TARGETING NEGATIVE REGULATORS OF IMMUNE SIGNALING PATHWAYS TO DISRUPT THE HIV-1 LIFE CYCLE

By

JARED PAUL TAYLOR

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2017

© 2017 Jared Paul Taylor

To the giants who came before us, on whose shoulders we stand

ACKNOWLEDGMENTS

I would like to thank my mentor Mark Wallet for giving me the opportunity to pursue my graduate research in his lab. He has been a wonderful mentor over the years and I have learned so much from him. I also would like to thank Maureen Goodenow,

Stephanie Karst, Naohiro Terada, and Edward Scott for serving on my advisory committee and for their support and advice. I would also like to thank all the Wallet lab members past and present who have been coworkers as well as friends for making the work environment enjoyable and collaborative.

I would also like to thank Dave Bloom for sponsoring me on the Infectious

Disease T32 training grant for three years. This afforded me many unique opportunities with funding for not only my graduate stipend and tuition, but also for supplies and traveling to conferences greatly enriching my graduate experience.

I want to also express my appreciation for all of the administrative staff that have helped with so many things through the years. The IDP office staff have been very helpful especially in the beginning of the program. I especially want to thank Karen Cox in the Pathology office for all of the many things she does for our lab. I also thank Kris

Minkoff and Debbie Burgess in the Microbiology and Molecular Genetics office for their support with things related to the Immunology and Microbiology concentration and the training grant.

I would also like to thank my family for their support throughout the years. I also thank my friends here in Gainesville who have made getting through the program much easier. Finally, I would like to thank my dog Winston who has been my constant loyal companion as I have gone through graduate school.

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TABLE OF CONTENTS

Page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 7

LIST OF FIGURES ...... 8

LIST OF ABBREVIATIONS ...... 10

ABSTRACT ...... 12

CHAPTER

1 BACKGROUND ...... 14

Discovery of HIV/AIDS ...... 14 Natural History of HIV/AIDS ...... 15 Where Do We Go from Here? ...... 16 HIV-1 Lifecycle ...... 17 Outcomes of the Life Cycle ...... 23 Immune Control of HIV-1 ...... 37

2 DEPLETION OF USP18 ENHANCES TYPE I IFN RESPONSIVENESS AND RESTRICTS HIV-1 INFECTION IN MACROPHAGES ...... 46

Introduction ...... 46 Methods and Materials...... 54 Results ...... 64 Discussion ...... 71 Figures ...... 78 Tables ...... 98

3 HARMINE BOOSTS REACTIVATION OF LATENT HIV-1 BY LATENCY REVERSING AGENTS ...... 100

Introduction ...... 100 Methods and Materials...... 105 Results ...... 108 Discussion ...... 112 Figures ...... 116 Tables ...... 130

4 CONCLUDING REMARKS ...... 131

5

LIST OF REFERENCES ...... 136

BIOGRAPHICAL SKETCH ...... 192

6

LIST OF TABLES

Table Page

2-1 Primer Sequences ...... 98

2-2 HIV-1-Induced Gene Expression ...... 99

3-1 PMA-Induced Gene Expression with Harmine...... 130

7

LIST OF FIGURES

Figure Page

2-1 Type I IFN restricts HIV-1 replication in TZM-bl cells ...... 78

2-2 IFNAR blocking inhibits HIV-1 replication ...... 79

2-3 HIV-1 induces an IFN-like response in MDMs ...... 80

2-4 USP18 expression is induced by HIV-1 and is dependent on type I IFN signaling ...... 81

2-5 USP18 knockdown by siRNA makes THP-1 cells and MDMs, but not TZM-bl cells, refractory to HIV-1 infection ...... 82

2-6 USP18 knockdown by shRNA in THP-1 cells ...... 84

2-7 shRNA knockdown of USP18 inhibits HIV-1 replication in THP-1 cells and enhances IFN-β-induced STAT activation ...... 85

2-8 siRNA knockdown of USP18 enhances STAT activation and modulates the transcriptome in IFN-β-treated MDMs ...... 87

2-9 ISG expression is enhanced in USP18 deficient THP-1 Cells and MDMs ...... 89

2-10 Workflow for generating iPSC-derived monocytes ...... 91

2-11 Characterization of iMacs ...... 92

2-12 iMacs support HIV-1 replication ...... 93

2-13 CRISPR/Cas9 knockout of USP18 in iPSCs ...... 94

2-14 USP18-/- iMacs have enhanced STAT phosphorylation ...... 95

2-15 Knockout of USP18 restricts HIV-1 replication in iMacs ...... 96

2-16 USP18 tunes the IFN response to allow for efficient HIV-1 replication ...... 97

