Immunomodulatory Effects of the Short Chain Fatty Acid Butyrate On

IMMUNOMODULATORY EFFECTS OF THE SHORT CHAIN FATTY ACID BUTYRATE ON

GUT MUCOSAL T CELLS IN THE SETTING OF HIV-1 INFECTION

JON J. KIBBIE

B.S., Loyola Marymount University, 2008

M.S. University of California Los Angeles, 2013

A thesis submitted to the

Faculty of the Graduate School of the

University of Colorado in partial fulfillment

of the requirements for the degree of

Doctor of Philosophy

Immunology Program

2019

The thesis for the Doctor of Philosophy degree by Jon J Kibbie Has been approved for the Immunology Program by

Peter Henson, Chair Arthur Gutierrez-Hartman Laurel Lenz Sean Colgan Thomas Campbell Cara C. Wilson. Advisor

Date: 05/17/2019

Kibbie, Jon Jason (Ph.D., Immunology) Immunomodulatory Effects of the Short Chain Fatty Acid Butyrate on Gut Mucosal T Cells in the Setting of HIV-1 Infection Thesis Directed by Professor Cara C. Wilson ABSTRACT The gastrointestinal tract is profoundly affected during acute and chronic HIV-1 infection. Early in the course of the infection, there is a breakdown in gut epithelial barrier integrity, high levels of viral replication, CD4 T cell depletion and inflammation in the gut mucosa. Importantly, People Living with HIV-1 (PLWH) lose significant numbers of protective T helper (Th) 17 and Th 22 populations in the gut. These populations of Th cells are essential in promoting epithelial barrier integrity and appropriate immune responses in the mucosa. The combined loss of protective Th cell populations and an increase in intestinal permeability contributes to the translocation of bacteria and bacterial products from the gut lumen into the lamina propria. Microbial translocation (MT) has been linked to mucosal and systemic inflammation and is predictive of disease progression and mortality in

PLWH. The gut microbiome has been shown to influence mucosal immune responses.

Therefore, its role in HIV-associated mucosal pathogenesis has been a high priority research focus. To address the role of the microbiome in HIV-1 associated gut pathogenesis and MT, several groups have sequenced the intestinal microbiomes of PLWH. Many of these studies have demonstrated a dysbiosis of the microbiome that associates with systemic and mucosal inflammation. A key feature of this dysbiosis is the lower relative abundance of bacteria that produce the regulatory short-chain fatty acid butyrate. Loss of these butyrate-producing bacteria (BPB) species may contribute to the breakdown in gut mucosal immune cell homeostasis and systemic inflammation providing a link between changes in the microbiome and the host immune response. The mechanisms by which low

iii abundances of butyrate-producing bacteria and likely lower tissue butyrate levels contribute to gut mucosal CD4 T cell activation and HIV-1 infection have yet to be determined.

The purpose of the studies outlined in this thesis was to investigate the importance of the findings of decreased BPB species during HIV-1 infection and likely lower tissue butyrate levels in an in vitro human gut lamina propria mononuclear cell (LPMC) model of T cell activation and infection. This model was used to understand the impact of butyrate on enteric bacteria driven T cell activation and HIV-1 infection. These studies demonstrated that butyrate at higher physiologic concentrations decreased enteric bacteria driven LP T cell activation, cytokine production, and LP CD4 T cell productive infection. Next, to address the mechanism by which butyrate alters LP CD4 T cell activation and infection, I used an in vitro human LPMC model of direct LP CD4 T cell activation and HIV-1 infection. These studies determined that butyrate decreased LP CD4 T cell activation, proliferation, and cytokine production in a concentration-dependent manner likely through histone deacetylase inhibition. In these studies, gut CD4 T cell productive HIV-1 infection in the setting of CD4 T cell activation was decreased at higher concentrations but increased at lower concentrations of butyrate likely through increased CCR5 expression and increased viral entry/reverse transcription. The findings outlined in this study suggest that the low abundance of mucosal butyrate-producing bacteria and the likely low tissue butyrate levels that occur in the setting of chronic HIV-1 infection may potentiate LP CD4 T cell infection, contribute to chronic immune activation and therefore might be a contributing factor in HIV-1 disease pathogenesis.

The form and content of the abstract are approved. I recommend its publication.

Approved: Cara C. Wilson

I dedicate this thesis to my wife, Kathleen Kibbie, who has been a steadfast supporter during my academic pursuits. Without her support, my achievements and progress would not have been possible. To my daughters Penny and Ellie, you have both brought me more joy and happiness than I could have ever asked.

Lastly, to my parents whose continual support has allowed me the

privilege of pursuing this career path.

ACKNOWLEDGMENTS I would first like to thank my mentor, Dr. Cara Wilson, for her guidance and support during my graduate career. Cara is a great role model of a physician-scientist, and I am grateful for her advice in pursuing a career both in the sciences and medicine and how to balance that career with my family life. During my Ph.D. training, my wife and I had two children, and I am grateful for the support and understanding Cara extended to me during this time. Next, I would like to thank Dr. Steph Dillon for her insight and direct help in developing this project. Without her studies addressing the relative abundance of butyrate- producing bacteria in the colonic mucosa of PLWH, this project would not have had the basis to proceed. Also, her countless hours of mentorship and guidance with this project as well as with my writing have been essential for my progression during my graduate career.

In addition, I would like to thank the Wilson and Santiago lab members past and present who have been a continued source of support and guidance. From the Wilson lab, I would specifically like to thank Dr. Moriah Castleman, Dr. Jay Liu, Christine Purba, and

Andrew Donovan. They provided intellectual insight, helped with experiments and gave the support and friendship that is essential during graduate training. From the Santiago Lab, I would specifically like to thank Dr. Mario Santiago, Dr. Kejun Guo, Brad Barret and Sean

Jones who offered technical help, intellectual support, and friendship.

Next, I would like to thank my Thesis Committee, Dr. Peter Henson (Chair), Dr.

Laurel Lenz, Dr. Arthur Gutierrez-Hartman, Dr. Sean Colgan, Dr. Jose Castillo-Mancilla and

Dr. Thomas Campbell. Their guidance and advice through my graduate career have been invaluable, and I am thankful for the time they took to help me develop the skills necessary for a career in science. I would also like to thank Dr. Jose Castillo-Mancilla for allowing me the opportunity to shadow him during his ID clinic at the University of Colorado Hospital. He was not only open to training me clinically but giving me candid advice on how to have a career as a physician-scientist while balancing my roles outside of work. I would also like to

vi thank both the Immunology Graduate program and the MSTP for their support and guidance during this time. Furthermore, I am grateful for the program administrators past and present, notably Elizabeth Bowen and Michele Parsons, who have helped guide me through the requirements needed to complete my graduate training.

Lastly, I would like to thank the sources of funding that supported this project. During my time in the graduate program, I was supported by Cara Wilson’s funding source R01

AI108404 and the CCTSI TL1 fellowship TR001082.

vii

TABLE OF CONTENTS CHAPTER

I. INTRODUCTION

HIV-1 Infection: Epidemiology…………………………………………………….1

HIV-1 Infection: The Virus, Life Cycle, and ART……………………………..…2

HIV-1 Infection: Pathogenesis…………………………………………………….4

HIV-1 Infection: Mucosal Immune Response……………………………………7

Mucosal Immunity in Health………………………………………………7

Mucosal Immunity During HIV-1 Infection………………….………….10

The Role of the Gut Microbiome in HIV-1 Infection……...…………………….13

Short Chain Fatty Acids………………………………………………….……….16

SCFA Mechanisms of Action…………………………………………………….19

Histone Deacetylase Inhibition………………………………………….19

G-protein Coupled Receptor and PPARγ Signaling……………….….20

Butyrate and Immune Cell Interactions…………………………………...…….21

Project Rationale: Alterations in Butyrate Producing Bacteria in PLWH...... 26

Scope of Thesis……………………………………………………………………26

II. MATERIALS AND METHODS

Human Intestinal LPMC Isolation…………….…………………………….……30

PBMC Isolation………………………………………….…………………………30

Growth of Bacteria Stocks…………………………………………………….….31

Preparation of CCR5-tropic CH40 Transmitted Founder Stocks and HIV-1Bal Stocks………………………………………………………………………………31

LPMC and PBMC Culture Model………………………………………….….….31

SCFA Assay Conditions………..…………………………………………..…….32

Small Molecule PPARγ, HATi, and HDACi Assays……………….………...…32

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Quantification of Secreted Cytokine Production………………….……………33

FLOW Acquisition and Analysis …………………………………………….….33

Measurement of LP CD4 T Cell Activation…………………………….……….33

Measurement of LP CD4 T Cell Proliferation………………………….……….34

Measurement of LP CD4 T Regulatory Cell Frequency………………………34

LP and PB CD4 T Helper Cell Proliferation ……………………………………34

Studies to Understand Butyrate’s Mechanism of Action

Quantification of Histone H3 Acetylation ………………………..…….35

LP CD4 T Cell GPR Expression ………………………..…………...…39

Flow-cytometry Based Measurement of Productive HIV-1 Infection…...……39

HIV-1 Infection Related Assays

HIV-1 Co-receptor Expression Analysis………….………………….…40

TZM-bl Assay………………………………………………………...…...40

HIV-1 RU-5 Quantification………………………………………...….….40

HIV-1 gag Quantification………………………………………..…….…41

Statistical Analysis……………………………………………………...…………42

III. EXOGENOUS BUTYRATE DECREASED ENTERIC PATHOBIONT DRIVEN LP T CELL ACTIVATION AND HIV-1 INFECTION

Preface…………………………….…………………………………………….…46

Background………………………………………………………………………...46

Results

Butyrate Reduces P. stercorea-driven Activation of LP T Cells in vitro………………….…………………………………………….……….48

Butyrate Reduces P. stercorea-driven Activation of HIV-1 Infected LP T Cells in vitro………..…………………………………….………….….51

Butyrate Decreases LPMC Production of IFNγ and IL-17 in Response to P. stercorea…………………………………………………………….51

Butyrate Inhibits P. stercorea Driven HIV-1 Infection in LP CD4+ T cells…………………...……………………………………………………51

Discussion………………………………………………………………………….52

IV. BUTYRATE DIFFERENTIALLY IMPACTS LP CD4 T CELL ACTIVATION AND INFECTION IN VITRO IN A CONCENTRATION DEPENDENT MANNER

Background………………………………………………………………...………54

Results

In vitro Exposure to Exogenous Butyrate Decreases TCR-mediated LP CD4 T Cell Activation and Proliferation Directly in a Concentration- Dependent Manner…………………………………………………….…56

Timing of Butyrate Exposure and TCR Stimulation is Important for the Inhibition of LP CD4 T Cell Proliferation……………………….….…...57

Higher Concentrations of Propionate and Acetate Relative to Butyrate were Required to Similarly Inhibit LP CD4 T Cell Activation and Proliferation………………………………….…………………….………59

Butyrate’s Inhibition of LP CD4 T Cell Activation and Proliferation is likely Mediated by HDAC Inhibition………………………….………….59

Known Butyrate Receptors, GPR109a and GPR43, are Minimally Expressed on LP CD4 T Cells…………………………………………..68

LP Th17 cell proliferation is decreased at Lower Concentrations of Butyrate Relative to other Th Subsets……….……………….…….….68

Butyrate Enhances HIV-1 Infection of LP CD4 T Cells at Lower Concentrations…………………………………………………………....70

Low Dose Butyrate Increases the Expression of CCR5 on Activated LP CD4 T Cells and HIV-1 Viral Entry………………………………….77

Apicidin Decreases LP CD4 T Cell Productive Infection and CD4 Expression………...………………………………………………………78

Discussion…………………………………………...…………………………….82

V. ADDITIONAL MECHANISMS OF BUTYRATE’S IMMUNOMODULATORY EFFECTS ON LP CD4 T CELL ACTIVATION AND PROLIFERATION

Background……………………………………………………………………...…86

Results

Inhibition of Histone Acetyltransferases does not Block the Anti- proliferative Effect of Butyrate on TCR Stimulated LP CD4 T Cells……………………………………………………………………...... 87

PPARγ Agonism Phenocopies the Anti-proliferative Effect of Butyrate but is not Reversed by a PPARγ Antagonist………………………..…89

Discussion……………………………………………………………………….....89

VI. DISCUSSION

Project Rationale and Outline…………………………………………………....92

Summary of Major Findings……………………………………………………...93

Potential Implications………………………………………………………..…....99

Limitations and Future Directions…………………………………………...…103

VII. REFERENCES……………………………………………………………….………..….106

VIII. APPENDIX A……………………………………………………………………………...127

LIST OF TABLES

Table

1.1 Butyrate Producing Bacteria and Pathways Utilized………………………………...…18

2.1 Flow Antibodies…………………………………………………………………………….45

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LIST OF FIGURES

FIGURES

1.1 HIV-1 Viral Life Cycle and the Sites of Action of ART…………………………..…….…6

1.2 The Gut Mucosal Immune Response During HIV-1 Infection………………….…..….14

1.3 SCFA Signaling Pathways in Cells……………………………………………………….25

1.4 Communities of Butyrate-producing Bacteria (BPB) are Reduced in Colon Mucosa of Untreated HIV-1 Infected Study Participants…………………………………….…..….29

2.1 Flow Gating Scheme for Stimulated LP CD4 T Cell Activation and Proliferation Studies….…………………………………………………………………………….….….36

2.2 Flow Gating Scheme for the Identification of T Helper Subsets………….…………...37

2.3 Butyrate Increases LP CD4 T Cell Histone Acetylation Levels (acH3K9)……….….38

2.4 Flow Gating Scheme for the Identification of Productive Infection of LP CD4 T Cells…………………………………………………………………………………………43

2.5 Flow Gating Scheme for the Quantification of CCR5 Expression on LP CD4 T Cells………………………………………………………………………………………….44

3.1 Butyrate Decreases P. stercorea Driven Lamina Propria CD4 and CD8 T Cell Activation and Cytokine Production in the Absence of HIV-1 Infection in vitro……………………………………………………………………………………….….49

3.2 Butyrate Reduces P. stercorea-enhanced LP T Cell Activation and HIV-1 Productive Infection….………………………………………………………………………………….50

4.1 Total LPMC Viability After Exposure to Butyrate………………………………….……58

4.2 Butyrate Reduces LP T Cell Activation and Proliferation in a Dose-Dependent Manner………………………………………………………………….60

4.3 Examination of FOXP3+ CD25+ LP CD4 T Cells After the Addition of Butyrate……………………………………………………………………………………..62

4.4 Propionate and Acetate both Decrease TCR Bead Driven LP CD4 T Cell Activation and Proliferation…………………………………………………………………….……...63

4.5 Butyrate’s HDACi Activity Likely Plays a Role in its Immunomodulatory Effect on Activated LP CD4 T Cells……………………………………………………………...….65

4.6 Butyrate Increases LP CD4 T Cell Histone Acetylation Levels (acH3K9)……..…….67

4.7 LP CD4 T Cells Express Low Levels of GPR43 and GPR109a……………………....69

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4.8 Butyrate Decreases T Helper Cytokine Production and the Proliferation of Gut T Helper Cells………….…………………………………………………………………….71

4.9 Butyrate Reduces PB T Cell Activation and Proliferation in a Dose-Dependent Manner………………………………………………………….…………………………..72

4.10 High and Low Concentrations of Butyrate Differentially Alter LP CD4 T Cell HIV-1 Infection Levels……………………………………………….……………….…….…..…74

4.11 Butyrate does not Alter the Frequencies of Productively Infected T Helper Subsets in the Absence of Activation……………………………………………………….……...…76

4.12 Butyrate Alters HIV-1 Co-receptor Expression on LP CD4 T Cells and Viral Entry…………………………………………………………………………………..….....79

4.13 Apicidin Decreases LP CD4 T Cell Productive Infection and CD4 Expression at High Concentrations…………………………………………………………………………..…81

5.1 HAT Inhibition Decreases Butyrate Driven H3K9 Acetylation but does not Impact Butyrate’s Effect on LP CD4 T Cell Proliferation…………………………….…….…...88

5.2 PPARγ Agonism Decreases LP CD4 T Cell Proliferation, but PPARγ Antagonism does not Inhibit this Effect………………………………………………………………...90

6.1 LP CD4 T cell Activation and HIV-1 Infection is Differentially Impacted by Butyrate in a Concentration-Dependent Manner……………………………………………...... …100

xiv

LIST OF ABBREVIATIONS

HIV-1 Human immunodeficiency virus 1 CD Cluster of differentiation

AIDS Acquired immunodeficiency syndrome TLR Toll-like receptor

SIV Simian immunodeficiency virus GPCR(GPR) G protein-coupled receptor

LPMC Lamina propria mononuclear cell WHO World Health Organization

PBMC Peripheral blood mononuclear cell MSM Men who have sex with men

BPB Butyrate-producing bacteria PLWH People living with HIV-1

MT Microbial translocation ART Anti-retroviral therapy

ELISA enzyme-linked immunosorbent assay PreP Pre-exposure prophylaxis

Th T helper Gag group specific antigen

RA Relative abundance Pol Polymerase

IFN Interferon Env envelope glycoprotein

DNA deoxyribonucleic acid cDNA complementary DNA

RNA ribonucleic acid mRNA messenger RNA

RT Reverse transcriptase NRTI Nucleoside reverse transcriptase inhibitor HDAC Histone deacetylase NNRTI Non-nucleoside reverse transcriptase inhibitor HAT Histone acetyltransferase INSTI Integrase strand transfer inhibitors HDACi Histone deacetylase inhibitor LTR Long terminal repeats

HATi Histone acetyltransferase inhibitor ssRNA Single-stranded RNA

PPARγ Peroxisome proliferator-activated DC Dendritic Cell receptor gamma

IL Interleukin APC Antigen presenting cell

LPS Lipopolysaccharide ILC Innate lymphoid cell

LTA Lipoteichoic acid GI Gastrointestinal

16SrRNA 16S ribosomal RNA IEL Intraepithelial lymphocytes

CFSE Carboxyfluorescein succinimidyl ester MAIT Mucosal associated invariant T cell

CHAPTER I INTRODUCTION

HIV-1 Infection: Epidemiology

The retrovirus human immunodeficiency virus type 1 (HIV-1) that causes acquired immunodeficiency syndrome (AIDS) has been a global health epidemic for over 30 years and continues to be one of the primary causes of mortality and morbidity worldwide[1].

According to the World Health Organization’s (WHO) most recent data, as of 2017, there have been more than 70 million known infections and at least 35 million individuals who have died due to HIV-1 related illnesses. HIV-1 prevalence is higher in several groups such as intravenous drug users, men who have sex with men (MSM), sex workers and those with barriers to care such as those who are incarcerated and that lack access to medical care.

While there are significant global populations of persons living with HIV-1 (PLWH), there is a wide prevalence of HIV-1 infection in many of the countries in western, central, eastern and southern Africa where the highest burdens of HIV-1 infection exist[1].

The WHO notes that there are currently 36.9 million PLWH and nearly 1 million individuals succumbed to HIV-1 related illnesses in 2017, and many of those individuals are in developing countries, where there are barriers to the access to adequate care and medical education[1]. Notably, the WHO has stated nearly 4% of adults in Africa are infected with HIV-1, which represents 2/3 of total global HIV-1 infections. The introduction of antiretroviral therapy (ART) has resulted in a dramatic decline in the incidence of HIV-1 related illnesses and deaths[1, 2] through the decrease in HIV-1 viral load[3] and thus CD4

T cell loss[3, 4]. Adherence to ART has resulted in an increased lifespan of PLWH and decreased the incidence of new infections as individuals who are virally suppressed are unlikely to infect others[5, 6]. Recently, ART based pre-exposure prophylaxis (PreP), has been approved for individuals at risk of HIV-1 infection and has been shown to decrease

HIV-1 transmission in adherent individualseffectively[7-9]. It is clear from the continued prevalence of HIV-1 that there is a need for additional treatments and better access to adequate medical care and education to quell the HIV-1 epidemic[1, 7].

HIV-1 Infection: The Virus, Life Cycle, and ART

The HIV-1 virus belongs to the genus Lentivirus, family Retroviridae and subfamily

Orthoretrovirinae[8]. Within each mature viral particle, there are two identical single strands of RNA (ssRNA) that are contained within a viral core protein[8, 9]. The viral genome is flanked by long terminal repeat sequences (LTR) that contain the essential promoter regions for viral transcription. The three primary structural genes are gag (group-specific antigen), pol (polymerase) and env (envelope glycoprotein). The proteins encoded by gag consist of the outer core membrane (p17), the capsid protein (p24) and the nucleocapsid (p7) while pol codes for the enzymes protease (p12), reverse transcriptase (p51), RNase H (p15) and integrase (p32). Lastly, env codes for the two envelope glycoproteins gp120 and gp41[8-10].

The HIV-1 genome also codes for several accessory proteins that include Tat and Rev, which help in the initiation of HIV-1 replication from integrated viral DNA and export of viral transcripts from the nucleous[8-10]. There are also several regulatory proteins that help the virus evade host immune responses, which include Nef, Vif, Vpr and Vpu [8, 9].

The HIV-1 viral life cycle begins with the high-affinity binding of HIV-1 envelope protein gp120 to CD4 on the cell surface[11, 12]. This interaction is necessary but not sufficient for viral entry, which requires subsequent contact of gp120 with a co-receptor. HIV-

1 co-receptors are the G-protein coupled chemokine receptors CCR5 and CXCR4[13-16].

Viral tropism predicts which co-receptors gp120 interacts with during infection. Initially, most of the transmitted viruses have tropism for CCR5 while later in advanced disease there is a shift to CXCR4 tropism[16-19]. The binding of HIV-1 gp120 to CD4 and the co-receptor elicits a conformational change in the viral envelope proteins driving fusion of the virus and target cell[20, 21]. Once inside the cell endosome, the viral capsid contents are released

2 into the cytoplasm[8, 10, 22]. The viral protein reverse transcriptase (RT) then transcribes the single-stranded HIV-1 RNA into complementary DNA (cDNA). During the early steps of reverse transcription, two template switches occur. These involve the transfer of both minus and plus strand strong stop DNA (RU5 sequence) facilitating the production of viral DNA[8,

22, 23]. While cDNA is being synthesized, the viral RNA is degraded by the viral enzyme

RNase H. Next, the single-stranded cDNA is converted into double-stranded DNA by reverse transcriptase’s DNA dependent DNA polymerase activity. HIV-1 DNA is then transported into the cell nucleus by the viral protein integrase which also acts to insert the viral DNA throughout the human host cell genome[10, 22, 24]. Once the viral DNA has undergone integration, the cell can either remain latently infected or the viral genome can be transcribed during cell division or activation. During transcription, the long terminal repeat

(LTR) promoter of the viral genome serves as a docking site for host cellular DNA dependent RNA polymerases along with the viral protein tat and other transcription factors that drive the synthesis of viral mRNA. After splicing of the viral RNA and translation, viral proteins along with the Gag-Pol precursor polypeptides are packed into nascent virions and bud out of the cell into the extracellular space through the host cell membrane[10, 22, 24].

Once this process is complete, the viral protein protease cleaves the Gag-Pol peptides forming the mature infectious virion[10, 22, 24].

Over the last 20 years, there have been several pharmacological advances in the treatment of HIV-1 infection. Most important of these are ART advances that target essential components of the HIV-1 life cycle. The first approved drug for the treatment of HIV-1 was a nucleoside reverse transcriptase inhibitor (NRTI), zidovudine[2, 25], which functions by targeting RT and interrupting the elongation of the viral DNA chain produced by RT[2, 26].

This drug, along with other nucleoside or nucleotide reverse transcription inhibitors, is a pro- drug that is phosphorylated by cellular enzymes once in the cell to become the active drug.

They are taken up by RT due to their similarity to cellular nucleotides but stop the elongation

3 of the DNA chain due to competitive inhibition of cellular nucleotides in a process called chain termination[2, 26]. Non-nucleoside reverse transcriptase inhibitors (NNRTIs) are another class of ART that target RT but do so by directly altering RT protein conformation and inhibiting its function[2, 27]. These drugs bind to RT outside of the active site and function through non-competitive inhibition. The next class of ART target the viral protease, which prevents the maturation and production of HIV-1 virions by interrupting the function of the viral protease protein[2, 28]. The integration of the HIV-1 genome into the host cell can also be disrupted by targeting viral integrase with integrase strand transfer inhibitors

(INSTIs) such as raltegravir[29]. This class of drugs blocks integrase from catalyzing the formation of bonds between the host and viral DNA, thus preventing the incorporation of viral DNA into the human genome[2, 29]. Lastly, there are drugs, termed entry inhibitors, that have been developed to block the interaction of HIV-1 with CCR5 (maraviroc)[30].

Furthermore, there is a class of fusion inhibitors that prevent the fusion of the virus with the target cell, such as Enfuvirtide[2, 31]. While these drugs are very effective at halting HIV-1 infection, they must be used in combination due to the ability of the virus to rapidly mutate and become resistant to individual drugs. Therefore most ART combinations in use have a combination of NRTI/NNRTI and another ART class or 2NRTI+NNRTI[2].

HIV-1 Infection: Pathogenesis

HIV-1 is transmitted through breaches in the mucosal or epidermal barriers, intravenous introduction or vertically from mother to child either in utero, during delivery or breast feeding[22]. During the initial transmission event, it is thought that single virions are responsible for establishing an infection. The virus that transmits the infection is known as a transmitted founder virus (TF), which through variance in glycosylation and amount of Env make them more infectious and adept at binding to HIV-1 receptors (CD4 and primarily

CCR5) and bind better to dendritic cells. Furthermore, these TF viruses can evade host antiviral immunity such as the type I interferon responses[18, 19, 32]. Once introduced into the

4 body, the free virus can infect CD4 T cells or be taken up and disseminated by dendritic cells (DC), which then infect CD4 T cells by trans-infection[33-35]. Regardless of the site of infection, HIV-1 rapidly replicates in and depletes the CD4 T cell populations throughout the body[36-41], particularly in the gastrointestinal (GI) tract mucosa[36, 40-45] which is likely due to the high abundance of CCR5+ effector memory CD4 T cells at this site[46]. During the acute phase of the infection, there is rapid dissemination of the virus as determined by increases in viral load in the systemic vascular circulation and the infection and depletion of peripheral blood (PB) CD4 T cells[47]. In addition to the loss in CD4 T cells, there is an increase in immune cell activation as characterized by increased frequencies of activated

CD8[48-50] and CD4 T cells (as determined by CD38 and HLA-DR expression)[50-52], circulating pro-inflammatory cytokines[53-56] and innate immune cell activation systemically[47, 55, 57, 58]. Notably, the increase in peripheral CD8 T cell activation has been associated with HIV-1 disease progression and mortality in PLWH[50, 51, 59]. Over the weeks and months following the acute phase of the infection, there is a detectable rebound in CD4 T cell populations in the periphery, an increase in HIV-1 reactive antibodies, a moderate decrease in immune activation and the establishment of a viral set point (a steady state viral load)[47, 60]. This period is characterized as the chronic phase of the infection. If left untreated these individuals can eventually progress to acquired immunodeficiency syndrome (AIDS), characterized by low CD4 T cell counts (below 200 cells/ul), CD4 T cell percentage of total lymphocytes ≤ 14% or the presence of an AIDS- defining illness. At this point in the disease progression, the individual becomes susceptible to AIDS-related illnesses that can ultimately lead to death [47, 61].

Figure 1.1. HIV-1 Viral Life Cycle and the Sites of Action of ART. HIV-1 infection begins with the attachment of the virus to the cell host proteins CD4 and CCR5/CXCR4 and subsequent fusion with the cell membrane. This process can be blocked by fusion and CCR5 inhibitors (1,2). The next primary target in the HIV-1 life cycle is Reverse Transcription, which there are several reverse transcriptase inhibitors (NRTI or NNRTIs) (3). The viral genome is then packaged and integrated into the host genome by the enzyme integrase which can be inhibited by integrase strand transfer inhibitors(4). Lastly, the nascent viral particles are released from the cell, and the viral protein protease cleaves the viral proteins forming the mature virion. This process can be inhibited by protease inhibitors(7). Figure and legend adapted from https://www.niaid.nih.gov/diseases- conditions/hiv-replication-cycle.

HIV-1 Infection: Mucosal Immune Response

Mucosal Immunity in Health

The GI tract mucosal barrier is an essential defense against pathogens and plays a role in the tolerization of the immune system to commensal bacteria and food antigens[62-

64]. The cells most intimately associated with the gut lumen are the epithelial cells, which have various roles that range from nutrient uptake in the small intestine, production of antimicrobial proteins, transport and secretion of IgA antibodies, mucin production by goblets cells, water absorption in the colon and a barrier to commensal and pathogenic microbes[63-65]. Gut epithelial cells are essential regulators of the mucosal immune response through the production of cytokines and other factors when they encounter both commensal and pathogenic microbes[64, 65]. Within the epithelial layer, there are dendritic cells that sample the gut lumen, and several adaptive immune cell subsets termed intraepithelial lymphocytes (IEL)[66, 67]. The IEL fraction of cells is comprised of Mucosal- associated invariant T (MAIT) cells, which respond to conserved microbial antigens, along with CD8 αα and αβ T cells as well as γδ and small populations CD4 T cells[66, 67].

Basolateral to the epithelial barrier lies the lamina propria (LP) and immunological inductive sites such as Peyer's Patch in the small intestine or tertiary inductive sites which are found throughout the GI tract[68]. Specifically, the LP is comprised of various mononuclear cells

(LPMC) that include myeloid DCs (mDC), T helper (Th) CD4 T cells, CD8 T cells, γδ T cells,

NK cells, innate lymphoid cells(ILCs), B cells, macrophages, mast cells and during inflammation neutrophils and plasmacytoid DCs (pDCs)[62, 63, 68-72]. In health these cells work in concert to promote epithelial barrier integrity and function through the production of pro-barrier forming cytokines such IL-22[73], which functions in part through the promotion of stem-cell mediated proliferation of epithelial cells and the induction of antimicrobial peptides by the epithelium [42, 62, 64, 65, 70, 74].

