential connection to autophagy in other systems, such as ticks, is worth exploring. Ticks feed only once before molting to the next life stage15. Therefore, it is likely that they engage autophagy pathways in times of starvation; however, this hypothesis has not been explored in tick biology. In other systems, autophagic clean-up of immune signaling complexes is one mechanism of resolving inflammation154. Future studies could determine if IMD pathway signaling is mitigated by autophagy. Perhaps p47 links these two processes, and future studies addressing the role of p47 in connecting immunity to other aspects of cell biology in I. scapularis are warranted.

6.4 Conclusions

In conclusion, this work addressed an open question in the field of tick immunity.

We previously established that the E3 ubiquitin ligase XIAP regulates the I. scapularis

IMD pathway, and that the IMD signaling relay is triggered in response to tick- transmitted microbes97,99. E3 ubiquitin ligases ubiquitylate substrate proteins; however, no substrate for XIAP had been identified. This work established that the evolutionarily conserved protein p47 binds to XIAP through the UBX domain. Furthermore, the data demonstrated that XIAP polyubiquitylates p47 in a K63-dependent manner, and that p47 promotes IMD signal transduction in ticks. Altogether, this work highlights the diversity

63 of arthropod immune signaling pathways and may inform future strategies to prevent tick-borne diseases.

REFERENCES

1. World Health Organization. Vector-borne diseases fact sheet. http://www.who.int/mediacentre/factsheets/fs387/en/. Published October 2017. Accessed November 30, 2018.

2. Bardosh KL, Ryan SJ, Ebi K, Welburn S, Singer B. Addressing vulnerability, building resilience: community-based adaptation to vector-borne diseases in the context of global change. Infect Dis Poverty. 2017;6(1):166.

3. Jongejan F, Uilenberg G. The global importance of ticks. Parasitology. 2004;129 Suppl:S3-14.

4. de la Fuente J, Waterhouse RM, Sonenshine DE, et al. Tick genome assembled: new opportunities for research on tick-host-pathogen interactions. Front Cell Infect Microbiol. 2016;6:103.

5. Sonenshine DE, Roe RM. Ticks, people, and animals. In: Sonenshine DE, Roe RM, eds. Biology of Ticks. Vol 1. 2nd ed. New York, NY: Oxford University Press; 2014:3-16.

6. de la Fuente J, Estrada-Pena A, Venzal JM, Kocan KM, Sonenshine DE. Overview: ticks as vectors of pathogens that cause disease in humans and animals. Front Biosci. 2008;13:6938-6946.

7. Estrada-Pena A, Ostfeld RS, Peterson AT, Poulin R, de la Fuente J. Effects of environmental change on zoonotic disease risk: an ecological primer. Trends Parasitol. 2014;30(4):205-214.

8. Gortazar C, Reperant LA, Kuiken T, et al. Crossing the interspecies barrier: opening the door to zoonotic pathogens. PLoS Pathog. 2014;10(6):e1004129.

9. Jones KE, Patel NG, Levy MA, et al. Global trends in emerging infectious diseases. Nature. 2008;451(7181):990-993.

10. Kernif T, Leulmi H, Raoult D, Parola P. Emerging tick-borne bacterial pathogens. Microbiol Spectr. 2016;4(3).

11. Kosoy OI, Lambert AJ, Hawkinson DJ, et al. Novel thogotovirus associated with febrile illness and death, United States, 2014. Emerg Infect Dis. 2015;21(5):760- 764.

12. Paules CI, Marston HD, Bloom ME, Fauci AS. Tickborne diseases - confronting a growing threat. N Engl J Med. 2018;379(8):701-703.

13. Durden LA, Beati L. Modern tick systematics. In: Sonenshine DE, Roe RM, eds. Biology of Ticks. Vol 1. 2nd ed. New York, NY: Oxford University Press; 2014:17-58.

14. Mans BJ, de Klerk D, Pienaar R, Latif AA. Nuttalliella namaqua: a living fossil and closest relative to the ancestral tick lineage: implications for the evolution of blood-feeding in ticks. PLoS One. 2011;6(8):e23675.