3-1 Chemical structure of DYRK1A inhibitors ...... 116

3-2 DYRK1A inhibitors boost LRA-induced HIV-1 reactivation in 5A8 cells ...... 117

3-3 Harmine boosts clinically-relevant LRA-induced HIV-1 reactivation in 5A8 cells ...... 118

3-4 Harmine boosts the MFI of GFP induced by LRAs ...... 119

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3-5 Harmine boosts bryostatin-induced gag expression in J-Lat cells ...... 120

3-6 Harmine boosts LRA-induced gag expression in J-Lat cells ...... 121

3-7 Harmine boosts the magnitude of SAHA reactivation, but not the frequency of reactivated cells ...... 122

3-8 DYRK1A CRISPR knockout in 5A8 cells ...... 123

3-9 DYRK1A knockout in 5A8 cells does not boost PMA-induced reactivation of latent HIV-1...... 124

3-10 Harmine boosts PMA-induced phosphor-ERK1/2 expression ...... 125

3-11 MEK/ERK inhibitor abrogates effect of harmine in PMA treated 5A8 cells ...... 126

3-12 Jurkat T cells that were transduced with lentivirus NFκB or NFAT luciferase reporters or a negative control lentivirus were treated with different doses of PKC overnight ...... 127

3-13 Ionomycin induces NFAT activation in Jurkat luciferase reporter cell line ...... 128

3-14 Heat map of gene expression in 5A8 cells treated with PMA with or without harmine...... 129

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LIST OF ABBREVIATIONS

AIDS Acquired immune deficiency syndrome

ART Antiretroviral therapy

CRISPR Clustered regularly interspaced short palindromic repeats

CTL Cytotoxic T lymphocytes

DYRK1A Dual-specificity tyrosine phosphorylation-regulated kinase 1A

ELISA Enzyme-linked immunosorbent assay

GALT Gut-associated lymphoid tissue

HAART Highly active antiretroviral therapy

HDAC Histone deacetylase enzyme

HIV Human immunodeficiency virus

HTLV Human T-lymphotropic virus

IFN Interferon

IFNAR Interferon α receptor iMac Induced pluripotent stem cell-derived macrophage iPSC Induced pluripotent stem cell

ISG Interferon-stimulated gene

ISG15 Interferon-stimulated gene 15

LAV Lymphadenopathy associated virus

LRA Latency reversing agents

LTR Long terminal repeat

NFAT Nuclear factor of activated T cells

NFκB Nuclear factor kappa B

PKC Protein kinase C

PMA Phorbol 12-myristate 13-acetate

10

P-TEFb Positive transcription elongation factor b

RT Reverse transcriptase

TCID50 Tissue culture infectious dose 50

TCR T cell receptor

USP18 Ubiquitin-specific proteinase 18

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

TARGETING NEGATIVE REGULATORS OF IMMUNE SIGNALING PATHWAYS TO DISRUPT THE HIV-1 LIFE CYCLE

By

Jared Paul Taylor

December 2017

Chair: Mark A. Wallet Major: Medical Sciences – Immunology and Microbiology

Treatment with combination antiretroviral therapy (ART) halts the progression of acquired immunodeficiency syndrome (AIDS) in patients infected with human immunodeficiency virus 1 (HIV-1). HIV-1 infects long-lived memory CD4+ T cells and

macrophages and can remain latent in these cells. Because latency can be reversed in

these cells and viral replication can resume, patients must remain on ART for life. For a

sterilizing cure to be achieved, these permanently-infected cells must be eliminated.

The goal of this study was to identify host proteins that can be potential targets for

pharmaceutical interventions to augment existing antiretroviral therapies.

One approach to eliminating the latently infected T cells, known as the “shock and kill” approach, seeks to “shock” infected memory T cells with a stimulus that reactivates latent virus so that infected cells can be killed by the cytopathic effects of the

virus or immune surveillance. Existing drugs that reverse latency in infected T cells,

known as latency reversing agents (LRAs), have had some efficacy, but do not

reactivate all latently infected cells. Using J-Lat cells as a model for reactivation of latent

HIV-1 we identified a novel use for the naturally-occurring plant compound harmine,

which targets host protein dual-specificity tyrosine phosphorylation-regulated kinase 1A

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(DYRK1A), in boosting the efficacy of existing LRAs. Treatment with harmine in combination with PKC agonists increased the amount of two factors that promote HIV-1 transcription, such as nuclear factor kappa B and ERK1/2, and decreased the amount of

HEXIM1 expression a factor that negatively regulates HIV-1 transcription. Thus, harmine in combination with existing LRAs boosted both the frequency and magnitude of reactivation in our J-Lat model.