The lamina propria of the gut is densely populated with CD4 T helper cells. These cells are largely comprised of Th1, Th17 and Th22 cells with lower frequencies of Th2 and T regulatory cells[75]. These cells play an essential role in the defense against pathogens and maintenance of the epithelial barrier. The majority of the Th cells in the mucosa are Th1 cells[75], which respond to activation and antigen presentation by producing cytokines such

IFNγ and the express the transcription factor Tbet[76]. The other Th subsets are present in the gut mucosa at much lower frequencies. The Th17 cells mainly produce IL-17 but can be polyfunctional and produce IFNγ and IL-22 and are characterized by the expression of the transcription factor RORγt[76]. The Th22 cells mainly produce IL-22 and express the transcription factor AHR[76]. The T regulatory cells produce multiple regulatory cytokines such as IL-10 and TGFβ and are largely characterized by the expression of the transcription factor FOXP3[76]. Lastly, Th2 cells are characterized by the expression of GATA3 and produce cytokines such as IL-4 and IL-13 which are essential in the defense against parasitic infections[76]. While these cells have unique roles in the gut mucosa, they share several similarities in the mechanisms by which they are activated as well as in their phenotype.

In general, there are several conserved steps in CD4 T cell activation which can be broken into three signals. The first signal consists of antigen presentation from antigen presenting cells (APCs) such as dendritic cells, macrophages, and B cells[77]. These cells take up antigen and process it into peptides that are presented in the major histocompatibility complex II (MHC)II. Once presented, this antigen is recognized by the T cell receptor (TCR), which in most CD4 T cells consist of α and β chains that have undergone sequence recombination (V-D-J (β) and V-J(α)) to recognize specific peptides presented by MHCII[77]. While this signal is important for T cell activation it is not sufficient.

The second signal that is involved in CD4 T cell activation is the engagement of co- stimulatory molecules on the cell surface, such as CD28, by co-stimulatory proteins on the

APC surface such as CD80 and CD86[77]. In addition to the activation of CD4 T cells through CD28, several other surface molecules alter T cell activation, such as OX40 and

CTLA-4[78]. The combination of these two signals is thought to be the primary mechanism by which CD4 T cells are activated. The exposure of CD4 T cells to cytokines, such as IFNγ and IL-2, is considered to be a third signal that drives CD4 T cell activation and proliferation as well as differentiation in naïve CD4 T cells[79]. The culmination of these signaling events at the cell surface drive intracellular changes such as the activation of the transcription factors NF-κB and AP-1 and increases in cytosolic calcium concentrations which then activate calcineurin and ultimately promote the activity of transcription factor NFAT[78]. The combination of the activation of these transcription factors drives CD4 T cell activation and proliferation.

The Th cells that reside in the gut mucosa have recently been described to have a unique phenotype and function relative to other Th subsets in the periphery and are primarily considered tissue-resident memory (TRM) CD4 T cells[80, 81]. These cells are resident in the mucosa and are generated in the mesenteric lymph nodes when APCs present gut-derived antigens[82]. These T cells can respond rapidly to antigen presentation and produce cytokines such as IL-17 and IFNγ[82, 83]. Phenotypically, they differ from the memory T cells in the peripheral circulation in that they have an effector memory phenotype

(CD45RO+CD62L low) and have high expression of CD69 (a marker of acute activation in other CD4 T cell populations) and lower expression of S1P1(sphingosine 1-phosphate receptor)[80, 82]. CD69 is typically upregulated on CD4 T cells early during activation when the cell needs to be sequestered, such as in an inductive site (i.e., lymph nodes). S1P1 expression on T cells allows them to egress and access the peripheral circulation after education in lymph nodes[84]. Therefore, there are likely signals in the mucosa driving the expression of these markers. The idea of gut-specific signals driving the expression of

CD69 on human CD4 T cells is supported by the observations in Bolduc et al. [85], who

9 noted that the bacterial metabolite acetate, which is found at high concentrations in the gut lumen, promoted the expression of CD69 on CD4 T cells.

Intestinal CD4 T cells are uniquely poised to interface with commensal and pathogenic bacteria, metabolites and food antigens found in the gut lumen. Several studies in mice have outlined how commensal and pathogenic bacteria play an essential role in the differentiation and maintenance of both Th17 and T regulatory cells in the gut mucosa[86-

90]. In an elegant study by Ivanov et al. [91], colonization of the murine small intestine with segmented filamentous bacteria (SFB) induced the differentiation of Th17 cells in the lamina propria and protected these mice from Citrobacter rodentium infection, a murine gut pathogen. While the mechanism of SFB driven Th17 differentiation has yet to be fully understood, several studies have indicated a role for antigen presentation from myeloid cells[86, 87]. Commensal gut bacteria have also been shown to drive T regulatory cell differentiation and function (IL-10 production and inhibitory capacity) in the murine gut mucosa. Specifically, bacteria from clusters IV and XIVa of the genus Clostridium and

Bacteroides fragilis have both been shown to induce the differentiation and function of these cells in the murine colonic mucosa and human in vitro studies[88-90]. While findings have yet to be fully replicated in in vivo human studies, they clearly outline a unique and essential relationship between the microbiota and CD4 T helper cell differentiation and responses.

Mucosal Immunity in HIV-1 Infection

There are early changes in the mucosa during the acute phase of HIV-1 infection in

PLWH[36, 44, 45]. However, it is technically challenging to study the immediate (hours- days) changes in the mucosa after initial infection with HIV-1 in humans. To extensively investigate this issue, several groups have used the primate homolog virus simian immunodeficiency virus (SIV) to characterize the initial events in infection. These studies have given insight into the rapidity of the infection, disruption of the epithelial barrier and depletion of the gut mucosal CD4 T cells [47, 92-95]. While the immunological events that

10 occur during SIVmac infection have been well characterized, the detailed kinetics of these events until recently had not been fully characterized. In an elegant study by Hensley-

McBain et al. [93], the kinetics of mucosal and systemic immune responses during pathogenic SIV infection were evaluated. An important finding from this study was that epithelial barrier integrity was lost early during infection (roughly three days). They determined by proteomic analysis that by three days post-infection there were significant decreases in proteins involved in cadherin-mediated cell to cell adhesion. Conversely, the changes in the immune response were not noted until nearly 14 days post infection. This observation that the epithelium was altered very early was consistent with a previous study that demonstrated the Paneth cells in the colon sensed SIV virus and produced IL-1β before an interferon-based antiviral response was detected and the production of IL-1β was associated with epithelial barrier disruption[94]. While the initial kinetics of epithelial barrier breakdown is less well studied in HIV-1, several studies have noted disruption of the epithelial barrier by the presence of breakdown products of the epithelium and alterations in tight junction proteins during HIV-1 infection[96-99]. Collectively, these studies demonstrate that a rapid and dramatic loss in epithelial barrier integrity precedes the loss of gut mucosal

CD4 T cells and immune cell activation and likely contributes to the increased microbial translocation (MT) noted in both SIV and HIV-1 infections. The increase in MT has been detected in PLWH by increases in serum LPS, LTA, 16srRNA and downstream indicators of bacterial component interaction with the innate immune system (soluble CD14) [100-106].

These indicators of MT are associated with disease progression[107] and increased systemic inflammation in PLWH[100, 101, 104, 105] and persist despite viral suppression on

ART[108, 109].

There is significant activation and dysregulation of the gut mucosal immune system during HIV-1 infection[37, 40, 43, 45, 57, 69, 71, 103, 106, 110-112]. One population of gut immune cells that are dramatically altered during HIV-1 infection is the CD4 T helper (Th)

11 cells in the LP throughout the GI tract. These cells are important regulators of epithelial cell function and provide defense against pathogens. They express high levels of HIV-1 co- receptor CCR5 and are more readily infected than their counterparts in the periphery[46,

113]. During HIV-1 infection, there is a preferential depletion of the Th17[37, 43] and

Th22[114] cells in the gut mucosa of PLWH. The importance of these Th populations in the gut mucosa is highlighted by individuals known as long-term non-progressors who are infected but able to control the HIV-1 virus in the absence of ART and appear to preserve their Th17 populations both in the gut and in the periphery[115, 116]. Furthermore, in the nonpathogenic SIV model, where monkeys (Sooty Mangabeys and African Green Monkeys) that are infected with SIV but remain asymptotic and do not progress to AIDS[117], there is a preservation of the Th17 cells in the mucosa and periphery[43, 118] unlike in the pathogenic SIV model where there is a marked decrease in these populations[118-120].

Together these observations indicate that the loss of these cells is of central importance to the progression of HIV-1. To further understand the importance of Th17 populations and the progression of HIV-1 there have been several in vitro studies carried out. These studies demonstrated increased frequencies of Th17 infection relative to other Th subsets through various mechanisms, including a lack regulatory RNAses that limit HIV-1 infection [121], higher expression of α4β7 [122], higher frequency of CCR5+ cells and production of lower amounts of CCR5 ligands(MIP1-α/β) [123], increased mTOR phosphorylation[124] and lastly increased presence of several intracellular proteins shown to make cells more permissive to

HIV-1 infection [125]. From these studies, it is clear that the mechanism by which Th17 cells are preferentially infected and depleted in vitro is likely multifaceted and needs further investigation.

In addition to changes in the Th populations, there are also alterations in other adaptive and innate immune responses. Several studies have demonstrated that PLWH have an increased frequency of activated myeloid DCs [57], pDCs [71], neutrophils [126,

127] as well as increased inflammatory cytokines such as type I interferons[127, 128]. Along with these changes, there is a further dysregulation of the adaptive immune response in the gut mucosa characterized by dysregulation of B cell function and antibody production[129] and the increased frequency and altered activation of CD8 T cells (Figure 1.2).

The Role of the Gut Microbiome in HIV-1 Infection The human gut microbiome consists of a diverse and vast population of bacteria, and other microbes such as fungi that are thought to be equally if not more abundant than host somatic cells[130]. In the last decade, there has been a multitude of studies examining the composition of the bacteria that co-exist with humans and how those populations vary by location, change during development and disease as well as the differences that arise from the environment, lifestyle and diet[130]. To date, much of the focus of these studies has been on those commensal bacteria found in the GI tract, particularly the stool. The gut microbiome, which is thought to be on the magnitude of 1x1012, plays a vital role in the digestion of starches and other nutrients and is integrally tied to both mucosal and systemic health[130]. Numerous studies have identified perturbations in the composition and diversity of the GI tract microbiome and found associations with these changes in a variety of human disease states, which include atherosclerosis[131], septic shock[132], obesity[133],

IBD[134-137] and psychological disorders such as anorexia nervosa[138] and autism[139].

Importantly, the microbiome is critical in shaping the immune system at mucosal and barrier surfaces. It has been demonstrated that early colonization of the gut lumen is essential in the formation of immune tolerance and epithelial barrier integrity[140, 141]. Furthermore, interruption of this normal development with treatments such as broad-spectrum antibiotics can result in an imbalance of immune cell activation and tolerance driving autoimmune disease and atopy[142, 143].

Figure 1.2. The Gut Mucosal Immune Response During HIV-1 Infection. There are several hallmarks of HIV-1 infection in the gut mucosa. The breakdown of the epithelial barrier and subsequent microbial translocation(MT) have been noted to occur to HIV-1 infection (1). There is also an increase in the frequency of activated innate immune cells (e.g., mDCs) and CD4 and CD8 T cells (2-4). Importantly, the CD4 T cell populations are found to be activated, infected and decreased in frequency (3). Lastly, there is also a dysbiosis of the colonic mucosal and stool microbiomes in PLWH (5). Figure adapted from Dillon et al. 2016 [144]

To address a potential link between the composition of the GI tract microbiome and the mucosal and systemic inflammation seen during HIV-1 infection, several studies have sequenced the 16S ribosomal RNA (rRNA) of the fecal and mucosal microbiomes in

PLWH[101, 103, 111, 145-154]. In both treated and untreated PLWH, there is evidence of a reduction in alpha diversity, a measure of bacterial taxa diversity within an individual, of the bacterial populations in the GI tract[145-148, 150]. Although, there have also been reports of no changes[101, 103, 111, 151, 152] or increases[149, 155] in alpha diversity as well.

Several of the early studies on the GI tract microbiome in PLWH noted consistent alterations in the composition of the microbiome, with increases in several Gram-negative bacteria, many with known proinflammatory potential such as Prevotella,[101, 103, 111, 146, 148,

149, 151, 152] and decreases in Bacteroides and several Gram-positive bacteria, many that contain regulatory bacteria such as families and genera that include butyrate-producing bacteria (BPB) species, such as Lachnospiraceae and Ruminococcaceae [103, 111, 145-

148, 150, 151, 153, 154]. In addition to HIV-1 infection, the lower relative abundance of butyrate-producing bacteria (BPB) such as those belonging to the genera Roseburia and

Faecalibacterium have been characterized in other inflammatory diseases of the colonic mucosa such as Crohn’s disease and ulcerative colitis[135-137]. Therefore, there is an association with the loss of these bacteria and gut inflammation. A study undertaken by

Dillon et al. [103] characterized the colonic mucosal microbiome of untreated PLWH. They demonstrated that there was an HIV-1 associated dysbiosis of the composition of the colonic microbiome that included an increased relative abundance of Gram-negative bacteria along with a lower relative abundance of bacteria in the Firmicutes phylum including

Lachnospiraceae and Ruminococcaceae. Subsequently, they demonstrated that the Gram- negative enteric bacteria increased in the microbiome of chronically infected and untreated

PLWH, can increase HIV-1 infection in an in vitro intestinal cell model of MT and this was mediated by increased expression of CCR5 on LP CD4 T cells[156] (Figure 1.2). While

15 there is an important association of the gut microbiome and HIV-1 infection, recent studies have demonstrated that the men who have sex with men (MSM) lifestyle may also contribute to changes in the fecal microbiome. The MSM lifestyle is thought to be responsible for the noted increased relative abundance of Prevotella in many studies when HIV-1 infected study participants are not matched for sexual behavior[155, 157-159]. The first of these studies carried out by Noguera-Julian et al. [160] examined the fecal microbiome of HIV-1 infected and uninfected individuals and controlled for MSM status. Their studies demonstrated that the fecal microbiomes of their study participants clustered by MSM (Prevotella) and non-

MSM (Bacteroides) lifestyle rather than by HIV-1 serostatus. There have been several follow up studies, which have demonstrated a similar Prevotella rich fecal microbiome in MSM[155] and mucosal microbiomes of individuals who participate in condomless receptive anal intercourse[158]. Importantly, in a study where MSM status was controlled for in a group of

PLWH[155], the relative abundances of stool species BPB, such as Faecalibacterium prausnitzii, as well as families and genera that contain BPB were negatively associated with blood T cell activation and microbial translocation. Therefore BPB relative abundance may still play a role In the regulation of inflammation in PLWH.

Short Chain Fatty Acids

One crucial component of the microbiome is the production of metabolites from the breakdown of ingested starches and proteins. Short chain fatty acids (SCFAs) are metabolic end products of the fermentation of indigestible carbohydrates and in some cases the metabolism of amino acids and peptides by the anaerobic bacteria that colonize the GI tract[161-165]. The primary SCFAs are acetate, propionate, and butyrate, which are present in the lumen of the human colon in a 60:20:20 molar ratio with total levels averaging

100 millimolar (mM) [161-165]. In studies of the human GI tract, SCFAs are present in the gut lumen, portal and systemic circulation[166, 167]. From these studies it is clear that large

16 amounts of SCFA are metabolized crossing the colonic epithelial barrier from the gut lumen(acetate: 53mM, propionate: 20 mM and butyrate: 20 mM), which is reflected in the log fold decreases of these SCFA in the portal circulation(acetate:258 µM, propionate:88 µM and butyrate: 30 µM) from the gut lumen. After passing through the liver, the concentrations of SCFA are further diminished in the systemic circulation with acetate (70 µM) remaining the highest while propionate (5 µM) and butyrate(4 µM) only present at trace to non- detectable levels[166, 167]. Acetate is a net metabolic end product of most anaerobic bacteria in the GI tract and is typically generated via two pathways. The most common route is starch fermentation by most anaerobic bacteria in the colon. The next pathway is specific to acetogenic bacteria, that synthesize acetate from free hydrogen and carbon dioxide in the gut lumen or from formic acid through the Wood–Ljungdahl pathway[162]. Propionate is produced by bacterial species in the Bacteroidetes and Firmicutes phyla through succinate, propanediol or acrylate metabolic pathways with the succinate pathway being dominant[162]. Butyrate is produced via two dominate pathways, the butyryl-CoA: acetate

CoA-transferase and butyrate kinase pathways[162, 168-170]. The butyryl-CoA: acetate

CoA-transferase pathway is the dominant pathway where butyryl-CoA is converted to butyrate in a single step. This pathway is the primary pathway used by BPB in the

Firmicutes phylum, such as those in the Roseburia, Faecalibacterium and Eubacterium genera. The less common route of butyrate production is via the butyrate kinase pathway mostly found to be used in Coprococcus species (Table 1.1) [162, 168-170]. These SCFAs serve multiple roles in the bacterial communities in which are produced, including altering the microenvironment by changing the pH and acting as a source of energy for other bacteria when they are metabolized in a process known as cross feeding[162, 164]. Lastly, they have essential roles in human GI tract health with microbial-derived butyrate acting as a significant energy source for colonic enterocytes and an important modulator of epithelial barrier homeostasis, mucosal immune cell responses and gut health[171-174].

Table 1.1 Bacteria Species SCFA produced Metabolic Pathways Anaerostipes_caccae Butyrate butyryl‐CoA:acetate CoA‐ transferase Anaerostipes_hadrus Butyrate butyryl‐CoA:acetate CoA‐ transferase Butyrivibrio_fibrisolvens Butyrate butyryl‐CoA:acetate CoA‐ transferase Coprococcus_catus Butyrate, butyryl‐CoA:acetate CoA‐ Propionate transferase; acrylate pathway Coprococcus_eutactus Butyrate Butyrate Kinase Coprococcus_comes Butyrate Butyrate Kinase Eubacterium_hallii Butyrate, butyryl‐CoA:acetate CoA‐ Propionate transferase; 1,2‐propanediol pathway Eubacterium_rectale Butyrate butyryl‐CoA:acetate CoA‐ transferase Roseburia_faecis Butyrate butyryl‐CoA:acetate CoA‐ transferase Roseburia_hominis Butyrate butyryl‐CoA:acetate CoA‐ transferase Roseburia_intestinalis Butyrate butyryl‐CoA:acetate CoA‐ transferase Roseburia_inulinivorans Butyrate, butyryl CoA:acetate CoA Propionate transferase; 1,2 propanediol ‐ pathway ‐ Anaerotruncus_colihominis Butyrate Not Determined‐ Faecalibacterium_prausnitzii Butyrate butyryl‐CoA:acetate CoA‐ transferase Subdoligranulum_variabile Butyrate Butyrate Kinase Eubacterium_biforme Butyrate butyryl‐CoA:acetate CoA‐ transferase Eubacterium_cylindroides Butyrate butyryl‐CoA:acetate CoA‐ transferase Odoribacter_splanchnicus Butyrate Amine metabolism Clostridium_botulinum Butyrate Butyrate Kinase Butyrivibrio_crossotus Butyrate Butyrate Kinase Butyrivibrio_fibrisolvens Butyrate butyryl‐CoA:acetate CoA‐ transferase Butyrivibrio_proteoclasticus Butyrate butyryl‐CoA:acetate CoA‐ transferase Clostridium_saccharolyticum Butyrate butyryl‐CoA:acetate CoA‐ transferase Clostridium_symbiosum Butyrate butyryl‐CoA:acetate CoA‐ transferase Table 1.1 Butyrate Producing Bacteria and Pathways Utilized. Adapted from [162, 170]

SCFA Mechanisms of Action Histone Deacetylase Inhibition In addition to the typical regulation of gene transcription by transcription factors, epigenetic modifications of both DNA and the nucleosome play an important role in the regulation of gene expression[175, 176]. Nucleosomes are structural units of chromatin that consist of DNA and the scaffolding histone proteins. Within this structural unit, epigenetic modifications can be made to both the DNA and the histone proteins[175, 176]. Histone modifications (i.e., acetylation) play an important role in the regulation of mammalian gene expression[175, 176]. One of the primary modifications of histones is the addition of acetate to lysine residues in the N-terminal of the core histone proteins (H2A, H2B, H3, H4)[175-

177]. In total there are potentially more than 28 sites where these modifications can occur on the N-terminal tail histones. The addition of acetyl groups is primarily carried out by histone acetyltransferases (HAT) proteins[175-177]. Histone deacetylase (HDAC) proteins function by removing acetyl groups from histone tails. There are in total of four classes of

HDACS ( I, IIa, IIb, III, and IV)[177-180]. These HDAC classes have individual properties and cellular locations, which allow them to uniquely impact chromatin state, gene expression and protein function[177-180]. The HDAC proteins are targets of both naturally occurring and small molecule inhibitors, which are termed Histone Deacetylase inhibitors (HDACi).

HDACi act by inhibition of the enzymatic activity of HDAC proteins and therefore promote the hyperacetylation of histone tails and the open state of chromatin which is due to both the inhibition of the HDAC proteins as well as the continued activity of the HAT proteins[177-

180].

The acetylation of histones is known to impact gene expression. When promoter

DNA is assembled onto a nucleosome that is hypoacetylated, transcription factors cannot bind and initiate transcription, which is likely due to the positive charge of the N-terminal lysines that interacts strongly with DNA phosphates and promotes tight coiling of the DNA

19 into heterochromatin[175-177]. Conversely, in most cases when the histones are acetylated at their N-terminal lysine residues they no longer actively interact with DNA and surrounding proteins allowing access to the DNA and transcription[175-177].

Butyrate, propionate and more recently acetate have been characterized as HDAC inhibitors (HDACi) with the ability to inhibit both class I and II HDAC proteins[85, 180-183].

Butyrate is an aliphatic acid that is a relatively weak HDAC inhibitor with activity in the millimolar range[180]. Butyrate targets and inhibits a large group of HDACs but does not have activity on HDAC-6, HDAC-10 and class III and IV HDACs[180, 181]. To act intracellularly, butyrate must be either transported by surface transporters MCT-1 or

SLC5A8, which are not specific for butyrate[184, 185] or likely diffuse through the plasma membrane. The mechanism of action of butyrate’s HDACi is as a noncompetitive inhibitor, which implies that butyrate binds to a unique site on the HDAC proteins to inhibit the enzyme’s function[180]. Interestingly, there appears to be a sequence-specific regulation of individual genes by butyrate due to the presence of a common DNA sequence that is believed to be a butyrate response element[186, 187]. Through its HDACi activity, butyrate has been shown to regulate a multitude of genes and cellular processes that include components of the mTOR pathway[188, 189], cell cycle progression [186, 187], proliferation[187, 190-192], activation[192, 193] and cell differentiation[187, 194-197]

(Figure 1.3).

G-protein Coupled Receptor and PPARγ Signaling GPCRs are a diverse and abundant class of seven transmembrane spanning proteins, which are crucial in the regulation of a variety of cellular functions. GPCRs have a varied distribution of expression in the body and can be regulated by multiple stimuli. These receptors signal internally through heterotrimeric G-proteins that are made up of either

α(Gαs, Gαi, Gαq, and Gα12), β, and γ subunits[198, 199]. The primary SCFA receptors are

GPR41, GPR43 and GPR109a, of which GPR109a is specific to both butyrate and niacin,

20 whereas GPR41 and GPR43 can be activated by any of the three primary SCFA[200-204].

GPR43 has an affinity for the smaller SCFA such as acetate and propionate while GPR41 preferentially binds larger SCFA (propionate and butyrate) and has a lower affinity for acetate[200-205]. GPR41, GPR43, and GPR109a have been demonstrated to be expressed on a multitude of human tissue, including human immune cells such as neutrophils and monocytes. During signaling GPR109a and GPR41 couple to Gαi/o, while GPR43 couples to Gαi/o and Gαq[199, 206] (Figure 1.3).

Studies have demonstrated butyrate’s ability to signal through the nuclear receptor

PPARγ in colonic enterocytes Alex et al. [207] showed that butyrate acts as an agonist for

PPARγ in human colonic enterocytes and that butyrate’s signaling through this pathway promotes angiopoietin-like 4 production, which plays a role in the regulation of systemic triglyceride levels. More recently, a study by Byndloss et al. [208] determined that loss of butyrate signaling though PPARγ shifted the metabolism of colonic enterocytes toward β- oxidation, which increased available oxygen at the epithelial barrier, and drove increased expression of Nos2 and thus increased luminal nitrate concentrations. They determined that the increase in oxygen and nitrate promoted the dysbiotic expansion of potentially pathogenic Escherichia and Salmonella. Thus, signaling through PPARγ likely represents an important pathway by which the presence of butyrate regulates host responses and the mucosal microbiome (Figure 1.3).

Butyrate and Immune Cell Interactions Due to the observed alterations in BPB frequency and butyrate stool concentrations in both animal models of intestinal inflammation and human disease, the impact of butyrate on immune responses has been widely studied. Butyrate is an essential energy source for epithelial cells and a modulator of epithelial cell function. In the colon, where butyrate exists at higher concentrations[166, 167], butyrate alters colon epithelial cell (colonocyte) function and provides for roughly 70-90% of colonocyte energy demands by providing two acetyl-

CoA molecules for the TCA cycle[171, 174, 192, 205, 209]. Butyrate also stabilizes HIF1α, which promotes an anaerobic environment at the colonocyte lumen interface[172], as well as decreases in the production of Nitric Oxide through agonism of PPARγ[208]. Butyrate has also been shown to alter the immunological responses of colonocytes through the upregulation of IL-10R[173], tight junction molecules such as Claudin-1 and downregulation of Cluadin-2[173, 209], and the production cytokines such as IL-18[204, 210].

The impact of butyrate on the adaptive and innate immune system has been studied widely in mice and human peripheral and immortalized leukocytes. Several studies have examined the effects of exogenous butyrate on in vitro T cell activation and proliferation.

Cumulatively, these studies demonstrated that butyrate decreased CD4 and CD8 T cell cytokine production (IL-17 and IFNγ)[190, 211], activation[190-192], and proliferation[190-

192, 212-214] in response to exogenous stimuli. The mechanisms that drove these observations were primarily apoptosis[190, 192, 212-214] and HDACi[192]. Investigations into butyrate’s impact on CD4 T cell differentiation highlighted the critical role of butyrate in T regulatory cell function and differentiation. These studies demonstrated that butyrate increased the frequency of FoxP3+ CD4 T cells through either GPR43[215] signaling or an increase in Foxp3 gene expression through HDACi[194, 195, 216]. Also, butyrate has been shown to promote the differentiation of Th1 and Th17 cells. These studies demonstrated that depending on cytokine milieu, butyrate along with pro-Th1 cytokines (IL-12 and IFNγ) drove increased frequencies of Tbet expressing Th1 cells[188, 196]. Similarly, butyrate along with pro-Th17 cytokines (IL-6 and IL-1β) drove increased expression of Rorc expressing Th17 cells[188, 197]. In addition, both the concentration of butyrate and the differentiation state of CD4 T cells have been shown to alter the impact of butyrate on CD4

T cells. Kesphol et al. [196] demonstrated that lower concentrations of butyrate (0.25 mM) increased the differentiation of Th regulatory cells, while higher concentrations of butyrate (1 mM) increased the frequencies of Th1 cells. Butyrate has also been shown to increase IFNγ

22 and granzyme B expression in CD8 T cells [217]. Furthermore, a recent study determined that the addition of butyrate to differentiating CD8 and Th1 cells increased their production of IL-10, which helped decrease disease severity in a mouse model of colitis [188].

Collectively, these observations demonstrate the effect butyrate on T cell function and development is varied and depends on the concentration of butyrate and state of differentiation of the T cell.

Butyrate has also been demonstrated to alter B cell function and antibody production. Through HDAC inhibition, butyrate has been shown to alter the differentiation of

B cells and the production of immunoglobulin from B cells [218, 219]. Furthermore, when fiber levels were altered in the mouse colon, there were changes in stool IgA and IgG levels.

These changes were determined to likely be mediated by epigenetic modifications of miRNAs silencing Aicda and Prdm1 mRNAs mediated by butyrate driven HDAC inhibition[218]. Therefore, SCFAs can alter B cell development, B cell responses, and antibody production.

Innate immune cells, which respond to microbial antigens and express TLR receptors, have been demonstrated to be impacted by exposure to butyrate. Neutrophils are an essential first line of defense against invading microbes, and the abundance and activation of these cells are also tied to tissue inflammation as seen in the mucosa of

PLWH[126]. Butyrate decreased the production of reactive oxygen species and phagocytosis of C albicans in a study of rat intraperitoneal neutrophils. These effects were limited to butyrate as acetate and propionate did not have the same impact [220].

Furthermore, it has been demonstrated that butyrate increased human neutrophil apoptosis after activation with the TLR4 ligand LPS[181]. Butyrate has also been shown to regulate antigen presenting cell (macrophage and dendritic cell (DCs)) function and differentiation.