15. Sonenshine DE, Roe RM. Biology of Ticks. Vol 2. 2nd ed. New York, NY: Oxford University Press; 2014.

16. Parola P, Raoult D. Ticks and tickborne bacterial diseases in humans: an emerging infectious threat. Clin Infect Dis. 2001;32(6):897-928.

17. Xu G, Fang QQ, Keirans JE, Durden LA. Molecular phylogenetic analyses indicate that the Ixodes ricinus complex is a paraphyletic group. J Parasitol. 2003;89(3):452-457.

18. Stanek G, Wormser GP, Gray J, Strle F. Lyme borreliosis. Lancet. 2012;379(9814):461-473.

19. Eisen RJ, Eisen L. The blacklegged tick, Ixodes scapularis: an increasing public health concern. Trends Parasitol. 2018;34(4):295-309.

20. Centers for Disease Control and Prevention. Life cycle of hard ticks that spread disease. https://www.cdc.gov/ticks/life_cycle_and_hosts.html. Published June 2015. Updated April 2017. Accessed December 10, 2018.

21. Kocan KM, de la Fuente J, Coburn LA. Insights into the development of Ixodes scapularis: a resource for research on a medically important tick species. Parasit Vectors. 2015;8:592.

22. Apanaskevich DA, Oliver Jr., JH. Life cycles and natural history of ticks. In: Sonenshine DE, Roe RM, eds. Biology of Ticks. Vol 1. 2nd ed. New York, NY: Oxford University Press; 2014:59-73.

23. Sonenshine DE, Roe RM. External and internal anatomy of ticks. In: Sonenshine DE, Roe RM, eds. Biology of Ticks. Vol 1. 2nd ed. New York, NY: Oxford University Press; 2014:74-98.

24. Richter D, Matuschka FR, Spielman A, Mahadevan L. How ticks get under your skin: insertion mechanics of the feeding apparatus of Ixodes ricinus ticks. Proc Biol Sci. 2013;280(1773):20131758.

25. Sonenshine DE, Anderson JM. Mouthparts and digestive system. In: Sonenshine DE, Roe RM, eds. Biology of Ticks. Vol 1. 2nd ed. New York, NY: Oxford University Press; 2014:122-162.

26. Simo L, Kazimirova M, Richardson J, Bonnet SI. The essential role of tick salivary glands and saliva in tick feeding and pathogen transmission. Front Cell Infect Microbiol. 2017;7:281.

27. Chmelar J, Kotal J, Langhansova H, Kotsyfakis M. Protease inhibitors in tick saliva: the role of serpins and cystatins in tick-host-pathogen interaction. Front Cell Infect Microbiol. 2017;7:216.

28. Kotal J, Langhansova H, Lieskovska J, et al. Modulation of host immunity by tick saliva. J Proteomics. 2015;128:58-68.

29. Nuttall PA. Wonders of tick saliva. Ticks Tick Borne Dis. 2019;10(2):470-481.

30. Nelder MP, Russell CB, Sheehan NJ, et al. Human pathogens associated with the blacklegged tick Ixodes scapularis: a systematic review. Parasit Vectors. 2016;9:265.

31. Steere AC, Strle F, Wormser GP, et al. Lyme borreliosis. Nat Rev Dis Primers. 2016;2:16090.

32. Weitzner E, McKenna D, Nowakowski J, et al. Long-term assessment of post- treatment symptoms in patients with culture-confirmed early Lyme disease. Clin Infect Dis. 2015;61(12):1800-1806.

33. Koedel U, Fingerle V, Pfister HW. Lyme neuroborreliosis-epidemiology, diagnosis and management. Nat Rev Neurol. 2015;11(8):446-456.

34. Mead PS. Epidemiology of Lyme disease. Infect Dis Clin North Am. 2015;29(2):187-210.

35. Adrion ER, Aucott J, Lemke KW, Weiner JP. Health care costs, utilization and patterns of care following Lyme disease. PLOS ONE. 2015;10(2):e0116767.

36. Nelson CA, Saha S, Kugeler KJ, et al. Incidence of clinician-diagnosed Lyme disease, United States, 2005-2010. Emerg Infect Dis. 2015;21(9):1625-1631.