In macrophages, type 1 interferons (T1 IFNs) can restrict HIV-1 replication by inducing the expression of an antiviral transcriptional program. Expression of ubiquitin specific proteinase 18 (USP18), a negative regulator of T1 IFN signaling, is induced by

HIV-1 in macrophages. In USP18 knockout cell models, including a novel induced pluripotent stem cell derived macrophage model, HIV-1 replication was significantly restricted. HIV-1 restriction was mediated by increased T1 IFN signaling and increased expression of antiviral genes in the absence of the negative regulator USP18.

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CHAPTER 1 BACKGROUND

Discovery of HIV/AIDS

In the United States in the early 1980s, there were several cases of people with

no known causes of immunodeficiency presenting with diseases that were usually

predictive of cellular immunodeficiency, such as Kaposi’s sarcoma, Pneumocystis carinii

pneumonia, and undifferentiated non-Hodgkins lymphoma [1–5]. In 1982, the term

acquired immune deficiency syndrome (AIDS) was coined to describe these cases of

opportunistic infections. However, the cause of AIDS was yet to be determined.

In 1983, two research groups reported the cause of AIDS to be a retrovirus. Luc

Montagnier named the virus his group isolated lymphadenopathy-associated virus

(LAV) [6]. Robert Gallo’s group named their virus HTLV-III since it bore similarity to other HTLV family members previously described by his group [7]. It was later found that these two viruses were identical and they were named human immunodeficiency virus (HIV) [8, 9].

In 1986, Luc Montagnier’s group isolated a virus that was related to HIV from two

AIDS patients in West Africa and named this virus LAV-II [10, 11]. LAV-I/HTLV-III and

LAV-II have since been named HIV-1 and HIV-2, respectively. HIV-1 and HIV-2 both

originated in Africa, although it is unclear how the viruses were transmitted to humans.

HIV-1 is thought to have been transmitted to humans from chimpanzees and HIV-2 is

thought to have been transmitted to humans from sooty mangabey in West Africa based

on similarity to SIV strains that naturally infect these species [12–14]. There are four

lineages of HIV-1, groups M, N, O, and P, which are thought to represent four separate

cross-species transmission events [15]. Group M is the lineage that is responsible for

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the pandemic form that has spread around the world. Although it has not been shown conclusively, it is thought that the viruses were transmitted to humans through the hunting, butchering, and consumption of bushmeat [16].

Natural History of HIV/AIDS

It was recognized early in the AIDS epidemic that patients presenting with

opportunistic infections had impaired immune systems. Clinicians noted that patients

had decreased circulating CD4+ T cells and B-cell dysfunction [17–20]. Later in vitro

experiments would show that HIV-1 had a cytopathic effect on T cells [21], which

seemed to explain the source of the immunodeficiency. However, it would be many

years before the mechanism of HIV-1-induced CD4+ T cell death would be worked out.

After transmission of HIV-1 to a new host, there is a period of approximately 7-21

days after infection of the first cell to when the virus is detectable in the blood. This

period of time is referred to as the eclipse phase [22–27]. Studies with SIV in rhesus

macaques have shown that during this clinically silent phase the virus propagates

locally in CD4+ T cells until the viral population is large enough to disseminate and

establish infection in secondary lymphoid organs [28, 29]. After the eclipse phase, the

virus rapidly replicates in the secondary lymphoid tissues, especially in the gut-

associated lymphoid tissue (GALT) where CD4+ T cells are rapidly depleted [30–35].

Once the level of circulating CD4+ T cells has dropped below 200 cells/μl, a

patient is diagnosed with AIDS [36]. This is most certainly a death sentence for a patient

if left untreated. With the advent of highly active antiretroviral therapy (HAART) [37, 38]

there has been a significant decrease in opportunistic illnesses and AIDS-related deaths

among those on HAART [39]. However, treatment is required for life. Despite

undetectable viral loads during therapy [37, 38, 40], cessation of therapy results in

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rebound of viremia and the progression to AIDS resumes [41]. This is due to the

existence of viral reservoirs that can persist for life despite therapy. Indeed, CD4+ T cells harboring replication competent virus can be found in patients even on HAART

[42–44]. To date, a sterilizing cure for HIV-1 has been unobtainable.

Where Do We Go from Here?

Although lifelong treatment of patients with HAART (now called ART) can prevent

the development of AIDS and extend life expectancy, there are still non-AIDS related

morbidities associated with chronic HIV-1 infection. ART is not readily available to all in

every country. The high economic burden of lifelong therapy, as well as lack of patient

compliance, drives a need for a sterilizing cure and/or a highly effective preventative

vaccine. Despite much progress in the understanding of the molecular biology and

disease processes of HIV-1 in the past few decades, there are still significant

knowledge gaps.