Butyrate impacts DC differentiation through the upregulation of CD1a as well as the imprinting of CD103 and α4β7 expression in differentiating dendritic cells[221]. In

23 differentiated macrophages butyrate, through HDACi, decreased the specific cytokine (IL-6 and IL-12) and functional (nitric oxide production) responses to TLR4 ligation and thus dampened responses to microbes[193]. Moreover, butyrate and propionate inhibited CD8 T cell activation by suppressing APC IL-12 production[222]. In cultured macrophages, butyrate depending on the differentiation media has been shown to promote the differentiation of the more homeostatic M2 macrophage populations[223] or a more anti-microbial subset characterized by the production of anti-microbial peptides S100A8/A9/A12[189]. While, there is evidence that butyrate programs DCs to drive the differentiation of T regulatory cells through both GPR109a signaling and HDACi[224] others have highlighted a deleterious role where butyrate administration to activated DCs drove the production of more IL-23 and oral butyrate administration during a murine model of colitis (DSS colitis) worsened symptoms potentially through DC production of IL-23 and Th17 activity[225].

Finally, butyrate’s ability to regulate innate lymphoid cells (ILCs) has also been investigated in vivo. Butyrate has been demonstrated to decrease the murine ILC3s in the terminal ileum which resulted in a reduction in the frequency of T regulatory cells and increased antigen-specific effector T cells[226]. Furthermore, a murine model of airway hyperresponsiveness demonstrated that butyrate inhibited IL-13 and IL-5 production by

ILC2s, which would likely protect against airway hyperresponsiveness[227]. Collectively, these studies demonstrated that butyrate is a potent mediator of innate immunity and that differentiation state, cytokine microenvironment and disease state all impact the effect of butyrate on these cell types.

Figure 1.3. SCFA Signaling Pathways in Cells. The three primary SCFAs (butyrate, acetate, and propionate) alter target cell function through three primary pathways. They can act as HDACi to change cell function by inhibiting HDAC proteins. The can also signal through surface receptors GPR41 and GPR43, which have an affinity for all three SCFA and GPR109a which has an affinity for butyrate and niacin. Butyrate can also act as a PPARγ agonist. Figure adapted from Louis et al. 2014[228].

Project Rationale: Alterations in Colonic Mucosal Butyrate Producing Bacteria During

Chronic HIV-1 Infection1

Several studies of the gut microbiome of PLWH have demonstrated an HIV-1 associated dysbiosis that is in part characterized by a lower relative abundance of taxa that contain BPB species. A study by Dillon, Kibbie et al. [229], demonstrated that the relative abundance of the summed total BPB species in the colonic mucosa was lower in PLWH.

On the species level, Roseburia intestinalis was lower in PLWH and the relative abundance of this bacteria negatively associated with several markers of systemic inflammation, viremia, and microbial translocation. Furthermore, when the ratio of a known HIV-1 associated enteric commensal bacteria (pathobiont) Prevotella stercorea to R. intestinalis was examined, there were strong positive associations with mucosal T cell activation and inflammation along with markers of microbial translocation(Figure 1.4). These observations suggest that there is a loss of colonic mucosal BPB species in PLWH, that occurs in conjunction with an increased abundance of pathobiont commensal bacteria in some individuals, and this promotes unregulated inflammatory responses in the mucosa and periphery as well as microbial translocation and HIV-1 replication.

Scope of Thesis

The infection and loss of Th cells in the gut mucosa and alterations in the GI tract microbiome are hallmarks of HIV-1 infection. The lower relative abundance of regulatory bacteria that contain BPB has been noted in HIV-1 infection as well as other inflammatory disorders of the gut mucosa. Therefore, it is of critical importance to understand how a loss of mucosal BPB species and thus likely decreased concentrations of mucosal butyrate, influence the function of gut CD4 T cells in the setting of inflammation and HIV-1 infection.

The focus of this thesis was two-part. First, I sought to understand the impact of butyrate on enteric HIV-1 associated commensal bacteria driven LP CD4 T cell activation and HIV-1BAL infection using a lamina propria mononuclear cells (LPMC) model of microbial translocation and acute HIV-1 infection. Next, I sought to understand how butyrate affects T cell receptor-mediated activation and HIV-1CH40 infection of LP CD4 T cells over a wider physiologic concentration range and carry out more extensive studies to determine the mechanism(s) by which butyrate is achieving this. For the first part of the thesis, I hypothesized that butyrate would decrease enteric bacteria-driven LP T cell activation, cytokine production, and HIV-1 infection. For the second part of the thesis, I hypothesized that exposure of LPMC or purified LP CD4 T cells to high physiologic concentrations of butyrate would dampen LP CD4 T cell activation, cytokine production, Th subset proliferation, and HIV-1 infection through its known activity as an HDACi.

When I modeled the impact of butyrate on pathobiont-driven gut LP T cell activation and HIV-1 infection, there were significant decreases in both LP CD4 and CD8 T cell activation as determined by the co-expression of HLA-DR and CD38 as well as by IL-17 and

IFNγ production at 2 mM butyrate. Furthermore, butyrate at this concentration significantly decreased the productive infection of LP CD4 T cells. The addition of butyrate at the lower concentration resulted in non-significant decreases in pathobiont driven T cell activation, cytokine production, and productive HIV-1 infection. Following up on these observations, I next determined that butyrate differentially affected T cell receptor-mediated LP CD4 T cell activation, proliferation, and HIV-1 infection depending on concentration. Specifically, at higher concentrations ≥0.25 mM, I noted that butyrate directly and likely via HDACi decreased LP CD4 T cell activation ( frequency of CD25+ and HLA-DR and CD38 co- expressing cells), proliferation and productive HIV-1 infection. Conversely, at lower physiologic concentrations of butyrate ≤0.125mM, there was a limited decrease in LP CD4 T cell activation but an increase in productive HIV-1 infection which was likely mediated via

27 increased CCR5 expression and viral entry or reverse transcription. These effects at lower concentrations were noted only in the presence of other LPMC and not directly on the LP

CD4 T cell populations. Collectively, the data presented in this thesis contribute to our understanding of the functional impact of alterations in BPB populations and likely mucosal tissue butyrate concentrations on LP CD4 T cell activation state and infection during HIV-1 infection. Furthermore, these observations provide insight into possible butyrate centric interventions for the treatment of HIV-1 mucosal pathogenesis and the sequela of mucosal inflammation that occurs in the setting of both treated and untreated HIV-1 infection.

Figure 1.4. Communities of Butyrate-producing Bacteria (BPB) are Reduced in the Colon Mucosa of Untreated HIV-1 Infected Study Participants. Relative abundances (RA) of the major colonic butyrate-producing species were evaluated in 16S rRNA sequence datasets derived from the stool and colonic biopsies of 14 HIV-1 uninfected control (n=14) and untreated, chronic HIV-1 infected (n=17-18) study participants. (A) Summed RA was determined in the stool, and colonic mucosa of HIV-1 uninfected (HIV-; n=14) (open circles) and untreated, chronic HIV-1 infected (HIV+; Stool n=18; colonic mucosa: n=17) (filled squares) study participants. (B) RA of Roseburia intestinalis in the colonic mucosa of HIV-1 uninfected (HIV-; n=14) (open circles) and untreated, chronic HIV-1 infected (HIV+; n=17) (filled squares) study participants. (C) The ratio of RA of P. stercorea to R. intestinalis (P. stercorea: R. intestinalis) in colonic mucosa of HIV-1 uninfected (HIV-; n=14) (open circles) and untreated, chronic HIV-1 infected (HIV+; n=14) (filled squares) study participants. RA of R. intestinalis was undetectable in 3 HIV-1 infected individuals. Therefore, a ratio was not calculated for these study participants (n=14). (D) Correlations between P. stercorea: R. intestinalis and the number of activated (CD38+HLA-DR+) colon CD4 and CD8 T cells and activated colon CD1c+ mDC (CD40 expression levels) in HIV-1 infected study participants (n=14 for colon T cell activation; n=11 for colon CD1c+ mDC activation). Lines represent median values (A-C). Statistical analysis was performed using the Wilcoxon matched-pairs signed rank test to determine differences in matched groups (A), the Mann-Whitney test to determine differences between unmatched groups (A-C) and Spearman test to determine correlations between variables (D)[229].

CHAPTER II MATERIALS AND METHODS

Human intestinal LPMC Isolation

Human jejunal tissue that was deemed macroscopically normal was obtained from patients undergoing elective abdominal surgery and considered to be discarded tissue.

Tissue samples from individuals with a clinical history of recent chemotherapy (less than eight weeks), hematologic malignancy, radiation, HIV-1 infection, immune-suppressing drug use or inflammatory bowel conditions were excluded from this study. Patients granted release of tissue to allow unrestricted use in research and all samples were de-identified with only age, sex, the reason for surgery and surgery-related treatment status information available. The use of these samples for research purposes was reviewed by the Colorado

Multiple Institutional Review Board (COMIRB) at the University of Colorado Anschutz

Medical Campus and was granted exempt research status. Tissue samples were processed as follows, jejunal tissue was first dissected, and the mucosa isolated mechanically. This was then followed by incubations with dithiothreitol (DTT) to remove mucus, ethylenediaminetetraacetic acid (EDTA) to remove the epithelial layer and lastly digestion with collagenase to isolate LPMC. LPMC were cryopreserved and stored in liquid nitrogen.

PBMC Isolation

PBMC were isolated from healthy whole blood donors (N=6) who voluntarily gave written informed consent. Whole blood collection was approved by COMIRB, University of

Colorado, Anschutz Medical Campus. Blood was drawn in Vacutainer tubes containing sodium heparin. PBMC were isolated using Ficoll density gradient centrifugation and cryopreserved for long term storage in liquid nitrogen.

Growth of Bacteria Stocks

Prevotella stercorea (DSM#18206, DSMZ, Braunschweig, Germany) was cultured at

37 °C under anaerobic conditions using BD GasPak EZ Anaerobe Pouch Systems (BD

Diagnostics, Franklin Lakes, NJ, USA) according to manufacturer’s protocols. P. stercorea was grown in liquid chopped meat broth (Hardy Diagnostics) which was supplemented with

1% Vitamin Supplements (ATCC), 0.05% Tween 80, 1% Trace Minerals (ATCC), 8.1 mM propionic acid, 29.7 mM acetic acid and 4.4 mM butyric acid (Sigma-Aldrich) for 5–7 days.

P. stercorea was enumerated by flow cytometry using BD Liquid Counting Beads (BD) and stored long-term at −80 °C in 10% glycerol. Cells were subsequently suspended in DPBS for working stocks and stored at −80 °C in single-use aliquots.

Preparation of CCR5-tropic CH40 Transmitted Founder Stocks and HIV-1Bal Stocks

HIV-1Bal (AIDS Research and Reference Program #510) and mock-infected supernatant stocks were generated and frozen in single-use aliquots stored at −80°C as previously described[156, 230]. The Beatrice Hahn laboratory generated transmitted founder

HIV-1 infectious molecular clone CH40. Plasmids containing CH40 were re-transformed and amplified in Stbl3 cells (Invitrogen) and purified using Qiagen maxi kit. The CH40 maxi-preps were verified by 13 HIV-specific primers to cover the entire genome. 40 μg of transmitted founder plasmids were used to transfect 293-T cells by CaCl2 transfection method. Virus- containing and uninfected supernatants were collected at 48 hours, concentrated by ultracentrifugation at 141,000 RPM over a 20% sucrose cushion to generate both viral and mock stocks. Viral stock concentrations were determined by an HIV-1 Gag p24 ELISA kit

(ABI). Frozen one-time use aliquots of virus and mock were frozen at -80°C for future use.

LPMC and PBMC Culture Model.

LPMC and PBMC were counted and plated at 1x106 LPMC/mL in 48 well culture plates using complete culture media (RPMI + 10% human AB serum + 1% penicillin/streptomycin/glutamine + 500µg/ml Zosyn (piperacillin/tazobactam) at 37˚C, 5%

CO2 and 95% humidity from 24 hrs. - Four days. T cell activating beads (anti-

CD3/CD2/CD28; Miltenyi) were added to culture conditions in a 1:25 (bead: LPMC) ratio and

Prevotella stercorea 2.5:1 ( bacteria: LPMC). For assays using purified cells, LP CD4 T cells were isolated using the EasySep immunomagnetic negative selection kit (Stemcell

Technologies) according to the manufacturer’s protocol. To control for variance in cell responses due to the isolation process, total LPMC were also incubated with the separation buffer in place of the CD4 T cell Isolation Antibody Cocktail and all subsequent steps in the protocol were the same. On average LP CD4 T cells were enriched to 93% of viable leucocytes (n=6).

SCFA Assay Conditions

For all conditions exposed to SCFA, butyric acid, glacial acetic acid, and propionic acid were diluted in culture media to working stocks (10-20 mM; 7.5pH) and then serially diluted at the time of assay set up (SCFA concentrations, acetate: 0.25-10mM, propionate

0.25-2mM, butyrate 0.0625-0.5mM). Unless otherwise noted, LPMC and PBMC were cultured with SCFAs added concurrently with TCR activating beads or Prevotella stercorea.

For studies examining the impact of butyrate pre-treatment on LP CD4 cell proliferation,

LPMC were plated and exposed to butyrate (butyric acid) for 0, 2, 6 and 8 hours before exposure to TCR activating beads. The conditions examining the impact of butyrate addition after TCR activating bead exposure, butyrate was at 0, 6, 18 and 36 hours after TCR bead exposure.

Small Molecule PPARγ, HATi and HDACi Assays

LPMC were set up as previously mentioned in the sections describing LP CD4 T cell activation and LP CD4 T cell proliferation. Rosiglitazone, T 0070907, C646, trichostatin A, apicidin, MGCD, and ITF2357 were diluted to working stocks of 1000nM in culture media and serially diluted at the time of assay set up. Apicidin, MGCD, and ITF2357 were kindly provided by Timothy McKinsey (University of Colorado Anschutz Medical Campus). The

32 impact of HDACi on TCR mediated LP CD4 T cell activation, and proliferation was determined by the examination of changes in LP CD4 T cell expression of activation markers (HLA-DR+CD38+ and CD25+) and proliferation (CFSEdim) while studies with HATi and PPARγ focused assays only used proliferation as a readout.

Quantification of Secreted Cytokine Production

Levels of IL-17, IL-2 and IFN in culture supernatants were quantified using Ready-

Set-Go ELISA kits (ThermoFisher). Assays were carried out according to the manufacturer’s protocols.

FLOW Acquisition and Analysis

All flow cytometry data were acquired on an LSRII flow cytometer (BD Biosciences) and analyzed using FlowJo version 10. The Cytometer Setup and Tracking feature using BD

FACSDIVA software (V6.1) and routine quality control were performed daily as previously described[231].

Measurement of LP CD4 T Cell Activation

After four days of culture with or without butyrate and activating stimuli, culture conditions were harvested, surfaced stained and analyzed by multicolor flow cytometry.

Detailed gating strategies are outlined in supplemental figures (Figure 2.1, Table 2.1)). To enumerate populations of interest, viable LP CD4 T cells were identified (Zombie dye negative(Biolegend) within a total lymphocyte gate based on forward and side scatter properties cells and doublets were excluded using forward-scatter-height versus forward- scatter-width properties. CD4 T cells were identified as CD3+(PerCpCy5.5, TONBO, Clone:

OKT3), CD4+ (V450, TONBO, Clone: RPA-T4) and CD8- (APC, TONBO, Clone: RPA-T8), cells. LP CD4 T cell activation was determined by enumerating the percentage of CD4 T cells that co-expressed CD38 (AF700, eBiosciences, Clone: HIT2) and HLA-DR (APC-Cy7,

BioLegend, Clone: L243) as well as the percentage of cells that expressed CD25 (PE, BD,

Clone: M-A251). In studies with P stercorea, activation was only enumerated by CD38 and

HLA-DR. For final reported values, isotype and FMO control values were subtracted and data displayed as the net value.

Measurement of LP CD4 T Cell Proliferation

In experiments enumerating proliferation of LP T cells, LPMC or purified LP CD4 T cells were labeled with 1M CellTrace CFSE (Invitrogen) per manufacturer’s instructions before setting up culture conditions. CFSE-labeled cells were exposed to various concentrations of SCFA and HDACi as mentioned above. LPMCs were cultured with or without TCR activating beads for four days at 37˚C, 5% CO2 and 95% humidity. Cultures were collected, and CD4 T cell proliferation (CFSEdim) was determined by multi-color flow cytometry. Proliferating CD4 T cells were enumerated by the percentage of CFSEdim CD4 T cells with the CFSE gate established on the unstimulated control condition(Figure 2.1,

Table 2.1).

Measurement of LP CD4 T Regulatory Cell Frequency

LPMC were cultured for either 24 hrs or four days in the presence or absence of butyrate (0.25 and 0.5mM). At baseline and after culture cells, were stained using FOXP3

Fix/Perm Buffer Set (Biolegend) with the following antibodies: CD45+(PerCpCy5.5, eBiosciences, Clone: 2d1), CD3+(PE-Dazzle, Biolegend, Clone: UCHT1), CD4+ (APC,

TONBO, Clone: RPA-T4) and CD8- (PE-Cy7, TONBO, Clone: RPA-T8), CD25 (PE, BD,

Clone: M-A251) and FOXP3 (AF488, eBiosciences, Clone: PCH101) and rat IgG2a

(eBiosciences) (Table 2.1).

LP and PB CD4 T Helper Cell Proliferation

LPMC or PBMC were labeled with CFSE initially and after 4 days of culture, cells were stimulated with a final concentration of phorbol myristate acetate (25ng/ml) and

Ionomycin (5ng/ml) in the presence of 1 μg/ml Brefeldin A (Golgi Plug) for 4 h at 37°C in 5%

CO2. LPMC were collected and measurement of proliferated Th cells determined using multi-color flow cytometry. LP CD4 T cells were identified using the following antibodies:

CD3 (PerCpCy5.5, TONBO, Clone: OKT3), CD4 (APC, Biolegend, Clone: RPA-T4) and CD8

(PE-Dazzle, TONBO, Clone: RPA-T8) and viability dye. Cells were fixed using Medium A

(Invitrogen, Carlsbad, CA) and permeabilized using Medium B (Invitrogen). Intracellular cytokine staining was then used to characterize Th subsets. The following antibodies were used IL-17 (V450, eBiosciences, CLONE: N49-653), IFN (AF700 BD, clone: B27), and IL-

22 (PE-Cy7, Biolegend, clone:22URTI). Matched isotypes were used to determine positive staining and CFSE labeling to assess proliferation. Th subsets were characterized as follows: Th17 were any LP CD4 T cell producing IL-17, Th1 were LP CD4 T cells that produced IFN and could also produce IL-22, Th22 were LP CD4 T cells that only produced

IL-22, and lastly there was a subset characterized the absence of the production of these three cytokines (Figure 2.2, Table 2.1)

Studies to Understand Butyrate’s Mechanism of Action

Quantification of Histone H3 Acetylation

To determine Histone H3 lysine 9 (H3K9) acetylation levels by flow cytometry, LPMC were cultured with or without butyrate (0.5mM) and TCR activating beads and harvested after 24 hours of culture. LP CD4 T cells were identified using the following antibodies: CD3

(PerCpCy5.5, TONBO, Clone: OKT3), CD4 (APC, Biolegend, Clone: RPA-T4) and CD8 (PE-

Dazzle, TONBO, Clone: RPA-T8) and viability dye. Cells were next fixed using Medium A and permeabilized using Medium B (Invitrogen) and stained according to the protocol outlined in [232] for acH3K9(Millipore) and a Donkey anti-rabbit IgG secondary antibody

(AF488, Biolegend) to determine acH3 expression. Expression levels of acH3 were determined by LP CD4 T cell mean fluorescence intensity (MFI) with secondary only values removed (Figure 2.3, Table 2.1).

Figure 2.1. Flow Gating Scheme for Stimulated LP CD4 T cell Activation and Proliferation Studies. LPMC were stimulated with TCR stimulating beads for four days. Cells were harvested and stained with fluorescent antibodies. In short, live cells were first gated using a viability exclusion dye, FSC-A and SSC-A next determined size and granularity. FSC-W and FSC-H then excluded doublets. Next CD3+ cells were gated, followed by CD4+CD8- cells. Activation (CD25, HLA-DR and CD38) were gated using FMO (CD25, HLA-DR) and Isotype (CD38) controls. Proliferation (CFSEdim) was gated based on corresponding conditions without exogenous stimulation.

Supplemental Figure 2.2. Flow Gating Scheme for the Identification of T helper Subsets Flow gating scheme for the identification of T helper subsets. In short, LPMC were activated with TCR stimulating beads for four days after which time T helper subsets were determined by cytokine expression after PMA/ionomycin stimulation + Golgi plug for the final four hours of culture. Proliferation (CFSEdim) was determined for the following populations: Th17 (all IL-17+ LP CD4 T cells), Th1(LP CD4 T cells that express IFNγ but not IL-17 but can express IL-22), Th22 (LP CD4 T cells that only express IL-22) and LP CD4 T cells that do not express IFNγ, IL-17 or IL-22(A).

Figure 2.3 Butyrate Increases LP CD4 T Cell Histone Acetylation Levels (acH3K9). Example flow plots for LPMC treated with butyrate 0.5mM or 0mM control in the setting or absence of TCR stimulation. LP CD4 T cells were gated, and acH3K9 MFI values determined acH3K9 expression after 24 hours. Secondary only staining was used for controls, and MFI values were removed for reported values.

H3K9 protein levels were also assessed using western blot in the absence of stimulation. Purified LP CD4 T cells were cultured in the presence of butyrate at 0.125 and

0.5mM for 24 hours. Protein was extracted using Tris lysis buffer in the presence of a protease inhibitor (Roche). Total protein was quantified for normalization using a Pierce

BCA protein assay kit. The following antibodies were used: anti–-actin (conc: 1:50,000,

Abcam) and anti-acH3K9 (conc: 1:10,000, Rabbit Polyclonal, Millipore). Peroxidase goat anti-rabbit IgG and peroxidase goat anti-mouse IgG were used according to manufactures recommendations. Pierce ECL substrate (Thermo Fisher Scientific) and Criterion XT Bis-Tris

Gels (Bio-Rad) were also used, and expression levels were quantified using ImageJ software.

LP CD4 T cell GPR Expression

GPR43 and GPR109a expression were determined at baseline and after four days culture on LP CD4 T cells. LP CD4 T cells were cultured in the presence or absence of TCR activation and butyrate (0.5mM). LP CD4 T cell flow staining was determined by CD3+

(PerCpCy5.5, TONBO, Clone: OKT3), CD4+ (TONBO, Clone: RPA-T4) and CD8- (PE-

Dazzle, TONBO, Clone: RPA-T8) and viability dye. Cells were also stained for GPR109a

(PE, R&D, clone: 245106) and GPR43 (AF647, Bioss Antibodies, clone: polyclonal) expression. Isotypes were used to determine positive staining (Table 2.1).

Flow-cytometry Based Measurement of Productive HIV-1 Infection

Total LPMCs and purified LP CD4 T cells were spinoculated with CH40 transmitted founder virus (100ng p24 per 1x106 LPMCs) in a final concentration of 2.5x106 LPMCs/ml.

LPMCs were mock infected using culture supernatants from 293T cell cultures and conditioned in the same manner as the HIV-1 infected cells. Spinoculation was carried as previously detailed[156, 231]. LPMCs were cultured with butyrate and stimulation as described earlier. After 4 days of culture, productive infection was measured by the frequency of total CD3+( PE-Dazzle, Biolegend, clone: UCHT1), CD8- (APC, Tonbo, clone:

RPA-T8) cells and Th subsets (enumerated after treatment with phorbol myristate acetate and Ionomycin in the presence of 1 μg/ml Brefeldin A), expressing the viral nucleocapsid protein p24 (PE, Beckman Coulter, clone: KC57). CD3+CD8- cells were used to determine the productive infection of total LP CD4 T cells since it is known that CD4 is down-regulated during HIV-1 infection[233]. p24 background staining in mock conditions was removed to determine true p24 positive populations in the HIV-1 infected conditions.

HIV-1 Infection Related Assays

TZM-bl Assay

Infectious titers present in culture supernatants were determined in TZM-bl reporter cells after 4 days of culture. TZM-bl cells were plated in 96-well plate white polystyrene microplates in culture media (DMEM+ 10% FBS + 1% PSG) with dextran sulfate (100 ng/ml). Culture supernatants from infection assays were then added to individual wells and incubated for 48 hours at 37˚C, 5% CO2 and 95% humidity. Brite lite luciferase reagent

(Perkin Elmer) was added to each well. A VictorX5 plate reader (Perkin Elmer) was then used to determine Relative Light Units (RLU) of luminescence.

HIV-1 Co-receptor Expression Analysis

LPMC and purified LP CD4 T cells were cultured as previously described. After four days of culture, cells were stained for extracellular expression of CCR5 (PE-Cy7, BD, clone:

2D7/CCR5) and CD4 (V450, Tonbo, clone: RPA-TA4). LP CD4 T cells were gated as previously mentioned, and CCR5 and CD4 MFI values of each were reported with isotype values removed(Figure 2.5, Table 2.1).

HIV-1 RU-5 Quantification

To quantify HIV-1 RU-5 DNA, LPMC were spinoculated, activated and exposed to butyrate as outlined above and exposed to Raltegravir (NIH AIDS Reagent Program) at 1M to prevent integration of HIV-1 into the host genome. After 24 hours of culture, LP CD4 T cells were sorted using Astrios Cell Sorter (University of Colorado Cancer Center), and DNA

40 was isolated using DNeasy mini kit (Qiagen). Primer and probe (TaqMan) combinations were used to quantify HIV-1 RU-5 and RNaseP (Thermofisher). Each reaction contained 10 pmol of primer and probe, and Gene Expression Mastermix (Life Technologies) was used according to manufacturer’s instructions. The following thermocycling conditions were used:

50°C 2 min and 95°C 10 min, then 40 cycles of 95°C 15s and specific annealing temperatures for each target were used (RU-5: 60°C 45 s; RNaseP: 60°C 60). qPCR reactions were run on the Biorad CFX96 rtPCR machine. Primer sequences: RU-5: F-

CTGGCTAGCTAGGGAACCCACTG,R-GAGTCCTGCGTCGAGAGAGCTCC,P-

TCAAGTCCCTGTTCGGGCGCCACTG. RNase P cell number quantification (Thermofisher) was used according to manufacturer’s recommendations.

HIV-1 Gag Quantification

To quantify HIV-1 gag RNA, LPMC were spinoculated, activated and exposed to butyrate as outlined above in the presence of Amprenavir (NIH AIDS Reagent Program

Bethesda, Maryland) at 1 µM to prevent multiple rounds of infection. After 48 hours of culture LP CD4 T cells were isolated by negative selection as outlined above (purity on average 88%) and RNA was isolated using RNeasy minikits (Qiagen). Primer and probe

(Taqman) combinations were used to quantify HIV-1 gag and GAPDH. Each reaction contained 10 pmol of primer and probe, and Gene Expression Mastermix (Life

Technologies, Carlsbad, CA) was used according to manufacturer’s instructions. The following thermocycling conditions were used: 50°C 2 min and 95°C 10 min, then 40 cycles of 95°C 15s and specific annealing temperatures for each target were used (GAPDH:

64.5°C 45 s; gag: 65.5°C 45 s). qPCR reactions were run on the Biorad CFX96 rtPCR machine. Primer sequences: GAPDH: F CCCATGTTCGTCATGGGTGT R-

TGGTCATGAGTCCTTCCACGAT, P-CTGCACCACCAACTGCTTAGCACCC; gag: F-

YCAAGCAGCYATGCAAATGTTAAA, R- CTATGTCACTTCCCCTTGGNTCT, P-

TCTATCCCATYYTGCAGCTTCCTCATTGAT

Statistical Analysis

Statistical analysis and graphing were performed using GraphPad Prism Version 6 for Windows 10. Parametric two-tailed tests were performed. Mann-Whitney tests were used to compare unmatched groups, and correlations analyzed using Spearman's test.

Differences between groups of matched paired data were performed using the Wilcoxon's matched-pairs signed-rank test, and multiple groups of matched paired comparisons were performed using the Friedman test with multiple Dunn comparisons. Significance was determined at P< 0.05. Inhibitory Capacity of 50% was determined using raw data values and log-transformed SCFA or apicidin concentrations using the following function log(inhibitor) vs. response with a variable slope (four parameters) with interpolation of unknowns from the standard curve.

Figure 2.4. Flow Gating Scheme for the Identification of Productive Infection of LP CD4 T Cells Flow gating scheme for the identification of productively HIV-1 infected (p24+) LP CD4 T cells after four days of infection in the setting of TCR stimulation. LPMC were harvested and stained by multicolor flow cytometry. Cells were gated as follows: Viable, single cell lymphocytes were gated first by CD3. CD3 cells were then gated into CD8 negative population due to the down-regulation of CD4 by HIV-1 infection. These cells were then gated by p24 expression with Mock control conditions acting as gating controls.

Figure 2.5. Flow Gating Scheme for the Quantification of CCR5 Expression on LP CD4 T Cells LPMC were cultured for four days in the setting of TCR stimulation and cells were then harvested, stained and examined by multi-color flow cytometry. Populations were gated as follows: Viable, single cell lymphocytes were gated first by CD3. CD3 cells were then gated into the CD4+ CD8- population. These cells were then gated by CCR5 expression with isotype stain acting as gating controls for CCR5. CD4 and CCR5 expression were determined by the MFI of CD4 and CCR5 in the CD4+ population with isotype MFI values removed for CCR5.

Table 2.1 Flow Antibodies

CHAPTER III EXOGENOUS BUTYRATE DECREASED ENTERIC PATHOBIONT DRIVEN LP T CELL

ACTIVATION AND HIV-1 INFECTION2

Preface

The data presented in this chapter is comprised of sections of a manuscript that I co- first authored with Dr. Stephanie Dillon. The clinical data generated for my co-first author work is detailed in the introduction.