37. Pritt BS, Respicio-Kingry LB, Sloan LM, et al. Borrelia mayonii sp. nov., a member of the Borrelia burgdorferi sensu lato complex, detected in patients and ticks in the upper midwestern United States. Int J Syst Evol Microbiol. 2016;66(11):4878-4880.

38. Pritt BS, Mead PS, Johnson DKH, et al. Identification of a novel pathogenic Borrelia species causing Lyme borreliosis with unusually high spirochaetaemia: a descriptive study. Lancet Infect Dis. 2016;16(5):556-564.

39. Wormser GP, Shapiro ED, Fish D. Borrelia miyamotoi: an emerging tick-borne pathogen. Am J Med. 2018;132(2):136-127.

40. Krause PJ, Fish D, Narasimhan S, Barbour AG. Borrelia miyamotoi infection in nature and in humans. Clin Microbiol Infect. 2015;21(7):631-639.

41. Rikihisa Y. Anaplasma phagocytophilum and Ehrlichia chaffeensis: subversive manipulators of host cells. Nat Rev Microbiol. 2010;8:328-339.

42. McClure EE, Oliva Chávez AS, Shaw DK, et al. Engineering of obligate intracellular bacteria: progress, challenges and paradigms. Nat Rev Microbiol. 2017;15(9):544-558.

43. Stuen S, Granquist EG, Silaghi C. Anaplasma phagocytophilum--a widespread multi-host pathogen with highly adaptive strategies. Front Cell Infect Microbiol. 2013;3:31.

44. Centers for Disease Control and Prevention. Anaplasmosis epidemiology and statistics. https://www.cdc.gov/anaplasmosis/stats/index.html. Published June 2018. Accessed December 17, 2018.

45. Pritt BS, Sloan LM, Johnson DK, et al. Emergence of a new pathogenic Ehrlichia species, Wisconsin and Minnesota, 2009. New Engl J Med. 2011;365(5):422-429.

46. Johnson DK, Schiffman EK, Davis JP, et al. Human infection with Ehrlichia muris-like pathogen, United States, 2007-2013(1). Emerg Infect Dis. 2015;21(10):1794-1799.

47. Pritt BS, Allerdice MEJ, Sloan LM, et al. Proposal to reclassify Ehrlichia muris as Ehrlichia muris subsp. muris subsp. nov. and description of Ehrlichia muris subsp. eauclairensis subsp. nov., a newly recognized tick-borne pathogen of humans. Int J Syst Evol Microbiol. 2017;67(7):2121-2126.

48. Hermance ME, Thangamani S. Powassan virus: an emerging arbovirus of public health concern in North America. Vector Borne Zoonotic Dis. 2017;17(7):453- 462.

49. Ebel GD. Update on Powassan virus: emergence of a North American tick-borne flavivirus. Annu Rev Entomol. 2010;55:95-110.

50. Krause PJ. Human babesiosis. Int J Parasitol. 2019. doi:10.1016/j.ijpara.2018.11.007. Epub ahead of print.

51. Centers for Disease Control and Prevention. Babesiosis data & statistics. https://www.cdc.gov/parasites/babesiosis/data-statistics/index.html. Published December 2016. Updated July 2018. Accessed December 17, 2018.

52. Diuk-Wasser MA, Vannier E, Krause PJ. Coinfection by Ixodes tick-borne pathogens: ecological, epidemiological, and clinical consequences. Trends Parasitol. 2016;32(1):30-42.

53. Vannier EG, Diuk-Wasser MA, Ben Mamoun C, Krause PJ. Babesiosis. Infect Dis Clin North Am. 2015;29(2):357-370.

54. Dunham-Ems SM, Caimano MJ, Pal U, et al. Live imaging reveals a biphasic mode of dissemination of Borrelia burgdorferi within ticks. J Clin Invest. 2009;119(12):3652-3665.

55. De Silva AM, Fikrig E. Growth and migration of Borrelia burgdorferi in Ixodes ticks during blood feeding. Am J Trop Med Hyg. 1995;53(4):397-404.