The current priorities for HIV/AIDS research published by the Office of AIDS

Research (OAR) at the National Institutes of Health are to reduce the incidence of HIV,

to develop next-generation HIV therapies, to conduct research toward a cure, and to

research HIV-associated comorbidities, coinfections, and complications. Research

towards a cure aims to discover the mechanisms that regulate viral and host

interactions that contribute to HIV persistence, latency, and reservoir formation

Strategies to develop immune-based therapies to control or eliminate persistently or

latently infected cells in combination with ART are a major goal of the OAR.

The work presented herein contributes to the goal of eliminating persistently or

latently infected cells. Instead of developing new antivirals that target the virus, the

current work seeks to understand what host proteins are required for the HIV-1 life cycle

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and if the life cycle can be disrupted by targeting these proteins. The first project focuses on the role of host protein USP18 in HIV-1 persistence in macrophages. The second project focuses on a novel use of the plant compound harmine to boost reactivation of latently infected T cells.

HIV-1 Lifecycle

HIV-1 infects CD4+ T cells and macrophages (see below). The life cycle of HIV-1

has been well characterized since the discovery of the virus. A detailed description of

each stage of the HIV-1 life cycle is presented below followed by the causes of host

pathogenesis and the establishment of the viral reservoir.

Binding and entry. Like all retroviruses, HIV-1 is an enveloped virus that enters

the host cell by receptor-mediated fusion of the virus membrane with the host cell

membrane [45]. The transmembrane viral glycoprotein Env is made up of a trimer of

gp120 and gp41 heterodimers. Entry into host cells requires binding of Env to CD4, the

primary receptor for HIV, on the host cell [46–48]. Env also requires binding to a

coreceptor, CXCR4 or CCR5 [49–52]. Viruses can be classified based on coreceptor

usage: R5 viruses utilize CCR5, X4 viruses utilize CXCR4, and R5X4 viruses can utilize

both [53]. New infections are almost exclusively by R5 or R5X4 viruses [54]. After

binding of the receptor and coreceptor, the viral membrane fuses with the host cell

membrane allowing for entry of the virus into the cell [55].

Reverse transcription. HIV-1 has two copies of a single-stranded RNA genome.

A key characteristic of retroviruses is that their RNA genomes are reverse transcribed

into double-stranded DNA by viral reverse transcriptase (RT) [45]. After viral entry into the cells, RT begins synthesis of complementary DNA to the RNA template. RT uses

Lys host tRNA3 , which binds to the primer binding site on the HIV-1 genomic RNA to

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Lys prime synthesis of the (-) strand [56]. The tRNA3 is required for the switch from

transcriptional initiation to elongation and cannot be substituted with short RNA or DNA

primers [57, 58]. The reverse transcription step, which occurs in the cytoplasm, is an

important part of the HIV-1 lifecycle and has been the target of many antiretroviral drugs

[59].

There are two main classes of RT inhibitors: non-nucleoside reverse

transcriptase inhibitors (NNRTI) and nucleoside/nucleotide reverse transcriptase

inhibitors (NRTI). NNRTIs function by inhibiting the active site of the RT enzyme. NRTIs

function as analogs of nucleotides that get incorporated into DNA by HIV-1 RT, but lack

the 3’ hydroxyl group necessary for adding the next nucleotide. This halts the synthesis

of that strand of DNA. The first antiviral drug to be complete clinical trials for the

treatment of AIDS was a NRTI called azidothymidine (AZT) in 1987 [60]. Currently,

there are 23 FDA approved reverse transcriptase inhibitors on the market [59].

Integration. After reverse transcription, the linear, double-stranded DNA is

imported into the nucleus as part of the nucleoprotein complex called the pre-integration complex (PIC) [61]. In the nucleus, viral integrase catalyzes the integration of the proviral genome into the host genome [62]. Integration sites are not restricted to a particular DNA sequence and can be found throughout the genome on every chromosome [63–67], but integration does have a preference for actively transcribed regions of DNA [68–76]. It has also been shown that HIV integration preferentially occurs near histone modifications associated with active transcription, such as acetylation of H3 and H4 and H3K4 methylation [77].

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The integration target sites in CD4+ T cells may be different than those in

monocyte-derived macrophages with the latter being more restricted [78]. Considering that HIV integration prefers actively transcribed genes, it is not surprising that two different cell types would have different hotspots since the two cell types have differential gene expression. After integration, transcription is driven by host machinery that interacts with the viral LTR promoter. There are currently six integrase inhibitors on the market available as part of the antiretroviral therapy regimen: dolutegravir, elvitegravir, isentress, raltegravir, tivicay, and vitekta.