Background

Human immunodeficiency virus (HIV)-1 disease is associated with dramatic alterations in the gut-associated lymphoid tissue (GALT) that occur early in the course of infection and contribute to HIV-associated chronic immune activation and disease progression. HIV-associated GI tract pathology is characterized by early and high-level HIV-

1 replication and a dramatic depletion of mucosal effector CD4 T cells [36, 40, 45] especially

T helper (Th) 17 and Th22 cells [43, 44, 102, 114-116, 234] in GALT. Concomitant with T cell depletion, increased frequencies of activated gut CD8 T cells [57, 235-237], alterations in innate immunity [57, 71, 126, 238-241], generalized gut inflammation [242, 243] and significant epithelial barrier damage [126, 236, 244-246] occur. The cumulative impact of these structural and immunological changes is an increase in translocation of commensal bacteria and bacterial products from the gut lumen into the lamina propria (LP) and ultimately into the systemic circulation, a process called microbial translocation (MT). MT has been associated with systemic immune activation [100] and predicts disease progression in untreated HIV-1 infected individuals [107]. Its persistence during effective

46 antiretroviral therapy (ART) predicts mortality and is associated with several co-morbidities and immune activation and inflammation [247, 248].

Intestinal bacterial communities (microbiome) and their metabolites are essential in shaping and maintaining gut immunity [249]. Recent studies by our group and others demonstrated dysbiotic fecal and intestinal mucosal microbiomes during both untreated and

ART-treated HIV-1 infection [57, 103, 111, 146, 147, 149, 151, 153, 250-255]. Our lab previously demonstrated that the relative abundance (RA) of Gram-negative Proteobacteria and Prevotella spp. were increased whereas Firmicutes and Bacteroides spp. were reduced in the colonic mucosa of individuals with untreated HIV infection. This dysbiotic profile was associated with MT and mucosal and systemic immune activation. Recent work exploring the impact of sexual preference on HIV-1 associated gut microbiome dysbiosis determined that a Prevotella rich dysbiosis was associated with men who have sex with men (MSM) sexual preference rather than with HIV-1 infection status [160]. Although the increase in RA of Prevotella in our study could have been due in part to sexual preference, our finding that mucosal Prevotella spp RA correlated with colonic myeloid dendritic cell (mDC), and T cell activation [57, 103] provides a link between dominant mucosa-associated microbes and inflammation in the setting of MT. We have also recently demonstrated that commensal bacteria of the Prevotella genus have direct pro-inflammatory potential by activating gut mDC and increasing HIV-1 replication and CD4 T cell death in an ex vivo human intestinal cell model [57, 256]. Therefore, although the increase in Prevotella observed in our study

[57, 103], as well as in others [146, 149, 255] could potentially contribute to HIV-1 associated gut pathology.

Microbial metabolites, byproducts produced by intestinal bacteria, are also important in influencing immune responses [165]. In our previous study [103], we observed a decrease in RA of Firmicutes, a phylum which includes bacteria known to produce the short chain fatty acid (SCFA) butyrate [257]. Butyrate, a fermentation product of insoluble dietary

47 fiber, is one of three primary SCFAs produced in the colon [258], and decreases in abundances of butyrate-producing bacteria (BPB) occur in several inflammatory diseases of the GI tract such as Crohn’s disease, ulcerative colitis and colon cancer[259-261]. Butyrate has been reported to play essential roles in colonic T regulatory cell differentiation [195, 204] and in modulating both gut and blood antigen presenting cell (APC) and T cell activation

[190, 193, 262, 263]. In addition, butyrate is an essential energy source for colonic epithelial cells that promotes epithelial barrier integrity [264-267]. However, despite the known immune-regulatory role of butyrate, little is known about its role in regulating mucosal inflammation in the setting of HIV-1 infection.

In this chapter, I investigated the impact of exogenous butyrate on Prevotella stercorea driven T cell activation and HIV-1 infection using an ex vivo human intestinal cell model of MT and acute HIV-1 infection.

Results Butyrate Reduces P. stercorea-driven Activation of LP T Cells in vitro

The clinical study observations outlined in the introduction demonstrated that increasing abundances of Prevotella spp. in conjunction with decreasing abundance of R. intestinalis associated with increased colon T cell activation. This observation led me to hypothesize that a loss of butyrate may exacerbate P. stercorea driven intestinal T cell activation. To address this, I used an ex vivo human LPMC model which allows us to mimic the effect of translocating bacteria (i.e., MT) on LP T cell function in the setting of HIV-1 infection. In these in vitro assays, butyrate at a high (2mM) and low (0.2mM) concentration were used which agree with published studies using butyrate[192, 193, 268].

To determine if butyrate modulated the activation of LP CD8 and CD4 T cells in the absence of HIV-1 infection, Mock-infected LPMC were exposed to Prevotella stercorea and butyrate (0.2 and 2mM) for four days. In the absence of butyrate, P. stercorea significantly

Figure 3.1 Butyrate decreases P. stercorea driven lamina propria CD4 and CD8 T cell activation and cytokine production in the absence of HIV-1 infection in vitro. Lamina propria mononuclear cells (LPMC) were spinoculated with mock-infected culture supernatants and cultured with or without P. stercorea and butyrate (0.2mM, 2mM) for four days and levels of T cell activation and cytokine production determined (n=9). Percentages of LP CD4 (A) and CD8 (B) T cells co-expressing CD38 and HLA-DR were assessed using multi-color flow cytometry. FMO and isotype control values have been subtracted. Levels of IFNγ (C) and IL-17A (D) in culture supernatants at day 4 were evaluated by ELISA. P. stercorea-specific (Δ) values are shown as net values (P. stercorea/mock minus mock alone). Statistical significance was determined using the Wilcoxon matched pairs signed rank test and the Friedman’s test with comparisons to no butyrate controls. (**P<0.01, ***P<0.001). Data displayed as box and whiskers are represented with the box extending from the 25th to 75th percentile with the median value represented as the central line in the box and maximum and minimum values represented by the whiskers.

Figure 3.2 Butyrate reduces P. stercorea-enhanced LP T cell activation and HIV-1 productive infection Lamina propria mononuclear cells (LPMC) were spinoculated with CCR5-tropic HIV-1Bal and cultured with or without P. stercorea and butyrate (0.2mM, 2mM) for 4 days and levels of T cell activation, cytokine production and productive HIV-1 infection determined (n=9). Percentages of LP CD4 (A) and CD8 (B) T cells co-expressing CD38 and HLA-DR were assessed using multi-color flow cytometry. FMO and isotype control values have been subtracted. Levels of IFNγ (C) and IL-17A (D) in culture supernatants at day 4 were evaluated by ELISA. Percentages of p24-expressing LP CD4 T cells (E-F) were assessed using multi-color flow cytometry. P. stercorea-specific (Δ) values are shown as net values (P. stercorea/HIV-1 minus HIV-1 alone). Statistical significance was determined using the Wilcoxon matched pairs signed rank test and the Friedman’s test with comparisons to no butyrate controls. (**P<0.01, ****P<0.0001). Data displayed as box and whiskers are represented with the box extending from the 25th to 75th percentile with the median value represented as the central line in the box and maximum and minimum values represented by the whisker

50 increased LP CD4 and CD8 T cell activation (co-expression of HLA-DR and CD38) and cytokine production (IFNγ and IL-17) (Figure 3.1). When exogenous butyrate was added to these cultures, butyrate at 2mM decreased LP T CD4 and CD8 T cell activation(Figure 3.1

A-B) and cytokine production(Figure 3.1 C-D) with minimal effects seen at 0.2mM butyrate.

Butyrate Reduces P. stercorea-driven Activation of HIV-1 Infected LP T Cells in vitro

In the absence of butyrate, the addition of P. stercorea to HIV-infected LPMC cultures significantly increased LP CD4 T cell activation levels to above those induced in with infection alone (Fig. 3.2A). The addition of 2mM butyrate significantly decreased P. stercorea-associated LP CD4 T cell activation (Fig. 3.3A) to that at or below levels observed with HIV infection in the absence of bacteria, whereas 0.2mM butyrate had minimal effect on

LP CD4 T cell activation. Similar suppressive effects of higher doses of butyrate were observed on LP CD8+ T cell activation, which was also increased in the presence of P. stercorea (Fig. 3.2B).

Butyrate Decreases LPMC Production of IFNγ and IL-17 in Response to P. stercorea

Based on the observation that butyrate reduced P. stercorea-driven LP T cell activation, I next evaluated T helper cytokine production as a functional readout of LP CD4 T cell activation. The addition of P. stercorea to HIV-infected LPMC cultures led to higher production of IFNγ and IL-17 in culture supernatants (Fig. 3.2C and D) compared to HIV- infected cultures alone. Butyrate inhibited both IFN-γ(Fig. 3.2C) and IL-17 (Fig. 3.2D) production in a dose-dependent manner, with complete suppression observed at the higher

2mM dose

Butyrate Inhibits P. stercorea Driven HIV-1 Infection in LP CD4+ T cells

We have previously shown that P. stercorea enhances LP CD4 T cell infection levels in our LPMC model[156]. Therefore, I aimed to investigate the impact of butyrate on P. stercorea-driven LP T cell infection. In agreement with this previous study, exposure of HIV-

1Bal-infected LPMC to P. stercorea in the absence of butyrate significantly increased the

51 percentage of p24+ LP CD4 T cells compared to HIV-1Bal infection alone (Fig. 3.2E).

Addition of 2mM butyrate suppressed P. stercorea driven LP CD4 T cell infection to levels observed with virus infection alone, whereas 0.2mM butyrate had a minimal effect (Fig.

3.2F). Butyrate had no noticeable impact on HIV-1Bal infection levels detected in the absence of P. stercorea (Fig. 3.2F).

Discussion Microbial translocation has been linked to mucosal and systemic inflammation and is predictive of disease progression and mortality in HIV-1 infected individuals [247, 248]. It is thus of critical importance to understand the mechanisms by which gut microbes influence immune function during HIV-1 infection. In this chapter, in vitro studies demonstrated that higher levels of butyrate inhibited pathobiont-driven LP T cell activation and HIV-1 infection of CD4 T cells, whereas lower levels of butyrate lacked this strong immunomodulatory effect. Taken together, these in vitro observations suggest that a loss of colonic mucosal

BPB, with a presumptive decrease in mucosal butyrate levels, in conjunction with an increased abundance of pro-inflammatory, pathobiont commensal bacteria promotes unregulated inflammatory responses and mucosal HIV-1 replication, which could further drive epithelial barrier damage and microbial translocation.

Butyrate plays several critical roles in colonic epithelial (colonocyte) function including by acting as a metabolic energy source for colonocytes[264-267], by increasing tight junction-mediated barrier integrity [269] and by protecting colonocytes from cellular damage and stress [172, 270]. Further, it has been shown that by decreasing NF-κB activation in metabolically stressed colonic epithelial cell lines, butyrate subsequently prevented transcytosis of E. coli in an in vitro epithelial cell/bacteria co-culture model [265].

Although the causality of these findings cannot be inferred, they suggest that reduced levels of butyrate may be a contributing factor to HIV-1-associated epithelial barrier breakdown that leads to MT.

Butyrate impacts cellular responses via G-coupled Protein Receptor (GPR) signaling

(GPR41, GPR43, and GPR109a) as well as through histone deacetylase inhibition (HDACi),

[258, 271] with an increased dosage favoring HDACi activity [272]. Butyrate functions through these two pathways to decrease immune cell activation [204, 259], inhibit T cell proliferation [190, 273] and increase apoptosis of activated T cells [190, 192]. In this study, higher doses of butyrate were associated with the most significant inhibition of T cell activation and CD4 T cell infection. Therefore, the butyrate-dependent decrease in the production of T cell cytokines following exposure to bacteria could be driven by inhibition of

T cell activation/proliferation, increased the death of activated T cells or, most likely, a combination of both. Further examination of butyrate’s GPR signaling and HDACi activity and the downstream effects on LP T cell activation/proliferation and cell death are warranted. Nevertheless, the data outlined in this chapter suggest that strategies to increase butyrate levels could potentially improve barrier integrity, reduce mucosal T cell activation, and decrease MT. The extent to which the HIV-associated mucosal pathology is reversible through dietary or microbiome manipulations awaits future study.

CHAPTER IV

BUTYRATE DIFFERENTIALLY IMPACTS LP CD4 T CELL ACTIVATION AND

INFECTION IN VITRO IN A CONCENTRATION DEPENDENT MANNER3

Background HIV-1 disease is characterized by a state of chronic immune activation that is associated with poor clinical outcomes and increased morbidity and mortality even in the presence of effective viral suppression with antiretroviral therapy[274, 275]. One of the primary mediators of HIV-1 associated immune activation is the profound breakdown in intestinal homeostasis that occurs early and throughout the disease. This is characterized by high levels of HIV-1 infection and CD4 T cell depletion in the intestinal mucosa[36, 37,

40, 43], including the depletion of T helper (Th) 17 populations[37, 43] that support epithelial barrier integrity via IL-22 production and protection from pathogens[276]. In persons living with HIV-1 (PLWH), several studies have noted lower abundances of genera in the gut microbiome that contain known butyrate-producing bacteria such as Lachnospira [145, 146,

148], Coprococcus [103, 145, 146], Roseburia [145, 146, 148, 154] and Faecalibacterium

[145, 146] . The abundances of several of these bacterial genera inversely associated with markers of inflammation. For example, one study determined that the relative abundance of

Faecalibacterium in the stool of PLWH who were on antiretroviral therapy (ART) negatively associated with serum sCD14 levels[146]. We previously demonstrated that untreated

PLWH had a lower relative abundance of Roseburia intestinalis, a known BPB, than uninfected controls in colonic mucosal biopsies. Furthermore, R. intestinalis was inversely associated with blood CD4 T cell activation, markers of microbial translocation, circulating inflammatory cytokines and viral load[229].

3 The data presented in this chapter are in preparation for submission.

Several studies have demonstrated that exogenous butyrate decreases activation of human peripheral blood (PB) and immortalized T cells as well as murine splenic T cells through alterations in the expression of proteins involved in activation, cell cycle regulation and cell death[190-192, 214]. In some studies, butyrate has also been shown to promote the differentiation of Th1 and Th17 cells from naïve human peripheral blood and murine splenic

CD4 T cells [188, 196]. Furthermore, butyrate decreased Th17-associated transcription factor activity and Th17-related gene expression in differentiating human Th17 cells but increased their expression in fully differentiated human Th17 cells[197]. Also, murine studies have highlighted that butyrate signaling through GPR43, and its activity as a histone deacetylase inhibitor (HDACi) are essential drivers of T regulatory cell differentiation[188,

194-196, 215]. Thus, the impact of butyrate on CD4 T cell function may be multifactorial depending on the type of T cells as well as their differentiation state.

In contrast to PB CD4 T cells, gut mucosal CD4 T cells are primarily resident effector memory cells that display different phenotypic and functional characteristics[80-82]. Given their proximity and consistent exposure to higher concentrations of butyrate, these cells may be more primed to respond to butyrate compared to the CD4 T cell populations in the periphery. Thus, information about how PB T cells responded to butyrate may not directly translate to what occurs in the gut mucosa. We previously demonstrated that the addition of high doses of exogenous butyrate decreased gut CD4 T cell activation by exposure to enteric bacteria and reduced levels of CD4 T cell HIV-1 infection in vitro [229]; however, the mechanisms by which butyrate modulated those responses have not presently been investigated. In this chapter, I utilized an in vitro model to comprehensively evaluate the impact of a range of physiologically relevant concentrations of butyrate on T cell receptor

(TCR) driven LP CD4 T cell activation and HIV-1 infection and to determine the mechanisms by which this occurs. I previously used this model to investigate the impact of butyrate on enteric pathobiont driven T cell activation. In the studies presented in this chapter, I used

TCR stimulation to study direct T cell activation. Comprehensively, these studies offer greater insight into how lower BPB and thus likely lower butyrate tissue concentrations may impact gut mucosal CD4 T cell function in the setting of HIV-1 infection in vivo.

Results In vitro Exposure to Exogenous Butyrate Decreases TCR-mediated LP CD4 T Cell

Activation and Proliferation Directly in a Concentration-dependent Manner.

TCR stimulatory beads were used to directly stimulate LP CD 4 T cells, to mimic the process of antigen presentation that would occur during microbial translocation. To determine the ability of butyrate to modulate LP CD4 T cell activation and proliferation in the setting of TCR-mediated activation, total LPMC were stimulated with or without TCR- stimulatory beads directed against CD3, CD2 and CD28, in the presence of a physiologic concentration range of butyrate (0.0625-0.5mM). The concentration range used was determined based on reported measured concentrations of butyrate in the gut lumen and portal circulation of human subjects [166, 167] and had limited impact on overall cell viability

(Fig. 4.1). The ability of butyrate to modulate LP CD4 T cell proliferation as measured by

CFSE dilution and T cell activation assessed by CD25 expression and HLA-DR and CD38 co-expression (markers associated with T cell activation and HIV-1 disease progression

[51]) were measured by multi-color flow cytometry. TCR-mediated stimulation of LPMC, in the absence of butyrate, induced significant increases in the percentage of LP CD4 T cells that had proliferated and expressed CD25 and co-expressed HLA-DR and CD38 (Fig 4.2a).

Increasing concentrations of butyrate led to a decrease in the percentage of proliferated LP

CD4 T cells in a dose-dependent manner (Fig. 4.2a). Butyrate also decreased the percentage of LP CD4 T cells that expressed CD25 and co-expressed HLA-DR and CD38 in a concentration-dependent manner with significant decreases observed at levels ≥0.25mM,

(Fig. 4.2a). In the absence of exogenous stimulation, the addition of butyrate to total LPMC did not impact proliferation or HLA-DR and CD38 co-expression on LP CD4 T cells;

56 however, butyrate increased the expression of CD25 on LP CD4 T cells by 1.4-fold at concentrations of ≥0.125mM. Given that butyrate is known to promote the differentiation of

Foxp3+ T regulatory cells[194, 195, 215], which also express high levels of CD25 [277], expression of Foxp3 by LP CD4 T cells in response to butyrate alone was measured. The addition of butyrate to LPMC cultures without exogenous stimulation did not result in increased percentages of Foxp3-expressing LP CD4 T cells after four days of culture (Fig

4.3).

To determine if the impact of butyrate on TCR-mediated proliferation and activation of LP CD4 T cells in LPMC occurred directly on T cells or required signals from non-T cells, negatively selected LP CD4 T cells were exposed to butyrate in the presence and absence of TCR stimulation. TCR bead stimulation of purified LP CD4 T cells in the absence of butyrate resulted in significantly increased percentages of proliferated LP CD4 T cells and those that expressed CD25 and co-expressed HLA-DR and CD38 (Fig 4.2b). Butyrate directly reduced TCR-mediated LP CD4 T cell proliferation and CD25 expression on LP CD4

T cells in a dose-dependent manner similar to that observed in LPMC cultures (Fig 4.2b).

Butyrate had a minimal direct effect on co-expression of HLA-DR and CD38 on purified CD4

T cells relative to that seen in LPMC cultures. Co-expression of HLA-DR and CD38 on LP

CD4 T cells were decreased in the presence of butyrate 1.39-fold compared to 1.72-fold in cultures with total LPMC at 0.5mM (Fig 4.2b). In the absence of TCR stimulation, the addition of butyrate directly increased the expression of CD25 on LP CD4 T cells in a dose- dependent manner (Fig 4.2b).

Timing of Butyrate Exposure and TCR Stimulation is Important for the Inhibition of LP

CD4 T Cell Proliferation.

To determine if the timing of butyrate exposure was critical to butyrate’s immunomodulatory activity on LP CD4 T cell proliferation, butyrate was added at various times before (pre-treatment) and after (post-treatment) the addition of the TCR-stimulatory

Figure 4.1. Total LPMC viability after exposure to butyrate LPMC were exposed to butyrate (0.0625mM - 4mM) for four days and total LPMC viability was determined by Trypan Blue and enumerated on a TC-120 cell counter (n=3).

58 beads to total LPMC cultures (Fig. 4.2c, 4.2d). For these assays, butyrate was added at

0.5mM, the dose at which the maximum immunomodulatory effect on LP T cell proliferation was observed (Fig. 4.2a). Exposure of LPMC to butyrate for ≤18hrs before TCR-mediated stimulation resulted in equivalent levels of LP CD4 T cell proliferation as observed when butyrate was added simultaneously (0hrs) with TCR stimulatory beads (Fig 4.2c). For the post-treatment assays, the addition of butyrate 6hrs after TCR-induced stimulation decreased proliferation to similar levels as those when butyrate was added simultaneously with TCR stimulation (Fig. 4.2d). However, the addition of butyrate at or after 18hrs of TCR stimulation limited butyrate inhibition of LP CD4 T cell proliferation, although the levels of proliferation were still lower than in the absence of butyrate (Fig 4.2d).

Higher Concentrations of Propionate and Acetate Relative to Butyrate were Required to Similarly Inhibit LP CD4 T Cell Activation and Proliferation.

To understand if butyrate’s immunomodulatory effects on TCR-mediated LP CD4 T proliferation and activation were shared by other SCFAs found in the gut lumen, total LPMC were similarly exposed to varying concentrations of acetate and propionate [166, 167].

Acetate and propionate decreased LP CD4 T cell proliferation and activation (percentages of CD25+ and HLA-DR+ CD38+ co-expressing LP CD4 T cells) in a concentration-dependent manner albeit at higher concentrations than butyrate (Fig 4.4a-c). Unlike butyrate, acetate and propionate had no impact on the frequency of CD25+ LP CD4 T cells in the absence of

TCR stimulation. However, like butyrate, acetate and propionate had minimal effect on LP

CD4 T cell activation (frequency of HLA-DR and CD38 co-expressing) and proliferation in the absence of TCR stimulation(data not shown).

Butyrate’s Inhibition of LP CD4 T Cell Activation and Proliferation is likely Mediated by HDAC Inhibition

To determine if butyrate, a known class I and II HDACi [180, 278], inhibited TCR- mediated LP CD4 T cell proliferation and activation via inhibition of HDACs, levels of

Figure 4.2. Butyrate reduces LP T cell activation and proliferation in a dose- dependent manner

Figure 4.2. Butyrate reduces LP T cell activation and proliferation in a dose- dependent manner Lamina propria mononuclear cells (LPMC) were cultured with or without TCR stimulating beads and butyrate (0.0625-0.5mM) for four days and levels of T cell activation and dim proliferation determined (n=3). Changes in the percentages of CFSE LP CD4 T cells and those expressing CD25 and co-expressing CD38 and HLA-DR were assessed from cultures with total LPMC (A) and purified LP CD4 T cells (B) using multi-color flow cytometry. FM2 (Fluorescence minus two) and isotype control values have been subtracted. Determination of the importance of the timing of butyrate addition to TCR bead activated cultures was dim carried out on total LPMC cultures and levels of CFSE cells were enumerated by flow cytometry after 4-day cultures. For the assays examining the importance of butyrate pre- treatment, LPMC were incubated with butyrate for 0, 2, 6 and 18 hours before the addition of TCR activating beads and cultured for four days post activation(C). For the assays examining the importance of butyrate post-treatment, LPMC were activated with TCR activating beads and then incubated with butyrate for 0, 6, 18 and 36 hours for the remainder of the 4-day incubation(n=3) (D). Statistical significance was determined using one-way ANOVA with Dunnett’s multiple comparison tests to 0mM butyrate or 0hr timepoint. Data represented as means, bars represent Standard Error of the Mean (SEM).

Figure 4.3. Examination of FOXP3+ CD25+ LP CD4 T cells after the addition of butyrate. LPMC were cultured with butyrate for four days and frequency of a.) CD25+ and b.) FOXP3+CD25+ LP CD4 T cells was determined(n=3) by multicolor flow cytometry. For CD4 Foxp3 staining the following gating strategy was used: viable, lymphocytes, singlets, CD3+, CD4+ and CD25+Foxp3+. Isotypes were used, and isotype values were removed from the final reported data. Data are mean values, and statistical significance was determined using either paired T-tests(a.) or one-way ANOVA with Dunnett’s multiple comparison tests to 0mM butyrate (b.). P value * <0.05 .Bars represent Standard Error of the Mean (SEM).

Figure 4.4. Propionate and Acetate both Decrease TCR Bead Driven LP CD4 T Cell Activation and Proliferation. LPMC were cultured with or without TCR stimulating beads and butyrate (0.5mM), acetate (0.25-10mM) or propionate (0.25-2mM) for four days and levels of T cell activation and proliferation were determined (n=3). Changes in the percentages of CFSEdim LP CD4 T cells and those expressing CD25 and co-expressing CD38 and HLA-DR were assessed from cultures with total LPMC using multi-color flow cytometry. FM2(HLA-DR and CD25) and isotype control values have been subtracted. Data are mean values and statistical significance was determined using one-way ANOVA with Dunnett’s multiple comparison tests to 0mM butyrate. Bars represent Standard Error of the Mean (SEM).

63 acetylation of histone 3 lysine 9 (H3K9), an indirect indicator of HDAC inhibition[179], were measured in LP CD4 T cells by multicolor flow cytometry. Twenty-four-hour exposure of unstimulated and TCR-stimulated LPMC to butyrate (0.5mM) resulted in significant increases in histone acetylation in LP CD4 T cells relative to controls without butyrate (Fig

4.5a). Butyrate-induced histone acetylation levels to similar extents in both TCR-stimulated and no stimulation conditions. Butyrate-mediated increases in histone acetylation in LP

CD4 T cells in the absence of activation were confirmed by western blot (Fig 4.6b).

The ability of a panel of known class I specific HDAC inhibitors (apicidin and MGCD) and pan HDAC inhibitors (TSA and ITF2357) to phenocopy butyrate’s effect on LP CD4 T cell proliferation was next investigated. Initial studies were carried out to determine the maximal dose of each HDAC inhibitor that did not induce toxicity (data not shown), and these maximal doses were used in subsequent assays. The addition of apicidin and MGCD decreased LP CD4 T cell proliferation to a greater degree than the addition of either pan

HDAC inhibitor (Fig 4.5b). The impact of apicidin on TCR mediated LP CD4 T cell activation and proliferation were next tested across a dose range (25-100nM) (Fig 4.5c). Apicidin decreased LP CD4 T cell proliferation and the expression of CD25 and co-expression of

HLA-DR and CD38 on LP CD4 T cells in a dose-dependent manner similar to butyrate (Fig

4.5c). In the absence of TCR stimulation, apicidin had little impact on LP CD4 T proliferation and the percentages of LP CD4 T cells expressing HLA-DR and CD38. However, similar to the observations of CD25 expression in the presence of butyrate, the percentage of CD25+

LP CD4 T cells increased on average 1.5-fold at all concentrations of apicidin tested (Fig

4.5c). To determine if apicidin also phenocopied butyrate’s ability to act directly on LP CD4

T cells, the maximal dose of apicidin (100nM) was added to purified LP CD4 T cells in the presence of TCR-stimulatory beads (Fig. 4.5d). Apicidin decreased LP CD4 T cell proliferation to a similar degree as observed with the highest dose of butyrate (0.5mM)

Figure 4.5 Butyrate’s HDACi Activity likely Plays a Role in its Immunomodulatory Effect on Activated LP CD4 T Cells.

Figure 4.5 Butyrate’s HDACi Activity likely Plays a Role in its Immunomodulatory Effect on Activated LP CD4 T Cells. LPMC cultured with or without TCR stimulating beads and butyrate (0.5mM) for 24 hours and levels of histone H3 lysine residue 9 acetylation levels were measured by intracellular flow cytometry(n=3). Butyrate driven changes in the mean fluorescence intensity of acH3K9 were compared to media controls using paired T-tests to determine significance(A). A panel of known HDACi was assayed for the ability to decrease LP CD4 T cell proliferation (CFSEdim) in total LPMC cultures stimulated with TCR stimulating beads, statistical significance was determined by comparing treatment condition to media control (B). Dose- dependent responses of apicidin (25-100nM) were assayed in total LPMC activated with TCR beads and percentages of CFSEdim LP CD4 T cells and those expressing CD25 and co-expressing CD38 and HLA-DR were assessed using multi-color flow cytometry(n=3) (C). FM2(HLA-DR and CD25) and isotype control values have been subtracted. Purified LP CD4 T cells were exposed to butyrate 0.5mM, apicidin 100nM or media control in the setting of TCR stimulation and the percentages of CFSEdim LP CD4 T cells and those expressing CD25 and co-expressing CD38 and HLA-DR were assessed using multi-color flow cytometry(n=4) (D). FM2 (HLA-DR and CD25) and isotype control values have been subtracted. Statistical significance was determined using raw data values and comparison of treatment condition to media controls by one-way ANOVA with Dunnett’s multiple comparison tests to 0 nM apicidin or paired T-tests. Data represented as means, bars represent Standard Error of the Mean (SEM) (*P<0.05, **P<0.01, ***P<0.001 ).

Figure 4.6. Butyrate Increases LP CD4 T Cell Histone Acetylation Levels (acH3K9) Purified LP CD4 T cells were exposed to exogenous butyrate for 24 hours, and H3K9 levels were evaluated by western blot at 0.125- and 0.5-mM butyrate. P (n=3). OD values were determined using Image J software comparing acH3 and b-actin expression in each sample.

(Fig. 4.5d). Similarly, apicidin also decreased the percentage of LP CD4 T cells expressing

CD25 and co-expressing HLA-DR and CD38 (Fig 4.5d).

Known Butyrate Receptors, GPR109a and GPR43, are Minimally Expressed on LP

CD4 T Cells.