56. Ribeiro JM, Mather TN, Piesman J, Spielman A. Dissemination and salivary delivery of Lyme disease spirochetes in vector ticks (Acari: Ixodidae). J Med Entomol. 1987;24(2):201-205.

57. Pal U, Yang X, Chen M, et al. OspC facilitates Borrelia burgdorferi invasion of Ixodes scapularis salivary glands. J Clin Invest. 2004;113(2):220-230.

58. Eisen L. Pathogen transmission in relation to duration of attachment by Ixodes scapularis ticks. Ticks Tick Borne Dis. 2018;9(3):535-542.

59. Ebel GD, Kramer LD. Short report: duration of tick attachment required for transmission of powassan virus by deer ticks. Am J Trop Med Hyg. 2004;71(3):268-271.

60. Buchon N, Silverman N, Cherry S. Immunity in Drosophila melanogaster--from microbial recognition to whole-organism physiology. Nat Rev Immunol. 2014;14(12):796-810.

61. Oliva Chavez AS, Shaw DK, Munderloh UG, Pedra JH. Tick humoral responses: marching to the beat of a different drummer. Front Microbiol. 2017;8:223.

62. Palmer WJ, Jiggins FM. Comparative genomics reveals the origins and diversity of arthropod immune systems. Mol Biol Evol. 2015;32(8):2111-2129.

63. Theopold U, Krautz R, Dushay MS. The Drosophila clotting system and its messages for mammals. Dev Comp Immunol. 2014;42(1):42-46.

64. Tsuzuki S, Matsumoto H, Furihata S, et al. Switching between humoral and cellular immune responses in Drosophila is guided by the cytokine GBP. Nat Commun. 2014;5:4628.

65. Gold KS, Bruckner K. Macrophages and cellular immunity in Drosophila melanogaster. Semin Immunol. 2015;27(6):357-368.

66. Jiravanichpaisal P, Lee BL, Soderhall K. Cell-mediated immunity in arthropods: hematopoiesis, coagulation, melanization and opsonization. Immunobiology. 2006;211(4):213-236.

67. Valanne S, Wang JH, Ramet M. The Drosophila Toll signaling pathway. J Immunol. 2011;186(2):649-656.

68. Michel T, Reichhart JM, Hoffmann JA, Royet J. Drosophila Toll is activated by Gram-positive bacteria through a circulating peptidoglycan recognition protein. Nature. 2001;414(6865):756-759.

69. Gobert V, Gottar M, Matskevich AA, et al. Dual activation of the Drosophila toll pathway by two pattern recognition receptors. Science. 2003;302(5653):2126- 2130.

70. Pili-Floury S, Leulier F, Takahashi K, et al. In vivo RNA interference analysis reveals an unexpected role for GNBP1 in the defense against Gram-positive bacterial infection in Drosophila adults. J Biol Chem. 2004;279(13):12848-12853.

71. Wang L, Weber AN, Atilano ML, Filipe SR, Gay NJ, Ligoxygakis P. Sensing of Gram-positive bacteria in Drosophila: GNBP1 is needed to process and present peptidoglycan to PGRP-SA. EMBO J. 2006;25(20):5005-5014.

72. Buchon N, Poidevin M, Kwon HM, et al. A single modular serine protease integrates signals from pattern-recognition receptors upstream of the Drosophila Toll pathway. Proc Natl Acad Sci U S A. 2009;106(30):12442-12447.

73. Gottar M, Gobert V, Matskevich AA, et al. Dual detection of fungal infections in Drosophila via recognition of glucans and sensing of virulence factors. Cell. 2006;127(7):1425-1437.

74. Lemaitre B, Kromer-Metzger E, Michaut L, et al. A recessive mutation, immune deficiency (imd), defines two distinct control pathways in the Drosophila host defense. Proc Natl Acad Sci U S A. 1995;92(21):9465-9469.

75. Kaneko T, Goldman WE, Mellroth P, et al. Monomeric and polymeric gram- negative peptidoglycan but not purified LPS stimulate the Drosophila IMD pathway. Immunity. 2004;20(5):637-649.

76. Leulier F, Parquet C, Pili-Floury S, et al. The Drosophila immune system detects bacteria through specific peptidoglycan recognition. Nat Immunol. 2003;4(5):478- 484.