Transcription of viral mRNA. After integration, the proviral genome serves as a template for transcription of viral mRNA driven by the LTR promoter. However, the LTR promoter is not sufficient for complete transcription without the HIV-1-encoded trans

activating factor called Tat. Sodroski and colleagues showed that if you put a reporter

plasmid in cells with just the LTR promoter that transcription was very inefficient, but if

the cells were also infected with HIV-1 then the promoter was greater than 1000-fold more efficient due to the presence of some trans activating factor [79].

It was later discovered that there was a region of DNA just 3’ to the LTR that

required stimulation from the trans activating factor for gene expression to occur [80].

This region is now called the transactivation-responsive region (TAR), which only

functions when immediately downstream of the LTR in the correct orientation [81]. A few

of the inefficiently transcribed early transcripts will be completely spliced allowing for

nuclear export and translation of Tat. Tat can now return to the nucleus where Dingwall

and colleagues demonstrated that Tat directly binds to the TAR, which forms a U-rich

stem-loop structure [82, 83].

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Tat binding to TAR plays a direct role in promoting elongation rather than initiation of transcription [84]. Herrmann and colleagues found that a kinase, later identified as CDK9 [85], binds specifically to Tat [86, 87]. CDK9 is part of the P-TEFb complex, which phosphorylates the C-terminal domain (CTD) of the RNA polymerase II large subunit promoting mRNA elongation [88, 89].

Translation of viral proteins. The gag and pol genes have overlapping ends in the HIV genome with the 3’ end of the gag gene overlapping the 5’ end of the pol gene by about 205-241 nucleotides in different reading frames [56, 90–92]. In order for both

polyproteins to be translated from the same mRNA, HIV-1 utilizes a frameshifting mechanism that depends on a U-rich region that causes a slippage of the ribosome

[93]. The resulting gene product is a Gag-Pol fusion protein that is produced at approximately a 1:20 ratio with Gag alone [94].

This ratio is important so that there are enough viral enzymes incorporated into

the new viruses. Virus-like particles can form and bud without Gag-Pol, but they are

non-infectious [95]. Gag-Pol only particles are also defective in that they have

premature activation of the protease activity in the producer cells that inhibits assembly

and budding of new virions [96–98]. It has been experimentally demonstrated that even

the slightest change in the ratio between Gag and Gag-Pol molecules can have an

effect on viral assembly and infectivity [99].

The Env protein is translated from incompletely spliced mRNAs that contain the

rev response element (RRE), which require the HIV-1 protein Rev for nuclear export.

Env is translated on the rough endoplasmic reticulum and contains an endoplasmic

reticulum signal sequence at its N-terminus that targets Env to the rough ER membrane

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[100, 101]. The transmembrane Env protein is trafficked to the plasma membrane through the Golgi apparatus and secretory pathway [102].

Viral assembly, budding, and maturation. After translation, Gag polyproteins traffic to the plasma membrane where assembly into virions and budding will occur. The matrix protein in Gag (MA) has the ability to bind to lipids in plasma membranes as well as nucleic acids [103]. It has been demonstrated that HIV virions are enriched in phosphoinositides [104], which is a result of basic residues in the MA protein that promote binding of Gag to phosphatidylinositol (4,5) bisphosphate (PI(4,5)P2 [105, 106].

After binding to PI(4,5)P2, a myristoyl group on the MA protein is exposed due to a

conformational change and inserts into the plasma membrane facilitating anchoring of

Gag to the plasma membrane [107–109].

Unspliced viral mRNAs that have exported from the nucleus dimerize through the

interaction of two mRNA molecules at the dimer initiation site located in the 5’ UTR of

gag [110, 111]. These dimerized mRNA molecules interact with the nucleocapsid region

of the Gag protein that is embedded in the membrane thereby directing the genomic

RNA to the site of virion assembly [112].

Viral budding begins spontaneously at the plasma membrane after enough Gag

particles have assembled from the cytoplasm. It was once believed that viruses

assembled in the endosomal compartment, but Jouvenet and colleagues demonstrated

that if late endosomal motility was blocked that HIV could still assemble and be released

from the cells [113]. Using real-time imaging with fluorescence resonance energy

transfer, fluorescence recovery after photobleaching, and total-internal-reflection

fluorescent microscopy, Jouvenet and colleagues could demonstrate in live cells the

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real-time kinetics of Gag assembly and budding. They showed that each cell would have between 50-150 distinct puncta that would form at the plasma membrane with approximately 5-6 minutes required for the complete assembly of one virion [114]. This

work was confirmed by Ivanchenko and colleagues that determined approximately 8-9

minutes was required for assembly of individual particles with ~1,500±700 seconds

required for release of particles from the cell [115].