To determine if GPR signaling might contribute to the actions of butyrate on LP CD4

T cell activation and proliferation, surface protein expression of GPR109a and GPR43 on LP

CD4 T cells directly ex vivo (baseline) and after TCR bead stimulation and butyrate exposure (4 days) was evaluated by multi-color flow cytometry (Fig 4.7a). Low percentages of LP CD4 T cells expressed GPR109a (mean 0.5% of total LP CD4 T cells ±0.01% SEM) and GPR43 (1.0% ± 0.6% SEM) at baseline (Fig 4.7b). TCR-bead stimulation of LPMC significantly increased the percentages of GPR43-expressing LP CD4 T cells, although these percentages remained low (<5% of total LP CD4 T cells). The addition of butyrate did not further alter expression (Fig 4.7c). Percentages of GPR109a+ LP CD4 T cells also increased with TCR stimulation. Similar to GPR43, GPR109a-expressing LP CD4 T cells constituted a very small fraction of total LP CD4 T cells (Fig. 4.7c). Percentages of

GPR109a were significantly decreased in the presence of butyrate relative to no butyrate, in both TCR-stimulated and unstimulated conditions (Fig 4.7c).

LP Th17 Cell Proliferation is Decreased at Lower Concentrations of Butyrate Relative to other Th Subsets.

The impact of butyrate on the production of T helper cytokines from purified LP CD4

T cells was next assayed. Butyrate decreased IL-17 production from TCR stimulated LP

CD4 T cells in a dose-dependent manner. IFNγ and IL-2 production were similarly decreased by butyrate with the greatest effect at 0.5mM (Figure 4.8a). The impact of butyrate on the proliferative capacity of Th cell subsets( Th1, Th17, and Th22) was next assessed by multicolor flow cytometry. TCR stimulation (in the absence of butyrate) drove significant increases in the proliferation of all gut Th cells enumerated relative to conditions

Figure 4.7. LP CD4 T Cells Express Low Levels of GPR43 and GPR109a Example flow stain profiled for GPR43 and GPR109a on LP CD4 T cells. Conditions represented are baseline expression and cells cultured for four days in the following conditions Media, Media + TCR stimulating beads, Butyrate (0.5mM) and butyrate (0.5mM) + TCR stimulating beads (A). Summary data of GPR43 and GPR109a on LP CD4 T cells on baseline cells (B) and cultured cells. Data are mean values, and statistical significance was determined using paired t-tests. Bars represent Standard Error of the Mean (SEM) (*P<0.05).

69 with no stimulation. (Fig 4.8b). The addition of butyrate significantly decreased the proliferation of all Th subsets at concentrations ≥0.125mM (Fig 4.8b). To determine if there were differences in the concentrations at which butyrate inhibited the proliferation of each Th subset by 50%, the half maximal inhibitory concentration (IC50) values were calculated.

Th17 cells had the lowest IC50 (0.140mM) while Th1 and Th22 subsets had similar IC50s

(0.227 and 0.270mM). The IC50 for cells that were IL-17-IL-22-IFNγ- was beyond the tested concentrations of butyrate (0.802mM). (Fig 4.8b).

To determine if the lower IC50 for butyrate seen in gut Th17 cells is specific to the gut or reflected a more general response of both tissue and peripheral Th cells, the impact of butyrate on TCR-mediated proliferation of Th cells in PB mononuclear cells (PBMC) was tested (Fig 4.9). Butyrate significantly decreased the proliferation of all Th subsets at concentrations ≥0.125mM. However, in contrast to LP Th subsets, butyrate inhibited PB

Th17 (IC50: 0.167) and Th1 (IC50: 0.178mM) cells to a similar extent as gut Th17 cells, while Th22 (IC50: 0.260mM) was more similar to the IC50s of the Th1 and Th22 cells in the gut. Unlike in the gut, PB IL-17-IL-22-IFNγ- cells were the most sensitive to butyrate (IC50:

0.127)( Fig 4.9).

Butyrate Enhances HIV-1 Infection of LP CD4 T Cells at Lower Concentrations.

We previously reported that high concentrations of butyrate (2mM) decreased productive HIV-1 infection of LP CD4 T cells driven by exposure to bacterial stimulation

[229] at concentrations that maximally inhibited LP CD4 T cell activation. Here I tested butyrate over a wider physiologic concentration range and with direct stimulation of LP CD4

T cells with TCR stimulation. The percentage of productively infected CD4 T cells was determined by the examination of LP CD4 T cells that, expressed the HIV-1 viral capsid protein (p24) by multicolor flow cytometry. Exposure of LPMC to TCR-beads significantly increased the percentage of productively infected CD4 T cells in 4-day cultures (Fig 4.10).

The addition of low doses of butyrate (≤0.125mM) in the presence of TCR stimulation

Figure 4.8. Butyrate Decreases T Helper Cytokine Production and the Proliferation of Gut T Helper Cells Purified LP CD4 T cells were cultured with or without TCR stimulating beads and butyrate (0.125-0.5mM) for four days and levels of secreted IL-17, IFNγ and IL-2 in the supernatants were quantified by ELISA(n=3) (A). Butyrate’s impact on TCR bead driven T helper subset proliferation was characterized in total LPMC exposed to butyrate (0.0625-0.5mM) for four days. Proliferation was determined by CFSE labeling and T helper subsets were identified by cytokine (IFNγ, IL-17 and IL-22) production after exposure to PMA and ionomycin stimulation for the last 4 hours of culture(n=6)(B). FMO and isotype control values have been subtracted. IC50 values were calculated, and statistical significance was determined using one-way ANOVA(*P<0.05) and was computed using raw data values and comparison of treatment condition to media controls. Data represented as means, bars represent Standard Error of the Mean (SEM).

Figure 4.9. Butyrate Reduces PB T Cell Activation and Proliferation in a Dose- dependent Manner. Butyrate’s impact on TCR bead driven T helper subset proliferation was characterized in total PBMC exposed to butyrate (0.0625-0.5mM) for four days. Proliferation was determined by CFSE labeling, and T helper subsets were identified by cytokine (IFNγ, IL-17, and IL-22) production after exposure to PMA and ionomycin stimulation + Golgi plug for the last 4 hours of culture. Isotype control values have been subtracted. IC50 values were calculated, and statistical significance was determined using one-way ANOVA and Dunnett’s multiple comparison tests to 0mM. Data represent mean CFSEdim frequencies (n=6).

72 resulted in a further increase in the percentage of productively infected LP CD4 T cells.

Conversely, higher butyrate concentrations (≥0.25mM) led to reductions in the percentage of productively infected LP CD4 T cells with 0.5mM butyrate suppressing infection to baseline levels (Fig 4.10a). To measure the effect of butyrate on the release of infectious virions throughout the 4-day culture period, infectious virions in the supernatants were next quantified. In agreement with percentages of p24-expressing LP CD4 T cells in the setting of

TCR-stimulation (Fig 4.10a), increased infectious HIV-1 titers were observed at lower concentrations of butyrate whereas reductions at 0.5mM to levels seen in the absence of

TCR-stimulation were noted (Fig .4). In the absence of TCR-stimulation, there was an observed increase in the production of infectious virions at 0.125mM butyrate (Fig 4.10b) which was not reflected in the percentages of p24+ LP CD4 T cells (Fig 4.10a). To determine if butyrate was acting directly on LP CD4 T cells to influence infection levels, assays using purified LP CD4 T were carried out. TCR-stimulation in the absence of butyrate increased percentages of p24-expressing LP CD4 T cells (Fig 4.10c) and infectious

HIV-1 titers (Fig 4.10d). In contrast to total LPMC cultures (Fig 4.10a, b), low dose (≤

0.125mM) butyrate did not increase the percentage of p24-expressing purified CD4 T cells or infectious HIV-1 titers (Fig 4.10c-d) with lower expression and titers observed at 0.125mM versus 0mM butyrate. Higher concentrations of butyrate (≥0.25mM) decreased p24+ LP

CD4 T cells and infectious virus titers (Fig 4.10 c-d).

The percentage of productively infected LP Th cell subsets in the presence of butyrate was next analyzed. LPMC activation promoted similar increases in the percentage of p24-expressing Th17, Th1 and Th22 above conditions without stimulation (Fig 4.10e).

Butyrate at low doses increased infection of Th17, Th1 and Th22 cells with significance reached at 0.0625mM for Th22 and 0.125mM for Th17 cells. Higher concentrations of butyrate decreased percentages of all p24-expressing Th cell subsets relative to no butyrate

(Fig 4.10e). The IL-17- IL-22-IFNg- LP CD4 T cells were infected at lower percentages than

Figure 4.10. High and Low Concentrations of Butyrate Differentially Alter LP CD4 T Cell HIV-1 Infection Levels.

Figure 4.10. High and Low Concentrations of Butyrate Differentially Alter LP CD4 T Cell HIV-1 Infection Levels. LPMC were spinoculated with the CCR5-tropic transmitted/founder virus CH40 and cultured with or without TCR stimulating beads and butyrate (0.0625- 0.5mM) for four days and levels of LP CD4 T cell productive infection(p24) (n=6) were assessed using multi-color flow cytometry (A). Mock-infected control values have been subtracted. HIV-1 infectious viral titers in culture supernatants were assayed by TZM-bl and values are reported as relative luciferase units/per ml (n=5)(B). To determine if butyrate directly impacts LP CD4 T cell infection, purified LP CD4 T cells were spinoculated with CCR5-tropic transmitted/founder virus CH40 and cultured with or without TCR stimulating beads and butyrate (0.0625- 0.5mM) for 4 days and levels of LP T CD4 T cell productive infection(p24) were quantified by multi-color flow cytometry(n=4)(C). HIV-1 infectious viral titers in culture supernatants from cultures with purified LP CD4 T cells were assayed by TZM-bl (D). Frequencies of productively infected T helper subsets were determined in total LPMC spinoculated with CH40, stimulated with TCR stimulating beads and exposed to a range of butyrate (0.0625- 0.5mM) and cultured for four days. T helper subsets were determined by intracellular cytokine staining after addition of PMA and ionomycin + Golgi plug during the last 4 hours of culture (n=6)(E). Statistical significance was determined using one-way ANOVA with Dunnett’s multiple comparison tests and was computed using raw data values and comparison of treatment condition to 0mM controls. Data represented as means, bars represent Standard Error of the Mean (SEM).

Figure 4.11. Butyrate does not Alter the Frequencies of Productively Infected T Helper Subsets in the Absence of Activation. Frequencies of productively infected T helper subsets were determined in total LPMC spinoculated with CH40, stimulated with TCR beads and exposed to a range of butyrate (0.0625-0.5mM) and cultured for four days. T helper subsets were determined by intracellular cytokine staining after addition of PMA/Ionomycin + Golgi plug during the last 4 hours of culture. (n=6) Bars represent Standard Error of the Mean (SEM).

76 the other Th subsets in conditions with TCR stimulation, but like other subsets, the frequency of productively infected cells increased at 0.125mM. In the absence of stimulation, butyrate alone did not substantially alter the frequency of p24+ Th cells (Fig

4.11).

Low Dose Butyrate Increases the Expression of CCR5 on Activated LP CD4 T Cells and HIV-1 Viral Entry. Expression levels of known HIV-1 co-receptors are known to alter the infectibility of

CD4 T cells[14, 40, 279].To investigate if increased LP CD4 T cell infection observed following low dose butyrate exposure was related to increased expression of the canonical

HIV-1 co-receptor CCR5 and CD4 [280], the impact of butyrate on surface expression of these molecules was determined using multi-color flow cytometry. TCR stimulation of total

LPMC in the absence of butyrate significantly increased CD4 expression(Fig 4.12a).

Butyrate decreased CD4 expression at concentrations ≥0.125mM (Fig 4.12a). However, the addition of butyrate to total LPMC in the absence of exogenous TCR stimulation did not significantly impact on CD4 expression. Conversely, butyrate increased CCR5 expression at lower concentrations (0.125mM) in the presence of TCR stimulation, on average by 1.42- fold (Fig 4.12a). In the absence of TCR stimulation, CCR5 expression increased on average by 1.29-fold at 0.125mM but did not reach statistical significance (Fig 4.12a).

To determine if the effect of low dose butyrate on CCR5 expression was via a direct effect on butyrate on these cells we next tested the ability of butyrate, at concentrations that had the most differential response on HIV-1 infection and CCR5 expression, for the ability to alter CCR5 and CD4 expression on purified LP CD4 T cells. In keeping with observations for total LPMC, TCR stimulation alone of purified CD4 T cells resulted in significant increases in

CD4 expression relative to no stimulation but did not significantly increase the expression of

CCR5 (Fig 4.12b). Similar to total LPMC, CD4 expression on purified CD4 cells was decreased by butyrate in a dose-dependent manner (Fig 4.12b). However, in contrast to

77 total LPMC, low dose butyrate (0.125mM) did not increase CCR5 and high dose (0.5mM) decreased CCR5 expression(Fig 4.12b).

To better understand the impact of butyrate on the HIV-1 lifecycle, both viral transcription (gag) and entry (HIV-1 RU-5) were next studied by qPCR. Butyrate is known to promote the transcription of HIV-1 virus in latently infected CD4 T cells[281]. Therefore HIV-

1 transcription (gag RNA) was measured in activated LP CD4 T cells exposed to butyrate.

The addition of butyrate resulted in a dose-dependent increase in HIV-1 gag RNA(Fig 4.12c)

HIV-1 RU-5 DNA represents the early DNA products of reverse transcription and can be used to quantify the amount of HIV-1 viral entry and reverse transcription [282]. Since the impact of butyrate on LP CD4 T cell infection and CCR5 expression was seen in the setting of other cells in the LPMC, we exposed HIV-1 infected LPMC to butyrate and TCR stimulation for 24 hours and then sorted the CD4 T cells. I next quantified HIV-1 RU-5 DNA levels in these purified LP CD4 T cells. RU-5 DNA levels increased at 0.125mM butyrate, while there were no observable effects at higher concentrations (Fig 4.12d).

Apicidin Decreases LP CD4 T Cell Productive Infection and CD4 Expression

To evaluate if butyrate’s effect on LP CD4 T cell infection was mediated via HDAC inhibition, the impact of apicidin on LP CD4 T cell HIV-1 infection and HIV-1 co-receptor expression was next evaluated (Fig 4.13). In keeping with butyrate’s ability to decrease productive infection at higher concentrations, apicidin in the setting of TCR stimulation reduced the percentage of p24+ LP CD4 T cells at concentrations ≥ 25mM (Figure 4.13).

The addition of apicidin in the setting of TCR-stimulation did not drive changes in CCR5 expression, however similar to butyrate at higher concentrations it did decrease CD4 expression on average -1.3-fold at 100nM (Fig 4.13).

Figure 4.12 Butyrate Alters HIV-1 Co-receptor Expression on LP CD4 T Cells and Viral Entry

Figure 4.12 Butyrate Alters HIV-1 Co-receptor Expression on LP CD4 T Cells and Viral Entry LPMC were cultured with or without TCR stimulating beads and butyrate (0.0625-0.5mM) for four days, and the levels of CCR5 and CD4 expression (MFI) were assessed using multi- color flow cytometry (N=6). Isotype control values have been subtracted for CCR5(A). Purified LP CD4 T cells were similarly stimulated with TCR stimulating beads and exposed to butyrate (0.125 and 0.5mM), and CCR5 and CD4 expression were determined(B). Data is displayed as raw MFI or frequency values. (C) Viral entry assay and reverse transcription, LPMC were infected for 24 hour, and the presence of HIV-1 RU-5 DNA was quantified in LP CD4 T cells. HIV-1 RU-5 DNA copy number was normalized to RNaseP cycle number (N=4). (D) HIV-1 gag RNA PCR assay. LPMC were spinoculated with CH40 HIV-1, exposed to increasing concentrations of butyrate, amprenavir or raltegravir and TCR stimulation for 48 hours. After 48 hours, LP CD4 T cells were isolated, and qPCR for HIV-1 gag RNA and GAPDH were carried out. Statistical significance was determined using one-way ANOVA with Dunnett’s multiple comparison tests to 0mM butyrate. Data represented as means, bars represent Standard Error of the Mean (SEM).

Figure 4.13. Apicidin Decreases LP CD4 T Cell Productive Infection and CD4 Expression at High Concentrations. A.) CH40 infected LPMC were cultured with TCR stimulating beads and apicidin (25- 100nM) for four days, and levels of productive infection (p24)were assessed using multi- color flow cytometry. Mock control values have been subtracted (n=3). B.) Total LPMC were activated with TCR stimulating beads and exposed to apicidin for four days (25-100nM), and CCR5 and CD4 expression were determined by MFI. CCR5 isotype expression values were removed from data. Data are mean values, and statistical significance was determined using one-way ANOVA with Dunnett’s multiple comparison tests(*P<0.05) relative to 0nM apicidin controls. Bars represent Standard Error of the Mean (SEM).

Discussion

Low abundances of BPB have been reported in the gut mucosa and stool of

PLWH[103, 145, 146, 229, 283]. However, studies extensively probing how low tissue concentrations of butyrate may impact local gut immunity, particularly gut CD4 T cells, in the setting of HIV-1 infection and the mechanisms by which this occurs, are limited. In this chapter, I utilized an in vitro human LPMC model to investigate the impact of butyrate on LP

CD4 T cell function both in the presence and absence of HIV-1. The responses to various concentrations of butyrate that would broadly reflect “low” and “high” physiologic butyrate tissue concentrations were used to investigate the impact of butyrate in this model. Butyrate directly and in a concentration-dependent manner decreased TCR mediated LP CD4 T cell activation and proliferation likely by acting as an HDAC inhibitor. Furthermore, while butyrate decreased the proliferation of all gut Th subsets enumerated there was an increased sensitivity to butyrate’s anti-proliferative effect in the Th17 subset which was not reflected in

PB Th subsets highlighting the gut-specificity of butyrate’s actions. A caveat to these findings is that the more sensitive subsets in both the LP and PB Th cells were the subsets that had the lowest average proliferation in response to TCR stimulation. This may indicate that LP T helper cells that are less responsive to TCR stimulation are more sensitive to butyrate. These findings have broad applicability to other diseases such as irritable bowel disease (IBD) that are characterized by low abundances of BPB and marked activation of

Th1 and Th17 cells in the gut mucosa[135, 136, 284]. In these settings, higher concentrations of butyrate would likely dampen Th17 and Th1 activation and proliferation in the gut mucosa thus limiting the overt T cell driven inflammation that is characteristic of these diseases.

A key finding of in this chapter is the apparent uncoupling of HIV-1 infection and LP

CD4 T cell activation at lower concentrations of butyrate whereby low dose butyrate decreased LP CD4 T cell activation but increased productive infection of LP CD4 T cells

82 which was reflected in increased infection of all Th subsets. Although CD4 T cell activation is generally considered to be tightly associated with HIV-1 infection both in vitro and in vivo

[285-288], we had previously demonstrated enteric bacteria-driven increases in LP CD4 T cell infection in vitro also occurred in the absence of extensive LP CD4 T cell activation[156].

In this setting, the increased infection was associated with increased CCR5 expression on

LP CD4 T cells in response to signals from other LPMC. Intriguingly, increased infection at low doses of butyrate was also similarly linked to increased CCR5 expression and viral entry, but this effect was lost in the absence of other LPMC suggesting increased CCR5 expression was mediated via butyrate’s effect on other cell types in the LPMC. Therefore, the regulation of cytokines produced by these cells could be a potential mechanism by which butyrate alters CCR5 expression indirectly. Butyrate has been shown to induce IL-10 production by antigen presenting cells(APCs) in vitro [225, 263] and other studies have implicated IL-10 in increasing human PB CD4 T cell CCR5 expression[289]. Taken together, these studies suggest a similar cross-talk between low dose butyrate, APCs and T cells could be driving increased CCR5 expression on LP CD4 T cells at lower concentrations of butyrate. Additionally, these studies also demonstrate that the increase in productively infected LP CD4 T cells is not reflective of an increase in viral transcription, which further affirms the observation the increase in productive HIV-1 infection by low concentrations of butyrate is CD4 T cell and HDAC inhibitor independent. Studies are underway to dissect the specific mechanisms by which low dose butyrate regulates factors that promote increased

CCR5 expression and HIV-1 viral entry into LP CD4 T cells.

Gut mucosal Th cells are essential mediators of intestinal barrier integrity, and dysregulation of their function and frequencies is a hallmark of inflammatory disorders of the gut mucosa[62]. For example, increased frequencies of cytokine-producing Th1 and Th17 cells in the gut mucosa are noted to be a key feature of IBD[284]. In this current study, butyrate decreased the proliferation of all Th cell subsets; however, the IC50 of Th17

83 proliferation occurred at lower doses of butyrate. There are several potential explanations for the sensitivity of Th17 cells to butyrate relative to other Th subsets in the gut mucosa.

Firstly, Th17 cells may have higher expression levels of know butyrate receptors such as

GPR43 and GPR109a. Indeed, although we observed the low frequency of GPR expression on total LP CD4 T cells, it is possible that this population of GPR-expressing

CD4 T cells represent the similarly small fraction of Th17 cells found in the LP.

Furthermore, CD147, a component of MCT-1, a known cellular butyrate transporter, was shown to be expressed on Th17 [184, 290]. Thus, increased sensitivity of LP Th17 cells may also reflect an increased ability to transport butyrate into these cells relative to other Th subsets.

Although this study primarily focused on butyrate’s ability to modulate LP CD4 T cell function in vitro, we also investigated functional responses after exposure to acetate and propionate, the other primary SCFAs found in the human gut [166, 167]. Both acetate and propionate decreased LP CD4 T cell activation and proliferation, albeit at higher concentrations than butyrate. This differential impact on LP T cell function may reflect the ability of these to inhibit class I HDAC activity since all 3 inhibit this class of HDACs, but butyrate is known to be the most potent. [85, 180, 181, 291]. Our observations also suggest that there may be redundancies in the ability of these SCFAs to modulate LP CD4 T cell responses in the gut mucosa. Thus, lower concentrations of one may have limited detrimental impact if higher concentrations of the other SCFAs are present.

While several studies have demonstrated butyrate’s ability to inhibit CD4 T cell responses when added concurrently with stimulation[190-192, 195, 196, 229], the ability of butyrate to alter existing LP CD4 T cell activation has yet to be thoroughly studied. We demonstrated that butyrate’s immunomodulatory effects are dependent on the addition of butyrate concurrently with LP CD4 T cell activation. The observations that butyrate’s immunomodulatory effects are lost if LP CD4 T cells are exposed to butyrate after 18hrs of

84 stimulation have significance in the clinical setting. Since the efficacy of interventions to increase butyrate may be diminished if T cell activation is well established and therefore in this setting butyrate would likely only prevent activation of newly recruited T cells to the gut mucosa. Indeed, this may explain the conflicting observations in butyrate intervention in IBD patients where some studies demonstrated clinical improvement while others showed no effect from the treatments [292-294].

In conclusion, this data outlined in this chapter is the first to demonstrate that butyrate can have differential impacts on gut human LP CD4 T cell activation and HIV-1 infection over a physiologic concentration range in vitro. To translate these findings into the clinical setting, future studies investigating the impact of butyrate administration during HIV-

1 infection on inflammation and T cells activation both in the mucosa and periphery need to be carried out.

CHAPTER V

ADDITIONAL MECHANISMS OF BUTYRATE’S IMMUNOMODULATORY EFFECTS ON

LP CD4 T CELL ACTIVATION AND PROLIFERATION

Background

Butyrate has been demonstrated to alter cell function through multiple pathways that include HDACi and signaling though GPRs and PPARγ[178, 185, 205, 207]. In the previous chapter, I determined the importance of HDACi and GPR signaling in the immunomodulatory effect of butyrate on TCR driven activation of LP CD4 T cells. These studies indicated that low frequencies of LP CD4 T cells expressed GPR43 and GPR109a, which indicated that this was not likely the dominant pathway by which butyrate functioned in this cell type. Next, the role of HDAC inhibition was investigated, which demonstrated that butyrate increased histone acetylation levels in LP CD4 T cells and its effects on TCR stimulation driven LP CD4 T cell activation and proliferation were phenocopied by a class I

HDACi. While not conclusive, these data indicate that inhibition of class I HDAC proteins are likely the primary mechanism by which butyrate alter LP CD4 T cell activation and proliferation.

In this chapter, I attempted to further confirm the role of HDAC inhibition using a histone acetyltransferase inhibitor (HATi). A HATi was used to determine if the increases in histone acetylation driven by butyrate were responsible for the observed changes in TCR driven LP CD4 T cell proliferation. HAT proteins are the counterpart to HDAC proteins in the cell and act to add acetyl groups to histones or other proteins while HDACs remove them[179, 272]. When butyrate, a known HDACi, is present the activity of HDACs is deceased and thus the acetylation levels in the cell increase due to unopposed HAT activity.

Therefore, I tested the ability of a HATi to decrease histone acetylation in our model and block the effect of butyrate on LP CD4 T cell proliferation.

PPARγ receptor signaling is a relatively understudied pathway by which butyrate has been demonstrated to signal. To date, there have been two critical studies that have shown a role for butyrate signaling through the PPARγ receptor in colonic epithelial cells[207, 208].

The studies demonstrated that butyrate was able to modulate host epithelial gene expression and production of reactive nitrogen species through PPARγ signaling. While these studies focused exclusively on PPARγ signaling in epithelial cells, there is precedence for the immunomodulatory impact of PPARγ signaling in T cells[295, 296]. Therefore, in this chapter, I also probed the importance of PPARγ signaling on the effect of butyrate on TCR stimulated LP CD4 T cells through the use of both PPARγ agonists and antagonist in the

LPMC model.

Results

Inhibition of Histone Acetyltransferases does not Block the Antiproliferative Effect of

Butyrate on TCR Stimulated LP CD4 T Cells.

Increased histone acetylation is a marker for HDAC inhibition and is mediated by the activity of HAT proteins[178]. To determine if the inhibition of HAT protein activity with a chemical HATi decreased histone 3 lysine 9 (H3K9) acetylation in LP CD4 T cells, LPMC were cultured in the presence or absence of 0.5mM butyrate and the p300 HATi C646 for 24 hours. H3K9 acetylation levels were next determined by multicolor flow cytometry. The addition of butyrate to LPMC resulted in increased H3K9 acetylation in LP CD4 T cells.

When the HATi was added to cultures with butyrate, there was a consistent decrease in

H3K9 acetylation after 24 hours of culture (Figure 5.1A).

Next, I sought to determine if the addition of the HATi to LPMC cultures with TCR stimulation and butyrate blocked the effect of butyrate on proliferation as determined by

CFSE staining. LPMC were cultured in the presence or absence of TCR stimulation and

M e d ia 2 5 H A T i 5 u M

2 0

1 5

LP CD4 T1 cells) 0

Proliferation d i m 5

(CFSE 0 0 0.0625 0.125 0.25 0.5

Butyrate Conc (mM)

Figure 5.1. HAT Inhibition Decreases Butyrate Driven H3K9 Acetylation but does not Impact Butyrate’s Impact on LP CD4 T Cell Proliferation. a.) LPMC were cultured with TCR stimulating beads and butyrate (0.5mM) in the presence or absence of a HATi (5µM) for 24 hours, and levels productive H3K9 acetylation was assessed using multi-color flow cytometry (n=3). b.) Total LPMC were activated with TCR stimulating beads and exposed to butyrate (0-0.5mM) and HATi (5µM) for four days and proliferation of LP CD4 T cells was determined by CFSE dilution(n=3). Data are mean values. Bars represent Standard Error of the Mean (SEM).

0.5mM butyrate and the HATi for four days. The cultures with the HATi were pretreated with the HATi for one hour. After four days of culture, the addition of the HATi did not significantly reverse the impact of butyrate on TCR driven LP CD4 T cell proliferation(Figure 5.1B)

PPARγ Agonism Phenocopies the Antiproliferative Effect of Butyrate but is not

Reversed by a PPARγ Antagonist.

Since PPARγ is known to be expressed in CD4 T cells and PPARγ signaling has been demonstrated to alter CD4 T cell differentiation and function[295, 296], I next tested the impact of PPARγ agonism on TCR stimulation driven LP CD4 T cell proliferation. LPMC were stimulated with TCR activating beads in the presence of a dose range of

Rosiglitazone, a known PPARγ agonist, for four days. Rosiglitazone decreased LP CD4 T cell proliferation at the highest non-toxic concentration of 100µM but had little activity at lower concentrations(Figure 5.2A).

Next, a non-competitive PPARγ antagonist, T 0070907, was tested for the ability to block butyrate’s antiproliferative effect in LP CD4 T cells. LPMC were pre-treated with a dose range of T 0070907a for 1 hour and then cultured with TCR stimulating beads and butyrate (0.125 and 0.5mM) for four days at which time proliferation was determined by multicolor flow cytometry. After four days of culture, butyrate decreased TCR stimulation driven LP CD4 T cell proliferation, and T 0070907 was unable to inhibit butyrate’s impact.

An important caveat to this assay was that T 0070907 was unable to inhibit rosiglitazone’s impact at 100µM (Figure 5.2B).

Discussion

The data presented in this chapter attempted to further clarify the mechanisms by which butyrate is inhibiting TCR driven LP CD4 T cell proliferation. The data presented here continue to address the mechanisms by which butyrate alters LP CD4 T cell proliferation.