77. Iatsenko I, Kondo S, Mengin-Lecreulx D, Lemaitre B. PGRP-SD, an extracellular pattern-recognition receptor, enhances peptidoglycan-mediated activation of the Drosophila Imd pathway. Immunity. 2016;45(5):1013-1023.

78. Paik D, Monahan A, Caffrey DR, Elling R, Goldman WE, Silverman N. SLC46 family transporters facilitate cytosolic innate immune recognition of monomeric peptidoglycans. J Immunol. 2017;199(1):263-270.

79. Myllymaki H, Valanne S, Ramet M. The Drosophila imd signaling pathway. J Immunol. 2014;192(8):3455-3462.

80. Georgel P, Naitza S, Kappler C, et al. Drosophila immune deficiency (IMD) is a death domain protein that activates antibacterial defense and can promote apoptosis. Dev Cell. 2001;1(4):503-514.

81. Leulier F, Vidal S, Saigo K, Ueda R, Lemaitre B. Inducible expression of double- stranded RNA reveals a role for dFADD in the regulation of the antibacterial response in Drosophila adults. Curr Biol. 2002;12(12):996-1000.

82. Chen P, Rodriguez A, Erskine R, Thach T, Abrams JM. Dredd, a novel effector of the apoptosis activators reaper, grim, and hid in Drosophila. Dev Biol. 1998;201(2):202-216.

83. Leulier F, Rodriguez A, Khush RS, Abrams JM, Lemaitre B. The Drosophila caspase Dredd is required to resist Gram-negative bacterial infection. EMBO Rep. 2000;1(4):353-358.

84. Meinander A, Runchel C, Tenev T, et al. Ubiquitylation of the initiator caspase DREDD is required for innate immune signalling. EMBO J. 2012;31(12):2770- 2783.

85. Paquette N, Broemer M, Aggarwal K, et al. Caspase-mediated cleavage, IAP binding, and ubiquitination: linking three mechanisms crucial for Drosophila NF- κB signaling. Mol Cell. 2010;37(2):172-182.

86. Lu Y, Wu LP, Anderson KV. The antibacterial arm of the Drosophila innate immune response requires an IκB kinase. Genes Dev. 2001;15(1):104-110.

87. Rutschmann S, Jung AC, Zhou R, Silverman N, Hoffmann JA, Ferrandon D. Role of Drosophila IKKγ in a toll-independent antibacterial immune response. Nat Immunol. 2000;1(4):342-347.

88. Silverman N, Zhou R, Stoven S, Pandey N, Hultmark D, Maniatis T. A Drosophila IκB kinase complex required for Relish cleavage and antibacterial immunity. Genes Dev. 2000;14(19):2461-2471.

89. Kleino A, Valanne S, Ulvila J, et al. Inhibitor of apoptosis 2 and TAK1-binding protein are components of the Drosophila Imd pathway. EMBO J. 2005;24(19):3423-3434.

90. Erturk-Hasdemir D, Broemer M, Leulier F, et al. Two roles for the Drosophila IKK complex in the activation of Relish and the induction of antimicrobial peptide genes. Proc Natl Acad Sci U S A. 2009;106(24):9779-9784.

91. Stoven S, Ando I, Kadalayil L, Engstrom Y, Hultmark D. Activation of the Drosophila NF-κB factor Relish by rapid endoproteolytic cleavage. EMBO Rep. 2000;1(4):347-352.

92. Stoven S, Silverman N, Junell A, et al. Caspase-mediated processing of the Drosophila NF-κB factor Relish. Proc Natl Acad Sci U S A. 2003;100(10):5991- 5996.

93. Kleino A, Myllymaki H, Kallio J, et al. Pirk is a negative regulator of the Drosophila Imd pathway. J Immunol. 2008;180(8):5413-5422.

94. Gulia-Nuss M, Nuss AB, Meyer JM, et al. Genomic insights into the Ixodes scapularis tick vector of Lyme disease. Nat Commun. 2016;7:10507.