Mutational studies of the p6 domain of Gag have revealed its essential role in

budding [116]. There is a Pro-Thr-Ala-Pro or PTAP domain that is essential for budding.

Mutations to these amino acids arrest the viral lifecycle at this late stage, hence the

name “late domain” [117]. It would later be demonstrated by several groups that the

PTAP domain was so important to the budding process because it recruits TSG101,

which is part of the ESCRT-I complex [118–121]. These findings established the role

that ESCRT machinery played in HIV-1 budding. Additional studies would show that although the recruitment of Tsg101 was necessary that it was not sufficient on its own, suggesting that another factor was required [122].

It would later be shown that the p6 late domain YPXL also plays a role in the recruitment of host factors. The YPXL domain has been shown to recruit an ESCRT protein called ALIX [123–126]. Mutations disrupting the binding of ALIX to the YPXL domain results in a two-fold reduction in viral replication in a HeLa reporter cell line called TZM-bl [127].

The interactions of PTAP and YPXL with TSG101 and ALIX are essential for initiation of the recruitment of ESCRT-III proteins. CHMP2 and CHMP4, members of the

ESCRT-III family have been identified as the most critical ESCRT-III proteins required

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for HIV-1 budding [128]. These interactions then promote the recruitment of the remaining ESCRT factors needed for pinching off the viral membrane from the host cell membrane, thus facilitating the final step of budding.

Final maturation of the budded retroviruses like HIV-1 is dependent upon viral protease activity; without Gag cleavage, the viruses are non-infectious [129–134]. Even

partial inhibition of HIV-1 protease is sufficient to disrupt virus assembly and results in

non-infectious viral particles [135]. As such, HIV-1 protease has become an important and effective target of antiretroviral drugs. There are currently at least ten FDA approved drugs that target HIV-1 protease [59].

Outcomes of the Life Cycle

There are three potential fates for the HIV-1 life cycle in a cell that contribute differently to pathogenesis. They are listed here, but are described in more detail under

HIV-1 Pathogenesis below:

1) HIV-1 completes the life cycle. HIV-1 can complete the lifecycle and new virions are produced. These new virions are important for propagation and dissemination in the host as well as for reseeding the viral reservoir. Free virions are also what allows the virus to be transmitted to new hosts.

2) Abortive infection of non-permissive cells. Non-permissive CD4+ T cells

that are in a resting state are infected with the virus, but the virus is unable to complete

its lifecycle. The viral products are detected by the cell, which undergoes a programmed

cell death. Although this stops the virus from propagating and spreading from these

cells, this is the direct cause of CD4+ T cell depletion that results in AIDS and the host’s

susceptibility to opportunistic infections.

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3) Viral latency. Viral latency is characterized by the integration of the virus

without expression of viral products from the viral promoter. Latent viruses can remain

in long-lived memory cells and can be induced at a future time to resume the HIV-1

lifecycle if the correct stimulus is present to reactivate the latently-infected cells. These latently infected cells make up most of the viral reservoir, which must be eliminated or controlled for sterilizing or functional cure, respectively, to be achieved.

Transmission and dissemination

Transmission. HIV-1 most commonly is transmitted sexually as it requires entry through a mucosal surface. However, it can also be transmitted through mucosal membrane exposure to contaminated body fluids, needle stick, and mother to fetus transmission [136]. Although there is likely a diverse quasispecies in the inoculum during transmission to a new host, studies have shown that there is a bottleneck where only one or a few founder viruses are responsible for the viral outgrowth following transmission [54, 137]. Acute infections are essentially not genetically diverse. Derdeyn and colleagues demonstrated that the viruses found in newly infected individuals were sensitive to neutralization from antibodies from the transmitting sexual partner [137].

This implies that characteristics needed for transmission are not selected for strongly in chronically infected patients.

Founder viruses that establish new infections are usually R5 or R5X4 viruses and not X4 viruses and have the ability to infect macrophages [138–142]. However, this is not because X4 variants are not present. More recent studies utilizing ultra-deep sequencing methods have shown that there is a greater number of X4 present in newly infected patients than was once believed, but that over the first 1-2 years of infection these populations dwindle and the R5 population becomes dominant [143, 144]. The X4

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viruses reemerge later in chronically infected patients and is a marker for immunodeficiency [145–147].