There are several important findings highlighted in this chapter. First, the p300 HAT inhibitor resulted in consistent decreases in H3K9 acetylation in LP CD4 T cells cultured with

Figure 5.2. PPARγ Agonism Decreases LP CD4 T Cell Proliferation, but PPARγ does not Inhibit this Effect. a.) LPMC were cultured with TCR stimulating beads and butyrate (0.5mM) as well as a concentration range of rosiglitazone (0.1-100µM) for four days and proliferation of LP CD4 T cells was determined by CFSE dilution using multi-color flow cytometry (n=1). b.) Total LPMC were activated with TCR stimulating beads and exposed to butyrate (0.125 and 0.5mM) and the PPARγ antagonist T 0070907 for four days and proliferation of LP CD4 T cells was determined by CFSE dilution (n=2). Bars represent Standard Error of the Mean (SEM).

90 butyrate. Also, there is a trend for the inhibition of butyrate’s effect on LP CD4 T cell proliferation by the HATi. Due to the small sample size, further assays are warranted to determine if the HATi is blocking butyrate’s antiproliferative impact on LP CD4 T cells.

Furthermore, these assays are confounded by the observation that p300 is involved in cellular proliferation and its inhibition can decrease cellular proliferation[297]. In our model, the use of a p300 HATi did modestly decrease the proliferative capacity of LP CD4 T cells independent of butyrate. Therefore, future studies focused on the impact of HAT inhibition on butyrate’s effect on LP CD4 T cell activation and infection may be informative.

The impact of PPARγ agonism on LP CD4 T cell proliferation was not unexpected since PPARγ agonism has been previously demonstrated to dampen the function and activation in both mouse and human PB CD4 T cells [295, 296]. While there did appear to be an impact of rosiglitazone at the highest concentration tested there are several limitations that must be addressed moving forward. First, the sample size must be increased to generate statistical differences and LP CD4 T cell activation and HIV-1 infection need to be evaluated in addition to proliferation. There also needs to be a broader concentration range of rosiglitazone tested centered around 100µM to determine if there are concentration- dependent effects of rosiglitazone. Lastly, the need for better positive controls for the

PPARγ antagonist due to the inability of the PPARγ antagonist to block the effect of butyrate and rosiglitazone on LP CD4 T cell proliferation. Collectively, the data presented in this chapter indicate that the mechanisms by which butyrate alter LP CD4 T cell activation and proliferation are likely multifactorial and more directed and specific assays are warranted to establish the mechanism or mechanisms by which butyrate alters LP CD4 T activity.

CHAPTER VI DISCUSSION

Project Rationale and Outline

Chronic immune activation, infection, and depletion of gut CD4 T cells and dysbiosis of the GI tract microbiome are hallmarks of HIV-1 infection and are thought to contribute to its progression[22, 100, 112, 274]. The lower relative abundance of BPB species has been demonstrated in the colonic mucosal microbiome of PLWH[229]. A key feature of these changes is the inverse association of the abundance of these groups of bacteria with indicators of local and systemic immune activation as well as microbial translocation and viral load[229]. The association of the altered abundance of BPB and gut mucosal inflammation has also been determined in other inflammatory conditions of the GI tract such as IBD[135-137]. The impact of alterations in butyrate concentrations on gut mucosa T cell activation and HIV-1 infectability has yet to be studied in detail.

The studies outlined in this thesis had two specific aims. The first aim of this thesis was to characterize the effect of exogenous butyrate on enteric pathobiont-driven gut mucosal T cell activation and HIV-1 infection. This was followed by the second aim that was to carry out mechanistic studies looking at the impact of a range of physiologic butyrate concentrations on direct TCR stimulation driven LP CD4 T cell activation and HIV-1 infection. For the first aim of the thesis, I hypothesized that butyrate would reduce enteric bacteria driven LP T cell activation, cytokine production, and HIV-1 infection. For the second aim of the thesis, I hypothesized that exposure of LP CD4 T cells to physiologic concentrations of butyrate would dampen LP CD4 T cell activation, cytokine production, Th subset proliferation, and HIV-1 infection directly and in a concentration-dependent manner through its known activity as an HDACi.

Since the detailed results have been presented in previous chapters, I will review the major findings from each chapter and discuss the implications of these findings on the understanding of HIV-1 pathogenesis and the potential of these findings to inform future studies. Furthermore, I also will discuss the limitations of the data presented in this thesis.

Summary of Major Findings

In the first aim of this thesis, I aimed to translate findings from the clinical study outlined in the introduction[229], by examining the impact of exogenous butyrate on enteric pathobiont driven LP T cell activation and HIV-1 using an in vitro model of microbial translocation. In these studies, the pathobiont Prevotella stercorea was used to drive LP

CD4 and CD8 T cell activation, cytokine production, and productive HIV-1 infection. When exogenous butyrate was added at a concentration of 2mM, there was a significant decrease in both LP CD4 and CD8 activation, production of inflammatory cytokines (IL-17 and IFNγ) and productive HIV-1 infection.

The purpose of the second aim was to address the impact of butyrate on LP CD4 T cell activation, proliferation, and HIV-1 infection and to understand the mechanism by which this occurs. In these initial studies, it was determined that butyrate decreased LP CD4 T cell activation and proliferation in a dose-dependent manner after four days of culture with the maximal effect seen at the highest concentration tested (0.5mM). In the absence of TCR stimulation butyrate increased the expression of CD25 on LP CD4 T cells at concentrations

≥0.125mM. Next, negatively selected LP CD4 T cells were tested similarly to determine if the effect of butyrate was mediated directly on an LP CD4 T cell or another cell type in the

LPMC. Similar to total LPMC, there was a dose-dependent decrease in TCR stimulation driven activation and proliferation as well as an increase in CD25 expression in the absence of TCR stimulation. While the concentration-dependent decrease in LP CD4 T cell activation and proliferation was expected the increase in the frequency of CD25+ LP CD4 T cells in the absence of activation was not. Since butyrate has been demonstrated to

93 promote the differentiation and function of T regulatory cells[194, 195, 215], which have higher expression levels of CD25[298], the frequency of FoxP3+ LP CD4 T cells was next tested. After four days of culture, butyrate had no observable effect on the frequency of LP

CD4 T cells that expressed FoxP3. Therefore, butyrate did not alter T regulatory cell frequency in our model, and the increase in CD25 expression may reflect the specific regulation of CD25 expression by butyrate.

Since butyrate acted directly and in a concentration-dependent manner, the subsequent assays that were carried out to address aim 2 examined the specificity of the effect of butyrate. First, the other primary SCFAs, acetate, and propionate were tested for their ability to decrease LP CD4 T cell activation and proliferation over a physiologic concentration range[166, 167]. Similar to butyrate, acetate and propionate decreased TCR stimulation driven LP CD4 T cell activation and proliferation in a concentration-dependent manner. However, the effects of these other SCFA were noted at higher concentrations than butyrate. These results indicate the effect of butyrate on LP CD4 T cell activation and proliferation are not unique and are shared by other SCFAs in the gut lumen. This observation may further inform on the mechanisms by which butyrate alters LP CD4 T cell responses since these SCFA share the ability to signal through common surface GPRs and function as HDAC inhibitors.

Butyrate is known to have multiple pathways by which it alters cell function[228]. Of those, the two better studied are HDAC inhibition and signaling through GPRs(GPR109a and GPR43)[228]. To explore whether either of these two pathways was involved in butyrate’s ability to alter LP CD4 T cell activation and proliferation, the ability of butyrate to function as an HDACi was examined. When LPMC were treated with butyrate for 24 hours, there was a significant increase in the amount of histone H3 lysine 9 acetylation as determined both by flow cytometry and western blot. With confirmation that butyrate was acting as an HDACi, the ability of known HDACis to phenocopy butyrate’s effect on LP CD4

T cell activation and proliferation was next examined. These studies demonstrated that class

I HDACis, notably apicidin, phenocopied butyrate’s ability to directly decrease TCR stimulation driven LP CD4 T cell activation and proliferation in a concentration-dependent manner. Collectively, these studies indicate that butyrate is likely functioning as an HDACi in

LP CD4 T cells and the inhibition of class I HDAC proteins is likely the mechanism by which butyrate is decreasing LP CD4 T cell activation and proliferation. Ideally, I would confirm these observations by blocking specific HDAC proteins either chemically or genetically.

Although butyrate was acting as an HDACi in our culture model, there was still a possibility that GPR signaling could be an essential component of butyrate’s immunomodulatory effect on LP CD4 T cells. To determine if GPR signaling was involved in butyrate’s activity in LP CD4 T cells, the expression of GPR43 and GPR109a protein was examined on LP CD4 T cells directly ex vivo and after four days of culture with or without butyrate and TCR stimulation. While there did appear to be some differences in the expression of both GPR43 and GPR109a on LP CD4 T cells after TCR stimulation and butyrate exposure, the frequency of LP CD4 T cells that expressed these GPRs was generally low (≤4% GPR43 and ≤1% GPR109a) and could not account for the magnitude of effects on T cell proliferation and activation. Although, butyrate signaling through GPRs is not likely responsible for the observed effect of butyrate on the majority of LP CD4 T cell it could impact a smaller subset of cells such as the Th17, Th22 or T regulatory cells. The importance of these surface receptors could be further interrogated by using chemical antagonist and agonist as well as genetic modulation of their expression.

The CD4 T cells that populate the gut mucosa are comprised of unique T helper subsets that serve important roles in gut barrier homeostasis[80, 299, 300]. During HIV-1 infection, gut Th17 and Th22 cells appear to be preferentially infected and depleted, and it is thought that the loss of these cells is central to HIV-1 pathogenesis[43, 102, 115]. With the importance of Th cells in HIV-1 disease progression, the impact of butyrate on the

95 proliferation of distinct Th subsets in the LPMC was next tested. Consistent with the findings in total LP CD4 T cells, butyrate decreased the proliferation of Th1, Th17, Th22 and non-

IFNγ, IL-17, IL-22 producing Th cells in a concentration-dependent manner. Of the subsets enumerated, the non-IFNγ, IL-17, IL-22 producing Th cells proliferated the most and Th17 cells proliferated the least in response to TCR stimulation. With the addition of butyrate, the

Th17 cells were the most sensitive while the non-IFNγ, IL-17, IL-22 producing Th cells were the least sensitive to the anti-proliferative effects of butyrate. Consistent with the higher relative sensitivity of LP Th17 cells to the antiproliferative effect of butyrate, the production of

IL-17 from purified LP CD4 T cells that had been stimulated with TCR beads was decreased at lower concentrations than both IL-2 and IFNγ. The observed sensitivity of LP Th17 cells to butyrate is an intriguing observation since it has previously been reported that human T cell IL-17 production, unlike IFNγ, is sensitive to GPR43 agonism[301] and thus the low frequencies of GPR43+ LP CD4 cells in our cultures may be reflective of the similarly low- frequency Th17 population.

To determine if the difference in Th subset sensitivity to butyrate was specific to the gut PB Th subsets were similarly tested. In PB Th cells Th17 cells responded to butyrate similarly to Th1 and Th22 cells. Interestingly, the non-IFNγ, IL-17, IL-22 producing PB Th cells proliferated the least in response to TCR stimulation and were the most sensitive to the antiproliferative effects of butyrate. From these data, several possible conclusions can be made. First, the magnitude of proliferation in the Th subsets in the PB and LP differ as does the sensitivity of these cells to butyrate. This could be the result of intrinsic differences between LP and PB Th cells, such as their activation state, or merely differences in the ability of these cells to respond to direct TCR stimulation and subsequently to butyrate. At this time further mechanistic studies are needed to determine the nature of these differential responses to butyrate.

Activated CD4 T cells are readily infected with HIV-1 and depleted in PLWH[286].

Therefore, the observation that butyrate decreased LP CD4 T cell activation and proliferation in a concentration-dependent manner led to the hypothesis that butyrate would similarly decrease HIV-1 infection of LP CD4 T cells due to their decreased activation and proliferation. To test this, I used an in vitro model of acute infection of LPMC with a transmitted founder virus. Butyrate reduced the frequency of productively infected LP CD4 T cells at concentrations ≥ to 0.25mM and increased the frequency of productively infected LP

CD4 T cells at concentrations ≤0.125mM after four days of infection. This was similarly reflected in the frequency of productively infected Th subsets and the production of infectious virions. The observation of a differential impact of butyrate at higher and lower concentrations is interesting since there appears to be an uncoupling of LP CD4 T cell activation and HIV-1 infection in the setting of lower concentrations of butyrate, and an activation-independent regulation of HIV-1 infection at these concentrations. To determine if the impact of butyrate was mediated by direct action on LP CD4 T cells, the effect of butyrate on HIV-1 infection in purified LP CD4 T cells was next assayed. The data from these assays indicated that the impact of butyrate at lower concentrations was indirect, while the effect at higher concentrations was direct.

Previous studies with enteric bacteria have shown that productive HIV-1 infection can be increased significantly in the absence of overt LP CD4 T cell activation through the upregulation of the HIV-1 co-receptor CCR5 by other cells in the LPMC[156]. Due to the established impact of CCR5 expression on HIV-1 infection, the expression of CCR5, as well as CD4, were examined in the setting of TCR stimulation and exogenous butyrate. Similar to the trend seen with productive HIV-1 infection, CCR5 expression increased at lower concentrations of butyrate ≤0.125mM and but was not affected by higher concentrations.

Conversely, low concentrations of butyrate did slightly increase the expression of CD4, but higher concentrations decreased its expression. These assays were next carried out with

97 purified LP CD4 T cells, which demonstrated that the effect of lower concentrations of butyrate on CCR5 expression was dependent on other cell types in the LPMC and the effect on CD4 expression was direct. Collectively, these data indicate that at lower concentrations of butyrate there is an indirect effect of butyrate that increases CCR5 expression and productive HIV-1 infection, while at higher concentrations there is a direct effect of butyrate on LP CD4 T cells which decreased CD4 expression and productive HIV-1 infection.

To determine if the changes in productive infection were due to an increase in viral entry and reverse transcription due to increased co-receptor expression or if more HIV-1 virus was being produced by these cells due to the known effect of HDACi on HIV-1 transcription[302] qPCR based methods examined viral entry, reverse transcription, and viral transcription. The data from these assays indicate there to be a slight increase in HIV-1 viral entry or reverse transcription at 0.125mM and no discernable effects at other concentrations. Conversely, there appeared to be a concentration-dependent increase in viral transcription. These data indicate that there does seem to be an impact on viral entry or reverse transcription at the concentrations of butyrate where there were observed increases in CCR5 expression and productive infection and that this is independent of changes in viral transcription. The viral transcription data is interesting because the increased relative amounts on transcription at 0.5mM do not translate to increased frequencies of productively infected LP CD4 T cells or the production of infectious virions. While the mechanism for the difference is unknown at this time, there are several possibilities for this observation. First, the increase in transcription could be due to the higher relative amounts of HDAC inhibition at 0.5mM. This, however, does not translate to differences in productive infection due to the substantial decreases in LP CD4 T cell activation, proliferation, and CD4 expression.

Secondly, the normalization of the data could be affecting the reported values. There is higher relative proliferation at the lower concentrations of butyrate which could account for a higher amount of GAPDH (housekeeping gene) expressed relative to HIV-1 gag RNA at

98 early timepoint thus resulting in lower reported values. Conversely, at 0.5mM there is a substantial decrease in LP CD4 T cell proliferation and therefore likely less GAPDH present relative to HIV-1 gag RNA. Collectively, these data indicate that butyrate is altering the HIV-

1 life cycle and further in-depth studies are warranted.

Potential Implications

Collectively, the in vitro data outlined in this thesis suggest that in untreated PLWH, low abundances of BPB and therefore likely low concentrations of tissue butyrate may be a contributing factor to high levels of viral replication and infection of LP CD4 T cells. This process is likely mediated indirectly through butyrate’s impact on other cells (e.g., APCs) to drive increased expression of the HIV-1 co-receptor CCR5 and increased viral entry or reverse transcription. This is in contrast to what would occur at higher concentrations of butyrate, where butyrate would directly decrease LP CD4 T cell activation, likely through

HDAC inhibition, thus limiting the infection of LP CD4 T cells. It is possible that this scenario at higher butyrate concentrations would have the most significant effect on T cells that are newly recruited into the gut mucosa and thus limits their subsequent unchecked activation and response to translocating dysbiotic bacteria[79, 103, 231, 285] (Figure 6.1).

In an attempt to link alterations in BPB communities in PLWH and the impact of a loss of tissue concentrations of butyrate, I tested the effect of butyrate on LP CD4 T cell activation and proliferation. While there is a precedence for butyrate’s ability to alter T cell activation and function, these data were the first to demonstrate an immunomodulatory effect of butyrate on human gut LP T cells. Translated to a clinical setting these observations could have an impact on the management of gut centric inflammatory disorders. By increasing butyrate concentrations (within the normal physiologic range) one would anticipate that there would be lower levels of T cell activation both in the gut mucosa and potentially systemically if blood concentrations increase sufficiently. However, due to the immunosuppressive nature of butyrate’s effects on T cells the increase in butyrate in

Figure 6.1 LP CD4 T Cell Activation and HIV-1 Infection are Differentially Impacted by Butyrate in a Concentration-dependent Manner. A model that translates our findings into what may be occurring in the gut mucosa of PLWH. In short, in the setting of the lower abundances of BPB and thus lower tissue butyrate levels, there is a relatively higher amount of LP CD4 T cell activation, CCR5 expression, HIV-1 infection, and inflammation. Conversely, with higher abundances of BPB and tissue butyrate levels, there is a lower relative amount of LP CD4 T cell activation, CD4 expression and subsequent HIV-1 infection likely mediated directly by butyrate through HDACi.

100 both the gut mucosa and systemic circulation would likely be beneficial in environments that have limited exposure to infectious diseases such as Mycobacterium tuberculosis (mTB).

This is highlighted by the observations in Segal et al. [303] who demonstrated that increased serum concentrations of butyrate and propionate in HIV-1 infected individuals were predictive of mTB susceptibility likely due to suppression of Th17 and Th1 responses.

Therefore, there may be a risk in increasing SCFA concentrations outside of normal ranges and caution may be warranted in interventions that increased SCFA concentrations as a target in populations susceptible to infectious disease such as PLWH in areas with endemic infectious disease.

Another important observation made in this thesis is that the antiproliferative effect of butyrate on LP CD4 T cells requires butyrate to either be present before TCR stimulation or very early after TCR stimulation. Once the LP CD4 T cells have been activated for at least

18 hours the antiproliferative capacity of butyrate is diminished. The temporal nature of the butyrate response is likely due to the mechanism by which butyrate alters LP CD4 T cell proliferation. For example, If HDAC inhibition is responsible for the decreased proliferation noted in the presence of butyrate as our model suggests, it is likely that it must be present early in the events of T cell activation to alter the capacity of these cells to undergo proliferation. This would likely occur via the regulation of gene expression and proteins involved in cell cycle (e.g., cyclins) progression. These observations also have clinical implications and may explain the inconsistent results of the modulation of butyrate concentrations either directly or through probiotic and prebiotic therapies in individuals with

IBD [261, 293, 294, 304]. Gut mucosal T cell activation is an important feature of HIV-1 infection and IBD, and it is possible that butyrate interventions would be more efficacious if given to individuals who were currently in milder disease states with limited T cell activation rather than those with more overt T cell activation. Conversely, if butyrate concentrations are increased for an extended period the activation of newly recruited CD4 T cells would likely

101 be dampened and thus act to limit the severity of the disease. Follow up studies on the temporal importance of butyrate addition on HIV-1 infection in our model are also warranted.

The majority of the data presented in this thesis examined the impact of butyrate on

LP CD4 T cell activation and function. However, it is important to note that the other primary

SCFAs, propionate and acetate, also inhibited LP CD4 T cell activation and proliferation in a concentration-dependent manner. These SCFA showed similar activity to butyrate albeit at higher concentrations. These data are not surprising since these SCFA share homology in the surface GPRs that they signal through (GPR41 and GPR43) and their function as an

HDACi[228]. Combined with the lack of significant expression of GPR43 on LP CD4 T cells, these data strengthen the observation that butyrate, as well as propionate and acetate, are likely inhibiting LP CD4 T cell activation and proliferation through HDAC inhibition. These observations also indicate that there are redundancies in the effect of these SCFA on LP

CD4 T cell activation. Therefore, the decrease in the concentration of one SCFA may be compensated for by the presence of the other two, and thus the modulation of total SCFA may be important to target in the regulation of gut mucosal T cell activation.

One of the most salient observations made in this thesis was that the impact of the addition of exogenous butyrate to HIV-1 infected and activated LP CD4 T cells did not result in a concentration-dependent decrease in infection. Indeed, lower concentrations of butyrate increased the productive infection of LP CD4 T cells likely through increased expression of

CCR5 and increased viral entry or reverse transcription. The observations that butyrate increases HIV-1 while simultaneously decreasing LP CD4 T cell activation are somewhat counterintuitive since it is traditionally thought that T cell activation and HIV-1 infection are tightly coupled events. The lack of these observations in cultures with purified LP CD4 T cells indicates a role for another cell type in the LPMC, such as an mDC, in mediating these effects of butyrate. Therefore, it is likely that butyrate is decreasing the activation and proliferation of LP CD4 T cells but at a very narrow concentration range also altering the

102 production of proteins such as cytokines in the LPMC fraction to drive increases in LP CD4

T cell infection. One potential mechanism could be the production of cytokines by bystander cells in LPMC. In an endotoxemia model, it has been demonstrated that rhIL-10 treatment of

PBMCs increased CCR5 expression on human CD4 T cells[289]. Butyrate has also been shown to alter the production of IL-10 from myeloid cells and T cells. Thus, it is possible that butyrate increases LP CD4 T cell infection at lower concentrations via the indirect modulation of cytokines produced in other cell types.

Limitations and Future Directions

There are several limitations to the data presented in this thesis and unanswered questions that remain to be addressed. There are limitations of the in vitro model used to study the effect of butyrate on gut mucosal T cell function and HIV-1 infection. First, the disaggregated nature of the cells used in the model does account for the importance of signals from epithelium and IELs that may alter the impact of butyrate on LP CD4 T cell function. Secondly, the LPMC are primary cells which are not amenable to genetic manipulation in our experience. This makes it challenging to study the importance of specific proteins in our model, such as HDAC proteins, by genetic manipulation. Lastly, the LPMC infection model reflects an acute HIV-1 infection, which is not directly a limitation but makes it more challenging to infer the impact of butyrate in the mucosa of chronically infected individuals.

The observations outlined in this thesis lay the foundation for a multitude of further studies that can be carried out to understand the impact of butyrate on gut mucosa T cell function and HIV-1 infection. It would be informative to look at changes in gene networks and specific genes in LP CD4 T cells treated with higher and lower concentrations of butyrate in the setting or absence of HIV-1 infection and TCR stimulation. This could be done by single-cell RNA sequencing of LP CD4 T cells to understand how butyrate is altering gene expression in specific Th cell subsets by using Th specific transcription factors

103

(e.g., Tbet) for Th subset identification. Furthermore, carrying out these assays on purified

LP CD4 T cells and LP CD4 T cells sorted after culture with total LPMC would generate data on alterations in gene expression due to a direct effect of butyrate relative to those driven by other cell types in the LPMC. If carried out this would be the first assay of its type looking at the impact of butyrate specifically on human gut LP CD4 T cells. From these assays, hypotheses could be generated to further tease out the specific mechanisms by which butyrate alters LP CD4 T cell function.

Furthermore, key follow up experiments to gene expression assays would likely include paired chromatin immunoprecipitation (ChiP) sequencing analysis to evaluate if histone modifications associated with the resultant changes in specific gene expression. As a follow up to these experiments, the inhibition, either chemically or genetically, of specific

HDAC proteins could next be studied to elucidate the mechanism of butyrate’s actions further. Taken together, these assays would inform on the mechanisms by which butyrate alters LP CD4 T cell function and offer insight into potentially druggable targets to limit HIV-1 pathogenesis and inflammation.

In addition, the use of clinical models would help to better understand the effects of butyrate on LP CD4 T cell function and HIV-1 infection and confirm findings from the in vitro studies. With the importance of the microbiome in GI tract health and the observation that

HIV-1 associated dysbiosis may contribute to chronic inflammation and disease progression, several studies have sought to determine if modulation of the microbiome by pro- and prebiotics are beneficial in PLWH. The results of these studies have been mixed but have shown promise[146-151]. One encouraging study in PLWH carried out by Serrano-Villar et al. [152] examined the impact of pre-biotics and glutamine (scGOS/lcFOS/glutamine) supplementation on the composition of the microbiota and immunological markers in PLWH.

The results of that study demonstrated that prebiotic supplementation reduced HIV-1 dysbiosis of the stool microbiota. Also, they noted a decline in peripheral T cell activation,

104 microbial translocation and an increase in butyrate production in individuals in the prebiotic treatment arm. Lastly, they noted that increased relative abundance of stool

Faecalibacterium and Lachnospira associated with increased stool butyrate concentrations and lower microbial translocation and systemic inflammation[152]. While these studies are promising, they use probiotics and prebiotic supplementation, which are both nonspecific interventions and limit the interrogation of the specific pathways involved in the positive clinical outcomes. To overcome this, a similar model using a specific butyrate therapy, such as orally available butyrate in the form of tributyrin[305], could be employed in treated and untreated PLWH. After exposure to butyrate, frequencies of and activation states of Th subsets in the gut mucosa of these individuals could be investigated in mucosal biopsies.

Furthermore, LP CD4 T cells could be isolated from these individuals and similarly tested on a limited scale in the LPMC model outlined in this thesis. Additionally, the investigation of specific genes and gene networks identified in the proposed genomics study could be investigated in vivo to further confirm butyrate’s mechanism of action on gut mucosal CD4 T cells.

In summary, the data presented in this thesis offer a foundation to further explore the mechanism by which altered concentrations of butyrate in the gut mucosal tissue impact LP

CD4 T cell activation and HIV-1 disease progression. Furthermore, alterations in gut butyrate concentrations offer an attractive target for therapeutic interventions for PLWH and others with inflammatory disorders of the GI tract. While not likely curative, the increase in butyrate concentrations in the gut mucosal tissue of PLWH may dampen the negative impact of HIV-1 infection in the gut mucosa and limit the chronic immune activation that is characteristic of HIV-1 infection.

105

REFERENCES

1. UNAIDS, Global AIDS update 2016. UNAIDS Report on Global AIDS Epidemic: Joint United Nations Progamme on HIV-1/AIDS, 2016.

2. Cohen, M.S., et al., Antiretroviral Therapy for the Prevention of HIV-1 Transmission. N Engl J Med, 2016. 375(9): p. 830-9.

3. Palella, F.J., Jr., et al., Declining morbidity and mortality among patients with advanced human immunodeficiency virus infection. HIV Outpatient Study Investigators. N Engl J Med, 1998. 338(13): p. 853-60.

4. Autran, B., et al., Positive effects of combined antiretroviral therapy on CD4+ T cell homeostasis and function in advanced HIV disease. Science, 1997. 277(5322): p. 112- 6.

5. Donnell, D., et al., Heterosexual HIV-1 transmission after initiation of antiretroviral therapy: a prospective cohort analysis. Lancet, 2010. 375(9731): p. 2092-8.

6. Cohen, M.S., et al., Prevention of HIV-1 infection with early antiretroviral therapy. N Engl J Med, 2011. 365(6): p. 493-505.

7. Richman, D.D., et al., The challenge of finding a cure for HIV infection. Science, 2009. 323(5919): p. 1304-7.

8. German Advisory Committee Blood, S.A.o.P.T.b.B., Human Immunodeficiency Virus (HIV). Transfus Med Hemother, 2016. 43(3): p. 203-22.

9. Foley B, L.T., Apetrei C, Hahn B, Mizrachi I, Mullins J, Rambaut A, Wolinsky S, and Korber B, HIV Sequence Compendium 2018. Theoretical Biology and Biophysics Group, 2018.

10. Goto, T., M. Nakai, and K. Ikuta, The life-cycle of human immunodeficiency virus type 1. Micron, 1998. 29(2-3): p. 123-38.

11. Silberman, S.L., et al., The interaction of CD4 with HIV-1 gp120. Semin Immunol, 1991. 3(3): p. 187-92.

12. Arthos, J., et al., Identification of the residues in human CD4 critical for the binding of HIV. Cell, 1989. 57(3): p. 469-81.

13. Wu, L., et al., CD4-induced interaction of primary HIV-1 gp120 glycoproteins with the chemokine receptor CCR-5. Nature, 1996. 384(6605): p. 179-83.

14. Dragic, T., et al., HIV-1 entry into CD4+ cells is mediated by the chemokine receptor CC-CKR-5. Nature, 1996. 381(6584): p. 667-73.

15. Deng, H., et al., Identification of a major co-receptor for primary isolates of HIV-1. Nature, 1996. 381(6584): p. 661-6.

16. Berger, E.A., et al., A new classification for HIV-1. Nature, 1998. 391(6664): p. 240.

106

17. Alkhatib, G. and E.A. Berger, HIV coreceptors: from discovery and designation to new paradigms and promise. Eur J Med Res, 2007. 12(9): p. 375-84.

18. Parrish, N.F., et al., Transmitted/founder and chronic subtype C HIV-1 use CD4 and CCR5 receptors with equal efficiency and are not inhibited by blocking the integrin alpha4beta7. PLoS Pathog, 2012. 8(5): p. e1002686.

19. Parrish, N.F., et al., Phenotypic properties of transmitted founder HIV-1. Proc Natl Acad Sci U S A, 2013. 110(17): p. 6626-33.

20. Pancera, M., et al., Structure of HIV-1 gp120 with gp41-interactive region reveals layered envelope architecture and basis of conformational mobility. Proc Natl Acad Sci U S A, 2010. 107(3): p. 1166-71.

21. Pancera, M., et al., Structure and immune recognition of trimeric pre-fusion HIV-1 Env. Nature, 2014. 514(7523): p. 455-61.