95. Rosa RD, Capelli-Peixoto J, Mesquita RD, et al. Exploring the immune signalling pathway-related genes of the cattle tick Rhipicephalus microplus: from molecular characterization to transcriptional profile upon microbial challenge. Dev Comp Immunol. 2016;59:1-14.

96. Bechsgaard J, Vanthournout B, Funch P, et al. Comparative genomic study of arachnid immune systems indicates loss of β-1,3-glucanase-related proteins and the immune deficiency pathway. J Evol Biol. 2016;29(2):277-291.

97. Shaw DK, Wang X, Brown LJ, et al. Infection-derived lipids elicit an immune deficiency circuit in arthropods. Nat Commun. 2017;8:14401.

98. Smith AA, Pal U. Immunity-related genes in Ixodes scapularis--perspectives from genome information. Front Cell Infect Microbiol. 2014;4:116.

99. Severo MS, Choy A, Stephens KD, et al. The E3 ubiquitin ligase XIAP restricts Anaplasma phagocytophilum colonization of Ixodes scapularis ticks. J Infect Dis. 2013;208(11):1830-1840.

100. Zhang S, Guha S, Volkert FC. The Saccharomyces SHP1 gene, which encodes a regulator of phosphoprotein phosphatase 1 with differential effects on glycogen metabolism, meiotic differentiation, and mitotic cell cycle progression. Mol Cell Biol. 1995;15(4):2037-2050.

101. Schuberth C, Richly H, Rumpf S, Buchberger A. Shp1 and Ubx2 are adaptors of Cdc48 involved in ubiquitin-dependent protein degradation. EMBO Rep. 2004;5(8):818-824.

102. Cheng YL, Chen RH. The AAA-ATPase Cdc48 and cofactor Shp1 promote chromosome bi-orientation by balancing Aurora B activity. J Cell Sci. 2010;123(Pt 12):2025-2034.

103. Sang TK, Ready DF. Eyes closed, a Drosophila p47 homolog, is essential for photoreceptor morphogenesis. Development. 2002;129(1):143-154.

104. Kondo H, Rabouille C, Newman R, et al. p47 is a cofactor for p97-mediated membrane fusion. Nature. 1997;388(6637):75-78.

105. Yuan X, Simpson P, McKeown C, et al. Structure, dynamics and interactions of p47, a major adaptor of the AAA ATPase, p97. EMBO J. 2004;23(7):1463-1473.

106. Dreveny I, Kondo H, Uchiyama K, Shaw A, Zhang X, Freemont PS. Structural basis of the interaction between the AAA ATPase p97/VCP and its adaptor protein p47. EMBO J. 2004;23(5):1030-1039.

107. Uchiyama K, Jokitalo E, Lindman M, et al. The localization and phosphorylation of p47 are important for Golgi disassembly-assembly during the cell cycle. J Cell Biol. 2003;161(6):1067-1079.

108. Wang Y, Satoh A, Warren G, Meyer HH. VCIP135 acts as a deubiquitinating enzyme during p97-p47-mediated reassembly of mitotic Golgi fragments. J Cell Biol. 2004;164(7):973-978.

109. Krick R, Bremer S, Welter E, et al. Cdc48/p97 and Shp1/p47 regulate autophagosome biogenesis in concert with ubiquitin-like Atg8. J Cell Biol. 2010;190(6):965-973.

110. Shibata Y, Oyama M, Kozuka-Hata H, et al. p47 negatively regulates IKK activation by inducing the lysosomal degradation of polyubiquitinated NEMO. Nat Commun. 2012;3:1061.

111. McClure Carroll EE, Wang X, Shaw DK, et al. p47 licenses activation of the immune deficiency pathway in the tick Ixodes scapularis. Proc Natl Acad Sci U S A. 2019;116(1):205-210.

112. Carlyon JA. Laboratory Maintenance of Anaplasma phagocytophilum. Curr Protoc Microbiol. 2005;Chapter 3:Unit 3A 2.

113. Yoshiie K, Kim HY, Mott J, Rikihisa Y. Intracellular infection by the human granulocytic ehrlichiosis agent inhibits human neutrophil apoptosis. Infect Immun. 2000;68(3):1125-1133.