This correlation supports a model that proposes that CXCR4 using viruses are more susceptible to immune control since it is only seen in immunocompromised patients [148, 149]. If mutations in the V3 region of Env, which determines cell tropism, were truly random then there wouldn’t be such a strong tendency towards synonymous mutations that don’t change the tropism. Rather, there would be an equal ratio of synonymous to non-synonymous mutations [150]. Furthermore, this strong selection against CXCR4 usage wouldn’t suddenly disappear during the course of disease progression at the same time as the immune system begins to be compromised [151].

CXCR4 usage requires an open conformation of the HIV-1 Env V3 region that is potentially more susceptible to neutralizing antibodies [152–154] and possibly T cell- mediated immunity as well [155]. Therefore, in an immunocompetent host there is selective pressure against X4 variants, but after becoming immunocompromised, there is no selection against X4 variants.

Dissemination. Studies with SIV in the macaque model have demonstrated that founder viruses establish infection in activated and resting CD4+ T cells that reside in

the mucosal site of infection [156] before moving to lymphoid organs and eventual

systemic dissemination [28]. CCR5 expressing T cells, mainly memory CD4+ T cells, are

the primary targets of infection [157–160]. HIV-1 establishes robust infection in the

GALT where there is an abundance of CCR5 expressing memory CD4+ T cells [161,

162].

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Abortive infection and CD4+ T cell depletion

AIDS is caused by the depletion of CD4+ T cells in lymphoid sites throughout the

body. The GALT is the main site of CD4+ T cell depletion due to the high numbers of

CCR5+ activated memory T cells present [30, 32, 33, 160]. New insights into the

mechanism behind the death of CD4+ T cells have been gained recently from studies by

Warner Greene’s group. They demonstrated two important findings: 1.) Approximately

95% of cells killed by HIV infection are cells that were abortively infected not

productively infected; 2.) The abortively infected cells are killed by pyroptosis, an

inflammatory form of programmed cell death mediated by caspase 1 [163, 164]. These

insights were gained by treating human lymphoid aggregate cultures with antiretroviral

drugs that targeted earlier steps of the HIV life cycle such as binding and entry. When

these drugs were used, there was no CD4+ T cell death. However, when antiretroviral drugs that targeted late stages of the lifecycle were used there was cell death, indicating that the cells were not productively infected when cell death was triggered [163].

The same group would publish findings a year later that show that IFI16, a recently discovered cytoplasmic sensor of double-stranded DNA [165] was responsible for sensing reverse transcription products from HIV-1 resulting in triggering of the inflammasome and initiation of pyroptosis [166]. This finding agrees with a study published by Jakobsen and colleagues the year before that demonstrated that IFI16 knockdown enhanced HIV-1 replication [167].

Warner Greene’s group would go on to publish two more important papers that demonstrate that blood-derived T cells are resistant to HIV-1-induced pyroptosis [168].

This study showed that if you take blood-derived CD4+ T cells and co-cultured them with lymphoid cells they became sensitized to pyroptosis. Upon re-isolation from lymphoid

26

co-cultures, the CD4+ T cells would return to being resistant to pyroptosis [168]. This

phenomenon was due to levels of IFI16 decreasing in the absence of co-culture with

lymphoid cells [168].

This same group would also report in the same year that caspase-1 mediated

pyroptosis by HIV-1 required cell-to-cell spread. Large quantities of free virus were not

sufficient to induce pyroptosis. However, if they co-cultured productively infected T cells

with non-permissive resting T cells, then pyroptosis would be induced [169]. These

findings are significant in that they show that the source of the cells and the context in

which the infection occurs can dramatically affect the results. Most in vitro studies are

done with blood-derived T cells, so it is no surprise why this mechanism of killing was

not discovered sooner.

Contribution of infected macrophages

While CD4+ T cells are susceptible to the cytopathic effects of HIV-1, macrophages are not, even though they are infected [170–173]. Macrophages express

the CCR5 coreceptor and CD4 allowing for HIV-1 binding and entry. However, since

macrophages express less CD4 than T cells [174], macrophage tropism requires

increased affinity of viral Env for CD4 and changes to the CCR5-binding portion of Env

[175–178]. Although macrophages can be infected early by founder viruses, studies

have shown that founder viruses are weakly macrophage-tropic [179] and that founder

viruses preferentially infect CD4+ T cells [180, 181]. However, infected macrophages

can be found in tissues in all stages of disease even in patients on ART [182–184].

While macrophages have been known to be infected by HIV-1 for some time now, it was only recently discovered that not all tissue macrophages are derived from blood-borne monocytes, but that there are self-renewing macrophages that reside in the

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tissues [185–187]. This discovery has profound implications for HIV-1 infection. If tissue macrophages are infected and self-renewing then macrophages may play a larger role as a reservoir for HIV-1 than previously thought. If all macrophages were derived from circulating monocytes then after the tissue-resident cells died all the infected macrophages would be purged as long as the patient was on ART. This finding reconciles data showing that during all stages of infection even on ART that infected macrophages were detectable. Baxter and colleagues have also shown that macrophages can become infected after phagocytosing infected T cells, which may also contribute to the pool of infected macrophages [188].