22. Deeks, S.G., et al., HIV infection. Nat Rev Dis Primers, 2015. 1: p. 15035.

23. Peng, H., et al., Enhancement or inhibition of HIV-1 replication by intracellular expression of sense or antisense RNA targeted at different intermediates of reverse transcription. AIDS, 1997. 11(5): p. 587-95.

24. Bauer, M., et al., Structural constraints of HIV-1 Nef may curtail escape from HLA-B7- restricted CTL recognition. Immunol Lett, 1997. 55(2): p. 119-22.

25. Fischl, M.A., et al., The efficacy of azidothymidine (AZT) in the treatment of patients with AIDS and AIDS-related complex. A double-blind, placebo-controlled trial. N Engl J Med, 1987. 317(4): p. 185-91.

26. Sluis-Cremer, N., D. Arion, and M.A. Parniak, Molecular mechanisms of HIV-1 resistance to nucleoside reverse transcriptase inhibitors (NRTIs). Cell Mol Life Sci, 2000. 57(10): p. 1408-22.

27. Sluis-Cremer, N. and G. Tachedjian, Mechanisms of inhibition of HIV replication by non-nucleoside reverse transcriptase inhibitors. Virus Res, 2008. 134(1-2): p. 147-56.

28. Lv, Z., Y. Chu, and Y. Wang, HIV protease inhibitors: a review of molecular selectivity and toxicity. HIV AIDS (Auckl), 2015. 7: p. 95-104.

29. Hicks, C. and R.M. Gulick, Raltegravir: the first HIV type 1 integrase inhibitor. Clin Infect Dis, 2009. 48(7): p. 931-9.

30. Ray, N., Maraviroc in the treatment of HIV infection. Drug Des Devel Ther, 2009. 2: p. 151-61.

31. He, Y., et al., Design and evaluation of sifuvirtide, a novel HIV-1 fusion inhibitor. J Biol Chem, 2008. 283(17): p. 11126-34.

32. Fenton-May, A.E., et al., Relative resistance of HIV-1 founder viruses to control by interferon-alpha. Retrovirology, 2013. 10: p. 146.

107

33. Puryear, W.B., et al., Interferon-inducible mechanism of dendritic cell-mediated HIV-1 dissemination is dependent on Siglec-1/CD169. PLoS Pathog, 2013. 9(4): p. e1003291.

34. Izquierdo-Useros, N., et al., HIV-1 capture and transmission by dendritic cells: the role of viral glycolipids and the cellular receptor Siglec-1. PLoS Pathog, 2014. 10(7): p. e1004146.

35. Akiyama, H., et al., Virus particle release from glycosphingolipid-enriched microdomains is essential for dendritic cell-mediated capture and transfer of HIV-1 and henipavirus. J Virol, 2014. 88(16): p. 8813-25.

36. Brenchley, J.M., et al., CD4+ T cell depletion during all stages of HIV disease occurs predominantly in the gastrointestinal tract. J Exp Med, 2004. 200(6): p. 749-59.

37. Chege, D., et al., Sigmoid Th17 populations, the HIV latent reservoir, and microbial translocation in men on long-term antiretroviral therapy. AIDS, 2011. 25(6): p. 741-9.

38. Kiertiburanakul, S., et al., Epidemiology of bloodstream infections and predictive factors of mortality among HIV-infected adult patients in Thailand in the era of highly active antiretroviral therapy. Jpn J Infect Dis, 2012. 65(1): p. 28-32.

39. Kumarasamy, N., et al., Factors associated with mortality among HIV-infected patients in the era of highly active antiretroviral therapy in southern India. Int J Infect Dis, 2010. 14(2): p. e127-31.

40. Mehandru, S., et al., Primary HIV-1 infection is associated with preferential depletion of CD4+ T lymphocytes from effector sites in the gastrointestinal tract. J Exp Med, 2004. 200(6): p. 761-70.

41. Schneider, T., et al., Loss of CD4 T lymphocytes in patients infected with human immunodeficiency virus type 1 is more pronounced in the duodenal mucosa than in the peripheral blood. Berlin Diarrhea/Wasting Syndrome Study Group. Gut, 1995. 37(4): p. 524-9.

42. Brenchley, J.M. and D.C. Douek, HIV infection and the gastrointestinal immune system. Mucosal Immunol, 2008. 1(1): p. 23-30.

43. Brenchley, J.M., et al., Differential Th17 CD4 T-cell depletion in pathogenic and nonpathogenic lentiviral infections. Blood, 2008. 112(7): p. 2826-35.

44. Mehandru, S., et al., Mechanisms of gastrointestinal CD4+ T-cell depletion during acute and early human immunodeficiency virus type 1 infection. J Virol, 2007. 81(2): p. 599-612.

45. Guadalupe, M., et al., Severe CD4+ T-cell depletion in gut lymphoid tissue during primary human immunodeficiency virus type 1 infection and substantial delay in restoration following highly active antiretroviral therapy. J Virol, 2003. 77(21): p. 11708-17.

108

46. Anton, P.A., et al., Enhanced levels of functional HIV-1 co-receptors on human mucosal T cells demonstrated using intestinal biopsy tissue. AIDS, 2000. 14(12): p. 1761-5.

47. Paiardini, M., et al., Lessons learned from the natural hosts of HIV-related viruses. Annu Rev Med, 2009. 60: p. 485-95.

48. Kestens, L., et al., Expression of activation antigens, HLA-DR and CD38, on CD8 lymphocytes during HIV-1 infection. AIDS, 1992. 6(8): p. 793-7.

49. Liu, Z., et al., Elevated CD38 antigen expression on CD8+ T cells is a stronger marker for the risk of chronic HIV disease progression to AIDS and death in the Multicenter AIDS Cohort Study than CD4+ cell count, soluble immune activation markers, or combinations of HLA-DR and CD38 expression. J Acquir Immune Defic Syndr Hum Retrovirol, 1997. 16(2): p. 83-92.

50. Giorgi, J.V., et al., Shorter survival in advanced human immunodeficiency virus type 1 infection is more closely associated with T lymphocyte activation than with plasma virus burden or virus chemokine coreceptor usage. J Infect Dis, 1999. 179(4): p. 859- 70.

51. Kestens, L., et al., Selective increase of activation antigens HLA-DR and CD38 on CD4+ CD45RO+ T lymphocytes during HIV-1 infection. Clin Exp Immunol, 1994. 95(3): p. 436-41.

52. Ramzaoui, S., et al., During HIV infection, CD4+ CD38+ T-cells are the predominant circulating CD4+ subset whose HLA-DR positivity increases with disease progression and whose V beta repertoire is similar to that of CD4+ CD38- T-cells. Clin Immunol Immunopathol, 1995. 77(1): p. 33-41.

53. Shebl, F.M., et al., Increased levels of circulating cytokines with HIV-related immunosuppression. AIDS Res Hum Retroviruses, 2012. 28(8): p. 809-15.

54. Thea, D.M., et al., Plasma cytokines, cytokine antagonists, and disease progression in African women infected with HIV-1. Ann Intern Med, 1996. 124(8): p. 757-62.

55. Haissman, J.M., et al., Plasma cytokine levels in Tanzanian HIV-1-infected adults and the effect of antiretroviral treatment. J Acquir Immune Defic Syndr, 2009. 52(4): p. 493- 7.

56. Breen, E.C., et al., Infection with HIV is associated with elevated IL-6 levels and production. J Immunol, 1990. 144(2): p. 480-4.

57. Dillon, S.M., et al., Gut dendritic cell activation links an altered colonic microbiome to mucosal and systemic T-cell activation in untreated HIV-1 infection. Mucosal Immunol, 2016. 9(1): p. 24-37.

58. Del Corno, M., et al., HIV-1 gp120 signaling through TLR4 modulates innate immune activation in human macrophages and the biology of hepatic stellate cells. J Leukoc Biol, 2016. 100(3): p. 599-606.

109

59. Hunt, P.W., et al., Impact of CD8+ T-cell activation on CD4+ T-cell recovery and mortality in HIV-infected Ugandans initiating antiretroviral therapy. AIDS, 2011. 25(17): p. 2123-31.

60. Bartha, I., et al., Estimating the Respective Contributions of Human and Viral Genetic Variation to HIV Control. PLoS Comput Biol, 2017. 13(2): p. e1005339.

61. 1993 revised classification system for HIV infection and expanded surveillance case definition for AIDS among adolescents and adults. MMWR Recomm Rep, 1992. 41(RR-17): p. 1-19.

62. van Wijk, F. and H. Cheroutre, Mucosal T cells in gut homeostasis and inflammation. Expert Rev Clin Immunol, 2010. 6(4): p. 559-66.

63. Tokuhara, D., et al., A comprehensive understanding of the gut mucosal immune system in allergic inflammation. Allergol Int, 2019. 68(1): p. 17-25.

64. Peterson, L.W. and D. Artis, Intestinal epithelial cells: regulators of barrier function and immune homeostasis. Nat Rev Immunol, 2014. 14(3): p. 141-53.

65. Pott, J. and M. Hornef, Innate immune signalling at the intestinal epithelium in homeostasis and disease. EMBO Rep, 2012. 13(8): p. 684-98.

66. Sheridan, B.S. and L. Lefrancois, Intraepithelial lymphocytes: to serve and protect. Curr Gastroenterol Rep, 2010. 12(6): p. 513-21.

67. Napier, R.J., et al., The Role of Mucosal Associated Invariant T Cells in Antimicrobial Immunity. Front Immunol, 2015. 6: p. 344.

68. Ahluwalia, B., M.K. Magnusson, and L. Ohman, Mucosal immune system of the gastrointestinal tract: maintaining balance between the good and the bad. Scand J Gastroenterol, 2017. 52(11): p. 1185-1193.

69. Brenchley, J.M. and D.C. Douek, The mucosal barrier and immune activation in HIV pathogenesis. Curr Opin HIV AIDS, 2008. 3(3): p. 356-61.

70. Liu, J., et al., Inside Out: HIV, the Gut Microbiome, and the Mucosal Immune System. J Immunol, 2017. 198(2): p. 605-614.

71. Lehmann, C., et al., Longitudinal analysis of distribution and function of plasmacytoid dendritic cells in peripheral blood and gut mucosa of HIV infected patients. J Infect Dis, 2014. 209(6): p. 940-9.

72. Shale, M., C. Schiering, and F. Powrie, CD4(+) T-cell subsets in intestinal inflammation. Immunol Rev, 2013. 252(1): p. 164-82.

73. Alabbas, S.Y., et al., The role of IL-22 in the resolution of sterile and nonsterile inflammation. Clin Transl Immunology, 2018. 7(4): p. e1017.

74. Eyerich, K., V. Dimartino, and A. Cavani, IL-17 and IL-22 in immunity: Driving protection and pathology. Eur J Immunol, 2017. 47(4): p. 607-614.

110

75. Bowcutt, R., et al., Isolation and cytokine analysis of lamina propria lymphocytes from mucosal biopsies of the human colon. J Immunol Methods, 2015. 421: p. 27-35.

76. Luckheeram, R.V., et al., CD4(+)T cells: differentiation and functions. Clin Dev Immunol, 2012. 2012: p. 925135.

77. MacLeod, M.K., J.W. Kappler, and P. Marrack, Memory CD4 T cells: generation, reactivation and re-assignment. Immunology, 2010. 130(1): p. 10-5.

78. Smith-Garvin, J.E., G.A. Koretzky, and M.S. Jordan, T cell activation. Annu Rev Immunol, 2009. 27: p. 591-619.

79. Curtsinger, J.M. and M.F. Mescher, Inflammatory cytokines as a third signal for T cell activation. Curr Opin Immunol, 2010. 22(3): p. 333-40.

80. Kumar, B.V., et al., Human Tissue-Resident Memory T Cells Are Defined by Core Transcriptional and Functional Signatures in Lymphoid and Mucosal Sites. Cell Rep, 2017. 20(12): p. 2921-2934.

81. Booth, J.S., et al., Characterization and functional properties of gastric tissue-resident memory T cells from children, adults, and the elderly. Front Immunol, 2014. 5: p. 294.

82. Schluns, K.S. and K.D. Klonowski, Protecting the borders: tissue-resident memory T cells on the front line. Front Immunol, 2015. 6: p. 90.

83. Schenkel, J.M. and D. Masopust, Tissue-resident memory T cells. Immunity, 2014. 41(6): p. 886-97.

84. Mackay, L.K., et al., Cutting edge: CD69 interference with sphingosine-1-phosphate receptor function regulates peripheral T cell retention. J Immunol, 2015. 194(5): p. 2059-63.

85. Bolduc, J.F., et al., Epigenetic Metabolite Acetate Inhibits Class I/II Histone Deacetylases, Promotes Histone Acetylation, and Increases HIV-1 Integration in CD4(+) T Cells. J Virol, 2017. 91(16).

86. Panea, C., et al., Intestinal Monocyte-Derived Macrophages Control Commensal- Specific Th17 Responses. Cell Rep, 2015. 12(8): p. 1314-24.

87. Goto, Y., et al., Segmented filamentous bacteria antigens presented by intestinal dendritic cells drive mucosal Th17 cell differentiation. Immunity, 2014. 40(4): p. 594- 607.

88. Atarashi, K., et al., Induction of colonic regulatory T cells by indigenous Clostridium species. Science, 2011. 331(6015): p. 337-41.

89. Atarashi, K., et al., Treg induction by a rationally selected mixture of Clostridia strains from the human microbiota. Nature, 2013. 500(7461): p. 232-6.

111

90. Round, J.L. and S.K. Mazmanian, Inducible Foxp3+ regulatory T-cell development by a commensal bacterium of the intestinal microbiota. Proc Natl Acad Sci U S A, 2010. 107(27): p. 12204-9.

91. Ivanov, II, et al., Specific microbiota direct the differentiation of IL-17-producing T- helper cells in the mucosa of the small intestine. Cell Host Microbe, 2008. 4(4): p. 337- 49.

92. Kewenig, S., et al., Rapid mucosal CD4(+) T-cell depletion and enteropathy in simian immunodeficiency virus-infected rhesus macaques. Gastroenterology, 1999. 116(5): p. 1115-23.

93. Hensley-McBain, T., et al., Intestinal damage precedes mucosal immune dysfunction in SIV infection. Mucosal Immunol, 2018. 11(5): p. 1429-1440.

94. Hirao, L.A., et al., Early mucosal sensing of SIV infection by paneth cells induces IL- 1beta production and initiates gut epithelial disruption. PLoS Pathog, 2014. 10(8): p. e1004311.

95. Cecchinato, V., et al., Altered balance between Th17 and Th1 cells at mucosal sites predicts AIDS progression in simian immunodeficiency virus-infected macaques. Mucosal Immunol, 2008. 1(4): p. 279-88.

96. Nazli, A., et al., Exposure to HIV-1 directly impairs mucosal epithelial barrier integrity allowing microbial translocation. PLoS Pathog, 2010. 6(4): p. e1000852.

97. Smith, A.J., et al., A role for syndecan-1 and claudin-2 in microbial translocation during HIV-1 infection. J Acquir Immune Defic Syndr, 2010. 55(3): p. 306-15.

98. Prendergast, A.J., et al., Intestinal Damage and Inflammatory Biomarkers in Human Immunodeficiency Virus (HIV)-Exposed and HIV-Infected Zimbabwean Infants. J Infect Dis, 2017. 216(6): p. 651-661.

99. Cheru, L.T., et al., I-FABP Is Higher in People With Chronic HIV Than Elite Controllers, Related to Sugar and Fatty Acid Intake and Inversely Related to Body Fat in People With HIV. Open Forum Infect Dis, 2018. 5(11): p. ofy288.

100. Brenchley, J.M., et al., Microbial translocation is a cause of systemic immune activation in chronic HIV infection. Nat Med, 2006. 12(12): p. 1365-71.

101. Dinh, D.M., et al., Intestinal microbiota, microbial translocation, and systemic inflammation in chronic HIV infection. J Infect Dis, 2015. 211(1): p. 19-27.

102. Klatt, N.R. and J.M. Brenchley, Th17 cell dynamics in HIV infection. Curr Opin HIV AIDS, 2010. 5(2): p. 135-40.

103. Dillon, S.M., et al., An altered intestinal mucosal microbiome in HIV-1 infection is associated with mucosal and systemic immune activation and endotoxemia. Mucosal Immunol, 2014. 7(4): p. 983-94.

112

104. Leon, A., et al., Association of microbial translocation biomarkers with clinical outcome in controllers HIV-infected patients. AIDS, 2015. 29(6): p. 675-81.

105. Timmons, T., et al., Microbial translocation and metabolic and body composition measures in treated and untreated HIV infection. AIDS Res Hum Retroviruses, 2014. 30(3): p. 272-7.

106. Tincati, C., D.C. Douek, and G. Marchetti, Gut barrier structure, mucosal immunity and intestinal microbiota in the pathogenesis and treatment of HIV infection. AIDS Res Ther, 2016. 13: p. 19.

107. Marchetti, G., et al., Microbial translocation predicts disease progression of HIV- infected antiretroviral-naive patients with high CD4+ cell count. AIDS, 2011. 25(11): p. 1385-94.

108. Cassol, E., et al., Persistent microbial translocation and immune activation in HIV-1- infected South Africans receiving combination antiretroviral therapy. J Infect Dis, 2010. 202(5): p. 723-33.

109. Marchetti, G., et al., Microbial translocation is associated with sustained failure in CD4+ T-cell reconstitution in HIV-infected patients on long-term highly active antiretroviral therapy. AIDS, 2008. 22(15): p. 2035-8.

110. Dillon, S.M., et al., Brief Report: Inflammatory Colonic Innate Lymphoid Cells Are Increased During Untreated HIV-1 Infection and Associated With Markers of Gut Dysbiosis and Mucosal Immune Activation. J Acquir Immune Defic Syndr, 2017. 76(4): p. 431-437.

111. Vujkovic-Cvijin, I., et al., Dysbiosis of the gut microbiota is associated with HIV disease progression and tryptophan catabolism. Sci Transl Med, 2013. 5(193): p. 193ra91.

112. Hunt, P.W., et al., Gut epithelial barrier dysfunction and innate immune activation predict mortality in treated HIV infection. J Infect Dis, 2014. 210(8): p. 1228-38.

113. Lapenta, C., et al., Human intestinal lamina propria lymphocytes are naturally permissive to HIV-1 infection. Eur J Immunol, 1999. 29(4): p. 1202-8.

114. Kim, C.J., et al., A role for mucosal IL-22 production and Th22 cells in HIV-associated mucosal immunopathogenesis. Mucosal Immunol, 2012. 5(6): p. 670-80.

115. Salgado, M., et al., Long-term non-progressors display a greater number of Th17 cells than HIV-infected typical progressors. Clin Immunol, 2011. 139(2): p. 110-4.

116. Ciccone, E.J., et al., CD4+ T cells, including Th17 and cycling subsets, are intact in the gut mucosa of HIV-1-infected long-term nonprogressors. J Virol, 2011. 85(12): p. 5880-8.

117. Gordon, S.N., et al., Severe depletion of mucosal CD4+ T cells in AIDS-free simian immunodeficiency virus-infected sooty mangabeys. J Immunol, 2007. 179(5): p. 3026- 34.

113

118. Favre, D., et al., Critical loss of the balance between Th17 and T regulatory cell populations in pathogenic SIV infection. PLoS Pathog, 2009. 5(2): p. e1000295.

119. McGary, C.S., et al., The loss of CCR6(+) and CD161(+) CD4(+) T-cell homeostasis contributes to disease progression in SIV-infected rhesus macaques. Mucosal Immunol, 2017. 10(4): p. 1082-1096.

120. Ryan, E.S., et al., Loss of Function of Intestinal IL-17 and IL-22 Producing Cells Contributes to Inflammation and Viral Persistence in SIV-Infected Rhesus Macaques. PLoS Pathog, 2016. 12(2): p. e1005412.

121. Christensen-Quick, A., et al., Human Th17 Cells Lack HIV-Inhibitory RNases and Are Highly Permissive to Productive HIV Infection. J Virol, 2016. 90(17): p. 7833-47.

122. Alvarez, Y., et al., Preferential HIV infection of CCR6+ Th17 cells is associated with higher levels of virus receptor expression and lack of CCR5 ligands. J Virol, 2013. 87(19): p. 10843-54.

123. El Hed, A., et al., Susceptibility of human Th17 cells to human immunodeficiency virus and their perturbation during infection. J Infect Dis, 2010. 201(6): p. 843-54.

124. Planas, D., et al., HIV-1 selectively targets gut-homing CCR6+CD4+ T cells via mTOR- dependent mechanisms. JCI Insight, 2017. 2(15).

125. Cleret-Buhot, A., et al., Identification of novel HIV-1 dependency factors in primary CCR4(+)CCR6(+)Th17 cells via a genome-wide transcriptional approach. Retrovirology, 2015. 12: p. 102.

126. Somsouk, M., et al., Gut epithelial barrier and systemic inflammation during chronic HIV infection. AIDS, 2015. 29(1): p. 43-51.

127. Sereti, I., et al., Decreases in colonic and systemic inflammation in chronic HIV infection after IL-7 administration. PLoS Pathog, 2014. 10(1): p. e1003890.

128. Dillon, S.M., et al., A compartmentalized type I interferon response in the gut during chronic HIV-1 infection is associated with immunopathogenesis. AIDS, 2018. 32(12): p. 1599-1611.

129. Mestecky, J., et al., Humoral immune responses to HIV in the mucosal secretions and sera of HIV-infected women. Am J Reprod Immunol, 2014. 71(6): p. 600-7.

130. Gilbert, J.A., et al., Current understanding of the human microbiome. Nat Med, 2018. 24(4): p. 392-400.

131. Jie, Z., et al., The gut microbiome in atherosclerotic cardiovascular disease. Nat Commun, 2017. 8(1): p. 845.

132. Wan, Y.D., et al., Gut Microbiota Disruption in Septic Shock Patients: A Pilot Study. Med Sci Monit, 2018. 24: p. 8639-8646.

114

133. Castaner, O., et al., The Gut Microbiome Profile in Obesity: A Systematic Review. Int J Endocrinol, 2018. 2018: p. 4095789.

134. Frank, D.N., et al., Molecular-phylogenetic characterization of microbial community imbalances in human inflammatory bowel diseases. Proc Natl Acad Sci U S A, 2007. 104(34): p. 13780-5.

135. Machiels, K., et al., A decrease of the butyrate-producing species Roseburia hominis and Faecalibacterium prausnitzii defines dysbiosis in patients with ulcerative colitis. Gut, 2014. 63(8): p. 1275-83.

136. Takahashi, K., et al., Reduced Abundance of Butyrate-Producing Bacteria Species in the Fecal Microbial Community in Crohn's Disease. Digestion, 2016. 93(1): p. 59-65.

137. Wang, W., et al., Increased proportions of Bifidobacterium and the Lactobacillus group and loss of butyrate-producing bacteria in inflammatory bowel disease. J Clin Microbiol, 2014. 52(2): p. 398-406.

138. Kleiman, S.C., et al., The Intestinal Microbiota in Acute Anorexia Nervosa and During Renourishment: Relationship to Depression, Anxiety, and Eating Disorder Psychopathology. Psychosom Med, 2015. 77(9): p. 969-81.

139. Ding, H.T., Y. Taur, and J.T. Walkup, Gut Microbiota and Autism: Key Concepts and Findings. J Autism Dev Disord, 2017. 47(2): p. 480-489.

140. Gensollen, T., et al., How colonization by microbiota in early life shapes the immune system. Science, 2016. 352(6285): p. 539-44.

141. Donnet-Hughes, A., et al., Potential role of the intestinal microbiota of the mother in neonatal immune education. Proc Nutr Soc, 2010. 69(3): p. 407-15.

142. Strzepa, A., et al., Broad spectrum antibiotic enrofloxacin modulates contact sensitivity through gut microbiota in a murine model. J Allergy Clin Immunol, 2017. 140(1): p. 121-133 e3.

143. Francino, M.P., Antibiotics and the Human Gut Microbiome: Dysbioses and Accumulation of Resistances. Front Microbiol, 2015. 6: p. 1543.

144. Dillon, S.M., D.N. Frank, and C.C. Wilson, The gut microbiome and HIV-1 pathogenesis: a two-way street. AIDS, 2016. 30(18): p. 2737-2751.

145. McHardy, I.H., et al., HIV Infection is associated with compositional and functional shifts in the rectal mucosal microbiota. Microbiome, 2013. 1(1): p. 26.

146. Mutlu, E.A., et al., A compositional look at the human gastrointestinal microbiome and immune activation parameters in HIV infected subjects. PLoS Pathog, 2014. 10(2): p. e1003829.

147. Nowak, P., et al., Gut microbiota diversity predicts immune status in HIV-1 infection. AIDS, 2015. 29(18): p. 2409-18.

115

148. Sun, Y., et al., Fecal bacterial microbiome diversity in chronic HIV-infected patients in China. Emerg Microbes Infect, 2016. 5: p. e31.

149. Lozupone, C.A., et al., Alterations in the gut microbiota associated with HIV-1 infection. Cell Host Microbe, 2013. 14(3): p. 329-39.

150. Monaco, C.L., et al., Altered Virome and Bacterial Microbiome in Human Immunodeficiency Virus-Associated Acquired Immunodeficiency Syndrome. Cell Host Microbe, 2016. 19(3): p. 311-22.

151. Yang, L., et al., HIV-induced immunosuppression is associated with colonization of the proximal gut by environmental bacteria. AIDS, 2016. 30(1): p. 19-29.

152. Volpe, G.E., et al., Associations of cocaine use and HIV infection with the intestinal microbiota, microbial translocation, and inflammation. J Stud Alcohol Drugs, 2014. 75(2): p. 347-57.

153. Perez-Santiago, J., et al., Gut Lactobacillales are associated with higher CD4 and less microbial translocation during HIV infection. AIDS, 2013. 27(12): p. 1921-31.

154. Vazquez-Castellanos, J.F., et al., Altered metabolism of gut microbiota contributes to chronic immune activation in HIV-infected individuals. Mucosal Immunol, 2015. 8(4): p. 760-72.

155. Armstrong, A.J.S., et al., An exploration of Prevotella-rich microbiomes in HIV and men who have sex with men. Microbiome, 2018. 6(1): p. 198.

156. Dillon, S.M., et al., Enhancement of HIV-1 infection and intestinal CD4+ T cell depletion ex vivo by gut microbes altered during chronic HIV-1 infection. Retrovirology, 2016. 13: p. 5.

157. Pescatore, N.A., et al., Short Communication: Anatomic Site of Sampling and the Rectal Mucosal Microbiota in HIV Negative Men Who Have Sex with Men Engaging in Condomless Receptive Anal Intercourse. AIDS Res Hum Retroviruses, 2018. 34(3): p. 277-281.

158. Kelley, C.F., et al., The rectal mucosa and condomless receptive anal intercourse in HIV-negative MSM: implications for HIV transmission and prevention. Mucosal Immunol, 2017. 10(4): p. 996-1007.

159. Noguera-Julian, M., et al., Gut Microbiota Linked to Sexual Preference and HIV Infection. EBioMedicine, 2016. 5: p. 135-46.

160. Noguera-Julian, M., et al., Gut Microbiota Linked to Sexual Preference and HIV Infection. EBioMedicine, 2016.

161. Macfarlane, G.T. and S. Macfarlane, Fermentation in the human large intestine: its physiologic consequences and the potential contribution of prebiotics. J Clin Gastroenterol, 2011. 45 Suppl: p. S120-7.

116

162. Louis, P. and H.J. Flint, Formation of propionate and butyrate by the human colonic microbiota. Environ Microbiol, 2017. 19(1): p. 29-41.

163. den Besten, G., et al., The role of short-chain fatty acids in the interplay between diet, gut microbiota, and host energy metabolism. J Lipid Res, 2013. 54(9): p. 2325-40.

164. Topping, D.L. and P.M. Clifton, Short-chain fatty acids and human colonic function: roles of resistant starch and nonstarch polysaccharides. Physiol Rev, 2001. 81(3): p. 1031-64.

165. Lee, W.J. and K. Hase, Gut microbiota-generated metabolites in animal health and disease. Nat Chem Biol, 2014. 10(6): p. 416-24.

166. Cummings, J.H., et al., Short chain fatty acids in human large intestine, portal, hepatic and venous blood. Gut, 1987. 28(10): p. 1221-7.

167. Bloemen, J.G., et al., Short chain fatty acids exchange across the gut and liver in humans measured at surgery. Clin Nutr, 2009. 28(6): p. 657-61.

168. Pryde, S.E., et al., The microbiology of butyrate formation in the human colon. FEMS Microbiol Lett, 2002. 217(2): p. 133-9.

169. Vital, M., et al., A gene-targeted approach to investigate the intestinal butyrate- producing bacterial community. Microbiome, 2013. 1(1): p. 8.

170. Vital, M., A.C. Howe, and J.M. Tiedje, Revealing the bacterial butyrate synthesis pathways by analyzing (meta)genomic data. MBio, 2014. 5(2): p. e00889.

171. Donohoe, D.R., et al., The microbiome and butyrate regulate energy metabolism and autophagy in the mammalian colon. Cell Metab, 2011. 13(5): p. 517-26.

172. Kelly, C.J., et al., Crosstalk between Microbiota-Derived Short-Chain Fatty Acids and Intestinal Epithelial HIF Augments Tissue Barrier Function. Cell Host Microbe, 2015. 17(5): p. 662-71.

173. Zheng, L., et al., Microbial-Derived Butyrate Promotes Epithelial Barrier Function through IL-10 Receptor-Dependent Repression of Claudin-2. J Immunol, 2017. 199(8): p. 2976-2984.