114. Labandeira-Rey M, Skare JT. Decreased infectivity in Borrelia burgdorferi strain B31 is associated with loss of linear plasmid 25 or 28-1. Infect Immun. 2001;69(1):446-455.

115. Zuckert WR. Laboratory maintenance of Borrelia burgdorferi. Curr Protoc Microbiol. 2007;Chapter 12:Unit 12C 11.

116. Munderloh UG, Liu Y, Wang M, Chen C, Kurtti TJ. Establishment, maintenance and description of cell lines from the tick Ixodes scapularis. J Parasitol. 1994;80(4):533-543.

117. Samakovlis C, Asling B, Boman HG, Gateff E, Hultmark D. In vitro induction of cecropin genes--an immune response in a Drosophila blood cell line. Biochem Biophys Res Commun. 1992;188(3):1169-1175.

118. de la Fuente J, Antunes S, Bonnet S, et al. Tick-pathogen interactions and vector competence: identification of molecular drivers for tick-borne diseases. Front Cell Infect Microbiol. 2017;7:114.

119. Radivojac P, Vacic V, Haynes C, et al. Identification, analysis, and prediction of protein ubiquitination sites. Proteins. 2010;78(2):365-380.

120. Pierce NW, Kleiger G, Shan SO, Deshaies RJ. Detection of sequential polyubiquitylation on a millisecond timescale. Nature. 2009;462(7273):615-619.

121. Mark KG, Loveless TB, Toczyski DP. Isolation of ubiquitinated substrates by tandem affinity purification of E3 ligase-polyubiquitin-binding domain fusions (ligase traps). Nat Protoc. 2016;11(2):291-301.

122. Meyer HH, Wang Y, Warren G. Direct binding of ubiquitin conjugates by the mammalian p97 adaptor complexes, p47 and Ufd1-Npl4. EMBO J. 2002;21(21):5645-5652.

123. Yuan X, Shaw A, Zhang X, et al. Solution structure and interaction surface of the C-terminal domain from p47: a major p97-cofactor involved in SNARE disassembly. J Mol Biol. 2001;311(2):255-263.

124. Soukenik M, Diehl A, Leidert M, et al. The SEP domain of p47 acts as a reversible competitive inhibitor of cathepsin L. FEBS Lett. 2004;576(3):358-362.

125. Zhang Q, Lenardo MJ, Baltimore D. 30 years of NF-κB: a blossoming of relevance to human pathobiology. Cell. 2017;168(1-2):37-57.

126. Li J, Chai QY, Liu CH. The ubiquitin system: a critical regulator of innate immunity and pathogen-host interactions. Cell Mol Immunol. 2016;13(5):560- 576.

127. Komander D, Rape M. The ubiquitin code. Annu Rev Biochem. 2012;81:203-229.

128. Choy A, Severo MS, Sun R, Girke T, Gillespie JJ, Pedra JH. Decoding the ubiquitin-mediated pathway of arthropod disease vectors. PLOS ONE. 2013;8(10):e78077.

129. Metzger MB, Pruneda JN, Klevit RE, Weissman AM. RING-type E3 ligases: master manipulators of E2 ubiquitin-conjugating enzymes and ubiquitination. Biochim Biophys Acta. 2014;1843(1):47-60.

130. Severo MS, Sakhon OS, Choy A, Stephens KD, Pedra JH. The 'ubiquitous' reality of vector immunology. Cell Microbiol. 2013;15(7):1070-1078.

131. Schmukle AC, Walczak H. No one can whistle a symphony alone - how different ubiquitin linkages cooperate to orchestrate NF-κB activity. J Cell Sci. 2012;125(Pt 3):549-559.

132. Chen L, Paquette N, Mamoor S, et al. Innate immune signaling in Drosophila is regulated by transforming growth factor β (TGFβ)-activated kinase (Tak1)- triggered ubiquitin editing. J Biol Chem. 2017;292(21):8738-8749.

133. Xu G, Jaffrey SR. Proteomic identification of protein ubiquitination events. Biotechnol Genet Eng Rev. 2013;29:73-109.

134. Meyer JM, Hill CA. Tick genetics, genomics, and transformation. In: Sonenshine DE, Roe RM, eds. Biology of Ticks. Vol 2. 2nd ed. New York, NY: Oxford University Press; 2014:61-87.