174. Donohoe, D.R., et al., Microbial regulation of glucose metabolism and cell-cycle progression in mammalian colonocytes. PLoS One, 2012. 7(9): p. e46589.

175. Delcuve, G.P., D.H. Khan, and J.R. Davie, Roles of histone deacetylases in epigenetic regulation: emerging paradigms from studies with inhibitors. Clin Epigenetics, 2012. 4(1): p. 5.

176. Bannister, A.J. and T. Kouzarides, Regulation of chromatin by histone modifications. Cell Res, 2011. 21(3): p. 381-95.

177. Seto, E. and M. Yoshida, Erasers of histone acetylation: the histone deacetylase enzymes. Cold Spring Harb Perspect Biol, 2014. 6(4): p. a018713.

117

178. Dokmanovic, M., C. Clarke, and P.A. Marks, Histone deacetylase inhibitors: overview and perspectives. Mol Cancer Res, 2007. 5(10): p. 981-9.

179. Thiagalingam, S., et al., Histone deacetylases: unique players in shaping the epigenetic histone code. Ann N Y Acad Sci, 2003. 983: p. 84-100.

180. Davie, J.R., Inhibition of histone deacetylase activity by butyrate. J Nutr, 2003. 133(7 Suppl): p. 2485S-2493S.

181. Aoyama, M., J. Kotani, and M. Usami, Butyrate and propionate induced activated or non-activated neutrophil apoptosis via HDAC inhibitor activity but without activating GPR-41/GPR-43 pathways. Nutrition, 2010. 26(6): p. 653-61.

182. Sekhavat, A., J.M. Sun, and J.R. Davie, Competitive inhibition of histone deacetylase activity by trichostatin A and butyrate. Biochem Cell Biol, 2007. 85(6): p. 751-8.

183. Waldecker, M., et al., Inhibition of histone-deacetylase activity by short-chain fatty acids and some polyphenol metabolites formed in the colon. J Nutr Biochem, 2008. 19(9): p. 587-93.

184. Lecona, E., et al., Kinetic analysis of butyrate transport in human colon adenocarcinoma cells reveals two different carrier-mediated mechanisms. Biochem J, 2008. 409(1): p. 311-20.

185. Ganapathy, V., et al., Transporters and receptors for short-chain fatty acids as the molecular link between colonic bacteria and the host. Curr Opin Pharmacol, 2013. 13(6): p. 869-74.

186. Lallemand, F., et al., Direct inhibition of the expression of cyclin D1 gene by sodium butyrate. Biochem Biophys Res Commun, 1996. 229(1): p. 163-9.

187. Siavoshian, S., et al., Butyrate and trichostatin A effects on the proliferation/differentiation of human intestinal epithelial cells: induction of cyclin D3 and p21 expression. Gut, 2000. 46(4): p. 507-14.

188. Park, J., et al., Short-chain fatty acids induce both effector and regulatory T cells by suppression of histone deacetylases and regulation of the mTOR-S6K pathway. Mucosal Immunol, 2015. 8(1): p. 80-93.

189. Schulthess, J., et al., The Short Chain Fatty Acid Butyrate Imprints an Antimicrobial Program in Macrophages. Immunity, 2019.

190. Bailon, E., et al., Butyrate in vitro immune-modulatory effects might be mediated through a proliferation-related induction of apoptosis. Immunobiology, 2010. 215(11): p. 863-73.

191. Fontenelle, B. and K.M. Gilbert, n-Butyrate anergized effector CD4+ T cells independent of regulatory T cell generation or activity. Scand J Immunol, 2012. 76(5): p. 457-63.

118

192. Zimmerman, M.A., et al., Butyrate suppresses colonic inflammation through HDAC1- dependent Fas upregulation and Fas-mediated apoptosis of T cells. Am J Physiol Gastrointest Liver Physiol, 2012. 302(12): p. G1405-15.

193. Chang, P.V., et al., The microbial metabolite butyrate regulates intestinal macrophage function via histone deacetylase inhibition. Proc Natl Acad Sci U S A, 2014. 111(6): p. 2247-52.

194. Arpaia, N., et al., Metabolites produced by commensal bacteria promote peripheral regulatory T-cell generation. Nature, 2013. 504(7480): p. 451-5.

195. Furusawa, Y., et al., Commensal microbe-derived butyrate induces the differentiation of colonic regulatory T cells. Nature, 2013. 504(7480): p. 446-50.

196. Kespohl, M., et al., The Microbial Metabolite Butyrate Induces Expression of Th1- Associated Factors in CD4(+) T Cells. Front Immunol, 2017. 8: p. 1036.

197. Salkowska, A., et al., Differentiation stage-specific effect of histone deacetylase inhibitors on the expression of RORgammaT in human lymphocytes. J Leukoc Biol, 2017. 102(6): p. 1487-1495.

198. Rosenbaum, D.M., S.G. Rasmussen, and B.K. Kobilka, The structure and function of G-protein-coupled receptors. Nature, 2009. 459(7245): p. 356-63.

199. Husted, A.S., et al., GPCR-Mediated Signaling of Metabolites. Cell Metab, 2017. 25(4): p. 777-796.

200. Brown, A.J., et al., The Orphan G protein-coupled receptors GPR41 and GPR43 are activated by propionate and other short chain carboxylic acids. J Biol Chem, 2003. 278(13): p. 11312-9.

201. Kim, M.H., et al., Short-chain fatty acids activate GPR41 and GPR43 on intestinal epithelial cells to promote inflammatory responses in mice. Gastroenterology, 2013. 145(2): p. 396-406 e1-10.

202. Kim, S., et al., Perspectives on the therapeutic potential of short-chain fatty acid receptors. BMB Reports, 2014. 47(3): p. 173-178.

203. Thangaraju, M., et al., GPR109A is a G-protein-coupled receptor for the bacterial fermentation product butyrate and functions as a tumor suppressor in colon. Cancer Res, 2009. 69(7): p. 2826-32.

204. Singh, N., et al., Activation of Gpr109a, receptor for niacin and the commensal metabolite butyrate, suppresses colonic inflammation and carcinogenesis. Immunity, 2014. 40(1): p. 128-39.

205. Kasubuchi, M., et al., Dietary gut microbial metabolites, short-chain fatty acids, and host metabolic regulation. Nutrients, 2015. 7(4): p. 2839-49.

206. Layden, B.T., et al., Short chain fatty acids and their receptors: new metabolic targets. Transl Res, 2013. 161(3): p. 131-40.

119

207. Alex, S., et al., Short-chain fatty acids stimulate angiopoietin-like 4 synthesis in human colon adenocarcinoma cells by activating peroxisome proliferator-activated receptor gamma. Mol Cell Biol, 2013. 33(7): p. 1303-16.

208. Byndloss, M.X., et al., Microbiota-activated PPAR-gamma signaling inhibits dysbiotic Enterobacteriaceae expansion. Science, 2017. 357(6351): p. 570-575.

209. Peng, L., et al., Butyrate enhances the intestinal barrier by facilitating tight junction assembly via activation of AMP-activated protein kinase in Caco-2 cell monolayers. J Nutr, 2009. 139(9): p. 1619-25.

210. Kalina, U., et al., Enhanced production of IL-18 in butyrate-treated intestinal epithelium by stimulation of the proximal promoter region. Eur J Immunol, 2002. 32(9): p. 2635- 43.

211. Zhang, M., et al., Butyrate inhibits interleukin-17 and generates Tregs to ameliorate colorectal colitis in rats. BMC Gastroenterol, 2016. 16(1): p. 84.

212. Kurita-Ochiai, T., K. Ochiai, and K. Fukushima, Butyric acid-induced T-cell apoptosis is mediated by caspase-8 and -9 activation in a Fas-independent manner. Clin Diagn Lab Immunol, 2001. 8(2): p. 325-32.

213. Kurita-Ochiai, T. and K. Ochiai, Butyric acid induces apoptosis via oxidative stress in Jurkat T-cells. J Dent Res, 2010. 89(7): p. 689-94.

214. Kurita-Ochiai, T., K. Fukushima, and K. Ochiai, Butyric acid-induced apoptosis of murine thymocytes, splenic T cells, and human Jurkat T cells. Infect Immun, 1997. 65(1): p. 35-41.

215. Smith, P.M., et al., The microbial metabolites, short-chain fatty acids, regulate colonic Treg cell homeostasis. Science, 2013. 341(6145): p. 569-73.

216. Cao, T., et al., The epigenetic modification during the induction of Foxp3 with sodium butyrate. Immunopharmacol Immunotoxicol, 2018: p. 1-10.

217. Luu, M., et al., Regulation of the effector function of CD8(+) T cells by gut microbiota- derived metabolite butyrate. Sci Rep, 2018. 8(1): p. 14430.

218. Kim, M., et al., Gut Microbial Metabolites Fuel Host Antibody Responses. Cell Host Microbe, 2016. 20(2): p. 202-14.

219. Kim, C.H., B cell-helping functions of gut microbial metabolites. Microb Cell, 2016. 3(10): p. 529-531.

220. Vinolo, M.A., et al., Suppressive effect of short-chain fatty acids on production of proinflammatory mediators by neutrophils. J Nutr Biochem, 2011. 22(9): p. 849-55.

221. Nascimento, C.R., et al., The short chain fatty acid sodium butyrate regulates the induction of CD1a in developing dendritic cells. Immunobiology, 2011. 216(3): p. 275- 84.

120

222. Nastasi, C., et al., Butyrate and propionate inhibit antigen-specific CD8(+) T cell activation by suppressing IL-12 production by antigen-presenting cells. Sci Rep, 2017. 7(1): p. 14516.

223. Ji, J., et al., Microbial metabolite butyrate facilitates M2 macrophage polarization and function. Sci Rep, 2016. 6: p. 24838.

224. Kaisar, M.M.M., et al., Butyrate Conditions Human Dendritic Cells to Prime Type 1 Regulatory T Cells via both Histone Deacetylase Inhibition and G Protein-Coupled Receptor 109A Signaling. Front Immunol, 2017. 8: p. 1429.

225. Berndt, B.E., et al., Butyrate increases IL-23 production by stimulated dendritic cells. Am J Physiol Gastrointest Liver Physiol, 2012. 303(12): p. G1384-92.

226. Kim, S.H., et al., Microbiota-derived butyrate suppresses group 3 innate lymphoid cells in terminal ileal Peyer's patches. Sci Rep, 2017. 7(1): p. 3980.

227. Thio, C.L., et al., Regulation of type 2 innate lymphoid cell-dependent airway hyperreactivity by butyrate. J Allergy Clin Immunol, 2018. 142(6): p. 1867-1883 e12.

228. Louis, P., G.L. Hold, and H.J. Flint, The gut microbiota, bacterial metabolites and colorectal cancer. Nat Rev Microbiol, 2014. 12(10): p. 661-72.

229. Dillon, S.M., et al., Low abundance of colonic butyrate-producing bacteria in HIV infection is associated with microbial translocation and immune activation. AIDS, 2017. 31(4): p. 511-521.

230. Steele, A.K., et al., Microbial exposure alters HIV-1-induced mucosal CD4+ T cell death pathways Ex vivo. Retrovirology, 2014. 11: p. 14.

231. Dillon, S.M., et al., HIV-1 infection of human intestinal lamina propria CD4+ T cells in vitro is enhanced by exposure to commensal Escherichia coli. J Immunol, 2012. 189(2): p. 885-96.

232. Rigby, L., et al., Methods for the analysis of histone H3 and H4 acetylation in blood. Epigenetics, 2012. 7(8): p. 875-82.

233. Chaudhuri, R., et al., Downregulation of CD4 by human immunodeficiency virus type 1 Nef is dependent on clathrin and involves direct interaction of Nef with the AP2 clathrin adaptor. J Virol, 2007. 81(8): p. 3877-90.

234. Kim, C.J., et al., Mucosal Th17 Cell Function Is Altered during HIV Infection and Is an Independent Predictor of Systemic Immune Activation. J Immunol, 2013. 191(5): p. 2164-73.

235. Critchfield, J.W., et al., Magnitude and complexity of rectal mucosa HIV-1-specific CD8+ T-cell responses during chronic infection reflect clinical status. PLoS One, 2008. 3(10): p. e3577.

121

236. Epple, H.J., et al., Acute HIV infection induces mucosal infiltration with CD4+ and CD8+ T cells, epithelial apoptosis, and a mucosal barrier defect. Gastroenterology, 2010. 139(4): p. 1289-300.

237. Shacklett, B.L., et al., Trafficking of human immunodeficiency virus type 1-specific CD8+ T cells to gut-associated lymphoid tissue during chronic infection. J Virol, 2003. 77(10): p. 5621-31.

238. Allers, K., et al., Macrophages accumulate in the gut mucosa of untreated HIV-infected patients. J Infect Dis, 2014. 209(5): p. 739-48.

239. Favre, D., et al., Tryptophan catabolism by indoleamine 2,3-dioxygenase 1 alters the balance of TH17 to regulatory T cells in HIV disease. Sci Transl Med, 2010. 2(32): p. 32ra36.

240. Mela, C.M., et al., Depletion of natural killer cells in the colonic lamina propria of viraemic HIV-1-infected individuals. AIDS, 2007. 21(16): p. 2177-82.

241. Sips, M., et al., Altered distribution of mucosal NK cells during HIV infection. Mucosal Immunol, 2012. 5(1): p. 30-40.

242. McGowan, I., et al., Increased HIV-1 mucosal replication is associated with generalized mucosal cytokine activation. J Acquir Immune Defic Syndr, 2004. 37(2): p. 1228-36.

243. Voigt, R.M., et al., HIV-associated mucosal gene expression: region-specific alterations. AIDS, 2015. 29(5): p. 537-46.

244. Epple, H.J., et al., Impairment of the intestinal barrier is evident in untreated but absent in suppressively treated HIV-infected patients. Gut, 2009. 58(2): p. 220-7.

245. Keating, J., et al., Intestinal absorptive capacity, intestinal permeability and jejunal histology in HIV and their relation to diarrhoea. Gut, 1995. 37(5): p. 623-9.

246. Sankaran, S., et al., Rapid onset of intestinal epithelial barrier dysfunction in primary human immunodeficiency virus infection is driven by an imbalance between immune response and mucosal repair and regeneration. J Virol, 2008. 82(1): p. 538-45.

247. Marchetti, G., C. Tincati, and G. Silvestri, Microbial translocation in the pathogenesis of HIV infection and AIDS. Clin Microbiol Rev, 2013. 26(1): p. 2-18.

248. Zevin, A.S., et al., Microbial translocation and microbiome dysbiosis in HIV-associated immune activation. Curr Opin HIV AIDS, 2016. 11(2): p. 182-90.

249. Kaiko, G.E. and T.S. Stappenbeck, Host-microbe interactions shaping the gastrointestinal environment. Trends Immunol, 2014. 35(11): p. 538-48.

250. Dinh, D.M., et al., Intestinal Microbiota, Microbial Translocation, and Systemic Inflammation in Chronic HIV Infection. J Infect Dis, 2014.

122

251. Ellis, C.L., et al., Molecular characterization of stool microbiota in HIV-infected subjects by panbacterial and order-level 16S ribosomal DNA (rDNA) quantification and correlations with immune activation. J Acquir Immune Defic Syndr, 2011. 57(5): p. 363- 70.

252. Gori, A., et al., Early impairment of gut function and gut flora supporting a role for alteration of gastrointestinal mucosa in human immunodeficiency virus pathogenesis. J Clin Microbiol, 2008. 46(2): p. 757-8.

253. Lozupone, C.A., et al., HIV-induced alteration in gut microbiota: driving factors, consequences, and effects of antiretroviral therapy. Gut Microbes, 2014. 5(4): p. 562- 70.

254. McHardy, I.H., et al., HIV infection is associated with compositional and functional shifts in the rectal mucosal microbiota. Microbiome, 2013. 1: p. 26.

255. Vazquez-Castellanos, J.F., et al., Altered metabolism of gut microbiota contributes to chronic immune activation in HIV-infected individuals. Mucosal Immunol, 2014.

256. Dillon, S.M., et al., Enhancement of HIV-1 infection and intestinal CD4+ T cell depletion ex vivo by gut microbes altered during chronic HIV-1 infection. Retrovirology, 2016. 13(1): p. 5.

257. Macfarlane, S. and G.T. Macfarlane, Regulation of short-chain fatty acid production. Proc Nutr Soc, 2003. 62(1): p. 67-72.

258. Hamer, H.M., et al., Review article: the role of butyrate on colonic function. Aliment Pharmacol Ther, 2008. 27(2): p. 104-19.

259. Segain, J.P., et al., Butyrate inhibits inflammatory responses through NFkappaB inhibition: implications for Crohn's disease. Gut, 2000. 47(3): p. 397-403.

260. Van Immerseel, F., et al., Butyric acid-producing anaerobic bacteria as a novel probiotic treatment approach for inflammatory bowel disease. J Med Microbiol, 2010. 59(Pt 2): p. 141-3.

261. Vieira, E.L., et al., Oral administration of sodium butyrate attenuates inflammation and mucosal lesion in experimental acute ulcerative colitis. J Nutr Biochem, 2012. 23(5): p. 430-6.

262. Meijer, K., P. de Vos, and M.G. Priebe, Butyrate and other short-chain fatty acids as modulators of immunity: what relevance for health? Curr Opin Clin Nutr Metab Care, 2010. 13(6): p. 715-21.

263. Saemann, M.D., et al., Anti-inflammatory effects of sodium butyrate on human monocytes: potent inhibition of IL-12 and up-regulation of IL-10 production. FASEB J, 2000. 14(15): p. 2380-2.

264. Hague, A., A.J. Butt, and C. Paraskeva, The role of butyrate in human colonic epithelial cells: an energy source or inducer of differentiation and apoptosis? Proc Nutr Soc, 1996. 55(3): p. 937-43.

123

265. Lewis, K., et al., Enhanced translocation of bacteria across metabolically stressed epithelia is reduced by butyrate. Inflamm Bowel Dis, 2010. 16(7): p. 1138-48.

266. Ploger, S., et al., Microbial butyrate and its role for barrier function in the gastrointestinal tract. Ann N Y Acad Sci, 2012. 1258: p. 52-9.

267. Wong, J.M., et al., Colonic health: fermentation and short chain fatty acids. J Clin Gastroenterol, 2006. 40(3): p. 235-43.

268. Gibson, P.R., et al., Colonic epithelial cell activation and the paradoxical effects of butyrate. Carcinogenesis, 1999. 20(4): p. 539-44.

269. Wang, H.B., et al., Butyrate enhances intestinal epithelial barrier function via upregulation of tight junction protein Claudin-1 transcription. Dig Dis Sci, 2012. 57(12): p. 3126-35.

270. Rosignoli, P., et al., Protective activity of butyrate on hydrogen peroxide-induced DNA damage in isolated human colonocytes and HT29 tumour cells. Carcinogenesis, 2001. 22(10): p. 1675-80.

271. Canani, R.B., et al., Potential beneficial effects of butyrate in intestinal and extraintestinal diseases. World J Gastroenterol, 2011. 17(12): p. 1519-28.

272. Giannini, G., et al., Histone deacetylase inhibitors in the treatment of cancer: overview and perspectives. Future Med Chem, 2012. 4(11): p. 1439-60.

273. Cavaglieri, C.R., et al., Differential effects of short-chain fatty acids on proliferation and production of pro- and anti-inflammatory cytokines by cultured lymphocytes. Life Sci, 2003. 73(13): p. 1683-90.

274. Deeks, S.G., R. Tracy, and D.C. Douek, Systemic effects of inflammation on health during chronic HIV infection. Immunity, 2013. 39(4): p. 633-45.

275. Lederman, M.M., et al., Residual immune dysregulation syndrome in treated HIV infection. Adv Immunol, 2013. 119: p. 51-83.

276. Guglani, L. and S.A. Khader, Th17 cytokines in mucosal immunity and inflammation. Curr Opin HIV AIDS, 2010. 5(2): p. 120-7.

277. Baecher-Allan, C., V. Viglietta, and D.A. Hafler, Human CD4+CD25+ regulatory T cells. Semin Immunol, 2004. 16(2): p. 89-98.

278. Rada-Iglesias, A., et al., Butyrate mediates decrease of histone acetylation centered on transcription start sites and down-regulation of associated genes. Genome Res, 2007. 17(6): p. 708-19.

279. Penn, M.L., et al., CXCR4 utilization is sufficient to trigger CD4+ T cell depletion in HIV-1-infected human lymphoid tissue. Proc Natl Acad Sci U S A, 1999. 96(2): p. 663- 8.

124

280. Bleul, C.C., et al., The HIV coreceptors CXCR4 and CCR5 are differentially expressed and regulated on human T lymphocytes. Proc Natl Acad Sci U S A, 1997. 94(5): p. 1925-30.

281. Imai, K., et al., Reactivation of latent HIV-1 by a wide variety of butyric acid-producing bacteria. Cell Mol Life Sci, 2012. 69(15): p. 2583-92.

282. Zack, J.A., et al., HIV-1 entry into quiescent primary lymphocytes: molecular analysis reveals a labile, latent viral structure. Cell, 1990. 61(2): p. 213-22.

283. Serrano-Villar, S., et al., The effects of prebiotics on microbial dysbiosis, butyrate production and immunity in HIV-infected subjects. Mucosal Immunol, 2017. 10(5): p. 1279-1293.

284. Imam, T., et al., Effector T Helper Cell Subsets in Inflammatory Bowel Diseases. Front Immunol, 2018. 9: p. 1212.

285. Hunt, P.W., et al., Relationship between T cell activation and CD4+ T cell count in HIV- seropositive individuals with undetectable plasma HIV RNA levels in the absence of therapy. J Infect Dis, 2008. 197(1): p. 126-33.

286. Stevenson, M., et al., HIV-1 replication is controlled at the level of T cell activation and proviral integration. EMBO J, 1990. 9(5): p. 1551-60.

287. Mahalingam, M., et al., T cell activation and disease severity in HIV infection. Clin Exp Immunol, 1993. 93(3): p. 337-43.

288. Benito, J.M., et al., Quantitative alterations of the functionally distinct subsets of CD4 and CD8 T lymphocytes in asymptomatic HIV infection: changes in the expression of CD45RO, CD45RA, CD11b, CD38, HLA-DR, and CD25 antigens. J Acquir Immune Defic Syndr Hum Retrovirol, 1997. 14(2): p. 128-35.

289. Juffermans, N.P., et al., Up-regulation of HIV coreceptors CXCR4 and CCR5 on CD4(+) T cells during human endotoxemia and after stimulation with (myco)bacterial antigens: the role of cytokines. Blood, 2000. 96(8): p. 2649-54.

290. Yang, H., et al., CD147 modulates the differentiation of T-helper 17 cells in patients with rheumatoid arthritis. APMIS, 2017. 125(1): p. 24-31.

291. Das, B., et al., Short chain fatty acids potently induce latent HIV-1 in T-cells by activating P-TEFb and multiple histone modifications. Virology, 2015. 474: p. 65-81.

292. Ganji-Arjenaki, M. and M. Rafieian-Kopaei, Probiotics are a good choice in remission of inflammatory bowel diseases: A meta analysis and systematic review. J Cell Physiol, 2018. 233(3): p. 2091-2103.

293. Scheppach, W., et al., Effect of butyrate enemas on the colonic mucosa in distal ulcerative colitis. Gastroenterology, 1992. 103(1): p. 51-6.

294. Steinhart, A.H., et al., Treatment of left-sided ulcerative colitis with butyrate enemas: a controlled trial. Aliment Pharmacol Ther, 1996. 10(5): p. 729-36.

125

295. Lei, J., et al., Peroxisome proliferator-activated receptor alpha and gamma agonists together with TGF-beta convert human CD4+CD25- T cells into functional Foxp3+ regulatory T cells. J Immunol, 2010. 185(12): p. 7186-98.

296. Foryst-Ludwig, A., et al., PPARgamma activation attenuates T-lymphocyte-dependent inflammation of adipose tissue and development of insulin resistance in obese mice. Cardiovasc Diabetol, 2010. 9: p. 64.

297. Bai, B., et al., CBP/p300 inhibitor C646 prevents high glucose exposure induced neuroepithelial cell proliferation. Birth Defects Res, 2018. 110(14): p. 1118-1128.

298. Ng, W.F., et al., Human CD4(+)CD25(+) cells: a naturally occurring population of regulatory T cells. Blood, 2001. 98(9): p. 2736-44.

299. Sathaliyawala, T., et al., Distribution and compartmentalization of human circulating and tissue-resident memory T cell subsets. Immunity, 2013. 38(1): p. 187-97.

300. Clark, R.A., Resident memory T cells in human health and disease. Sci Transl Med, 2015. 7(269): p. 269rv1.

301. D'Souza, W.N., et al., Differing roles for short chain fatty acids and GPR43 agonism in the regulation of intestinal barrier function and immune responses. PLoS One, 2017. 12(7): p. e0180190.

302. Bohan, C., D. York, and A. Srinivasan, Sodium butyrate activates human immunodeficiency virus long terminal repeat--directed expression. Biochem Biophys Res Commun, 1987. 148(3): p. 899-905.

303. Segal, L.N., et al., Anaerobic Bacterial Fermentation Products Increase Tuberculosis Risk in Antiretroviral-Drug-Treated HIV Patients. Cell Host Microbe, 2017. 21(4): p. 530-537 e4.

304. Eeckhaut, V., et al., Butyricicoccus pullicaecorum in inflammatory bowel disease. Gut, 2013. 62(12): p. 1745-52.

305. Glueck, B., Y. Han, and G.A.M. Cresci, Tributyrin Supplementation Protects Immune Responses and Vasculature and Reduces Oxidative Stress in the Proximal Colon of Mice Exposed to Chronic-Binge Ethanol Feeding. J Immunol Res, 2018. 2018: p. 9671919.

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APPENDIX A

Colorado Clinical and Translational Sciences Institute (CCTSI) TL1 Fellowship Clinical

Experience

The Colorado Clinical and Translational Sciences Institute (CCTSI) offers several one- year TL1 fellowships to students carrying out basic science research on the Anschutz Medical

Campus. This fellowship aims to promote the integration and interface of basic science research and clinical medicine at the predoctoral level. As a TL1 fellowship recipient, we were expected to attend monthly TL1 journal clubs and guest presentations as well as complete a full day of shadowing in the clinic per month. Also, fellowship recipients are required to attend the national TL1 meeting and complete required coursework in biostatistics, epidemiology, and disease-focused basic science courses. After the completion of the requirements and thesis defense, each fellowship recipient will be awarded a certificate acknowledging the successful completion of the fellowship.

As a CCTSI TL1 fellowship recipient for the 2017-2018 year, I completed the following course work: Tissue Biology and Disease Mechanisms (IDPT 7646), Statistics for the Basic

Sciences (BIOS 6606) and Epidemiology (EPID 6630). As a medical scientist training program

(MSTP) student I also completed pre-clinical coursework in human anatomy, physiology and disease pathophysiology that helped fulfill these coursework requirements. As part of the TL1 fellowship, I presented my data at the 2018 National TL1 conference in Washington, D.C. in the form of a poster titled “The microbial-derived short-chain fatty acid butyrate directly and differentially inhibits gut T helper cell subset activation and proliferation.” While at the conference, I was able to attend several talks and information sessions about career opportunities, networking, and translational science. Also, the conference allowed several opportunities for networking with individuals at various career levels, including predoctoral and postdoctoral candidates as well as several senior investigators and mentors. One of the most

127 impactful experiences of the conference was the opportunity to go to the offices of the state of Colorado representatives in the House and Senate to discuss the importance of translational research and continued funding for both the NIH and the TL1 program with them.

Lastly, my clinical experience was carried out with Dr. Jose Castillo-Mancilla at the University of Colorado Hospital in Aurora, Colorado.

The basis of my thesis project was formed around the observation that during HIV-1 infection the breakdown of the gut barrier is associated with chronic immune activation which is a linked to increased morbidity and mortality[101, 252, 278, 310]. To better understand the potential clinical relevance and implications of my thesis project, I was mentored by Dr. José

Castillo-Mancilla. Due to the impact of systemic inflammation and immune activation in

PLWH, the goal of my shadowing experience with Dr. Castillo-Mancilla was to observe him during his weekly inpatient clinic, at the University of Colorado Hospital where he currently serves a patient population that includes 80 PLWH. Many of these individuals whom I encountered in the clinic had been chronically infected and have several non-AIDS related comorbidities such as neurocognitive decline, cardiovascular and metabolic diseases which are known to be associated with chronic immune activation and inflammation[101, 252, 278,

310]. During these shadowing opportunities, I observed Dr. Castillo-Mancilla manage his patients’ care and attended the visits of specific patients to gain a better understanding of their longitudinal care and how living with HIV-1 inflammation-associated comorbidities impacted their daily lives.

Aside from the clinical challenges that these individuals faced, I was also able to learn about their struggles with their diagnosis and the impact that having HIV-1 has had on their personal and professional lives. One of my most memorable experiences was interviewing a patient who had been living with HIV-1 since the early 1990s. This patient detailed the struggles of having HIV-1 before effective ART and the issues that some of the early ART regimes had. Also, they recounted losing numerous close friends to the disease as well as

128 their current struggles with conditions linked to chronic inflammation such as their struggle with metabolic and cardiovascular disease. While I had understood the clinical progression of

HIV-1 and the associated comorbidities, listening to an individual who had struggled with this disease for over twenty years, put the importance not only my work but all of the work in the

HIV-1 field into perspective. Moreover, this exposure will help tailor my research goals to be translationally relevant to the treatment and care those struggling with chronic HIV-1 infection and to be a proficient and understanding physician-scientist by not only understanding the scientific issues that need to be addressed to combat this disease but also the human connection required to provide adequate care.

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