135. de la Fuente J, Kocan KM, Almazan C, Blouin EF. RNA interference for the study and genetic manipulation of ticks. Trends Parasitol. 2007;23(9):427-433.

136. Kocan KM, Blouin E, de la Fuente J. RNA interference in ticks. J Vis Exp. 2011(47).

137. Narasimhan S, Montgomery RR, DePonte K, et al. Disruption of Ixodes scapularis anticoagulation by using RNA interference. Proc Natl Acad Sci U S A. 2004;101(5):1141-1146.

138. Aalto AL, Mohan AK, Schwintzer L, et al. M1-linked ubiquitination by LUBEL is required for inflammatory responses to oral infection in Drosophila. Cell Death Differ. 2018. Epub ahead of print. doi: 10.1038/s41418-018-0164-x

139. Aggarwal K, Rus F, Vriesema-Magnuson C, Erturk-Hasdemir D, Paquette N, Silverman N. Rudra interrupts receptor signaling complexes to negatively regulate the IMD pathway. PLoS Pathog. 2008;4(8):e1000120.

140. Lhocine N, Ribeiro PS, Buchon N, et al. PIMS modulates immune tolerance by negatively regulating Drosophila innate immune signaling. Cell Host Microbe. 2008;4(2):147-158.

141. Zaidman-Remy A, Herve M, Poidevin M, et al. The Drosophila amidase PGRP- LB modulates the immune response to bacterial infection. Immunity. 2006;24(4):463-473.

142. Kleino A, Ramia NF, Bozkurt G, et al. Peptidoglycan-sensing receptors trigger the formation of functional amyloids of the adaptor protein Imd to initiate Drosophila NF-κB Signaling. Immunity. 2017;47(4):635-647.e6.

143. Pelka K, De Nardo D. Emerging concepts in innate immunity. Methods Mol Biol. 2018;1714:1-18.

144. Gay NJ, Gangloff M, O'Neill LA. What the Myddosome structure tells us about the initiation of innate immunity. Trends Immunol. 2011;32(3):104-109.

145. Xu G, Paige JS, Jaffrey SR. Global analysis of lysine ubiquitination by ubiquitin remnant immunoaffinity profiling. Nat Biotechnol. 2010;28(8):868-873.

146. Rota-Stabelli O, Daley AC, Pisani D. Molecular timetrees reveal a Cambrian colonization of land and a new scenario for ecdysozoan evolution. Curr Biol. 2013;23(5):392-398.

147. Dunning Hotopp JC, Lin M, Madupu R, et al. Comparative genomics of emerging human ehrlichiosis agents. PLoS Genetics. 2006;2:e21.

148. Goto A, Okado K, Martins N, et al. The kinase IKKβ regulates a STING- and NF- κB-dependent antiviral response pathway in Drosophila. Immunity. 2018;49(2):225-234.e4.

149. Martin M, Hiroyasu A, Guzman RM, Roberts SA, Goodman AG. Analysis of Drosophila STING reveals an evolutionarily conserved antimicrobial function. Cell Rep. 2018;23(12):3537-3550.e6.

150. Liu Y, Gordesky-Gold B, Leney-Greene M, Weinbren NL, Tudor M, Cherry S. Inflammation-induced, STING-dependent autophagy restricts Zika virus infection in the Drosophila brain. Cell Host Microbe. 2018;24(1):57-68.e3.

151. Costa A, Jan E, Sarnow P, Schneider D. The Imd pathway is involved in antiviral immune responses in Drosophila. PLoS One. 2009;4(10):e7436. 152. Stach L, Freemont PS. The AAA+ ATPase p97, a cellular multitool. Biochem J. 2017;474(17):2953-2976.

153. Ye Y, Tang WK, Zhang T, Xia D. A mighty "protein extractor" of the cell: structure and function of the p97/CDC48 ATPase. Front Mol Biosci. 2017;4:39.

154. Cadwell K. Crosstalk between autophagy and inflammatory signalling pathways: balancing defence and homeostasis. Nat Rev Immunol. 2016;16(11):661-675.