HIV-1 infection in macrophages primes macrophages and makes them more sensitive to stimulation by TLR ligands in a way that mimics the effects of the inflammatory cytokine interferon-gamma (IFN-γ) [189]. HIV-1 primed macrophages that are exposed to LPS have a similar, but distinct activation profile compared to so-called

M1 macrophages (also known as classically activated macrophages) [190]. HIV-1 primes macrophages through STAT signaling, which is beneficial to the virus, since it requires STAT signaling for its replication [191]. Although these studies were done in vitro they have significance for in vivo infection.

In vivo, HIV-1 infection is associated with major alterations in both the structure and function of the intestine [192–196]. HIV-1-induced immune activation of monocytes/macrophages and the resulting intestinal immune dysfunction are associated with these changes and a breakdown of the intestinal epithelial barrier. The massive T cell depletion in the GALT seen early in infection damages the GALT and

HIV-1 exposure to epithelial tissue has been shown to compromise barrier integrity,

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which both result in microbial translocation – leakage of gut microbial products into the portal venous circulation – that is associated with chronic inflammation in HIV-1 infected patients [197, 198]. This microbial translocation results in chronic activation of macrophages as evident by elevated levels of LPS and soluble CD14 levels, which are independent of viral replication [199].

HIV-1 infected patients have signs of chronic immune activation independent of

ART treatment, including elevated IL-6, hsCRP, amyloid A, amyloid P, D-dimer, prothrombin fragment 1+2 and cystatin C levels [200, 201]. These elevated IL-6 levels, which are likely produced by monocytes/macrophages, and elevated D-dimer levels were associated with all-cause mortality in ART treated patients [201]. Soluble CD14 levels are also elevated in HIV-1-infected patients and are predictive of mortality [202].

A study of aging patients with HIV-1 were assessed for these inflammatory biomarkers

and had elevated levels of soluble CD14, CRP, and IL-6 as well as an increase in

CD57+ cells, which are indicative of immunosenescence seen in aging. However, the

increased immune activation was not associated with physical impairments often seen

in aging individuals [203].

Monocytes and macrophages also play a big role in the pathogenesis of

neurocognitive dysfunctions collectively termed HIV-Associated Neurocognitive

Disorders (HAND). Early in the AIDS epidemic, autopsies of 70 AIDS patients revealed

progressive dementia that was associated with motor and behavioral dysfunction,

impaired memory, and concentration with about 20% of these patients exhibiting severe

dementia [204]. In the mild cases of dementia, patients had lymphocytic infiltrates in the

perivascular space and brown-pigmented macrophages in the brain, whereas in

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advanced cases, clusters of foamy macrophages and multinucleated cells were detected [205]. Another study looking at AIDS patients with neurocognitive dysfunction

found that cells resembling monocytes or macrophages were staining positive for HIV-1

antigen, suggesting that the neurological symptoms detected in these AIDS patients

were due to the virus in the brain [206]. In later years with more sophisticated

technology, laser microdissection techniques were used to establish that brain

macrophages as well as astrocytes and neurons were infected with integrated HIV-1,

which adds an additional challenge in purging the viral reservoir as neurons are

terminally differentiated, long-lived cells [207–209].

The mechanism by which HIV-1 infected macrophages in the brain result in CNS

damage is unknown. However, it is thought to be the result of one or both of two

different mechanisms. HIV-1 proteins gp120, Tat, Vpu, and Vpr have all been shown to

have direct neurotoxic effects on neurons in vitro [210]. A second possible mechanism

is secondary effects of factors secreted from infected macrophages, such as TNF,

platelet activating factor, arachidonic acid, quinolinic acid, and other metabolites [210].

In summary, macrophages and monocytes clearly play an important role in the

pathogenesis of AIDS and or AIDS-related comorbidities, such as HAND. For a

sterilizing cure to be achieved, elimination of infected macrophages and monocytes will

be just as important as the elimination of infected T cells. With microglial/macrophages

reservoirs in the brain where antiretroviral drugs may not have access to will present a

unique challenge in achieving a cure.

Viral latency

Despite a lack of detectable plasma viremia in patients on long-term ART [37, 38,

40], replication-competent HIV-1 can still be found in CD4+ T cells from these individuals

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[42–44]. If ART is halted plasma viremia rebound is detectable within weeks [41]. These findings pointed out the existence of HIV-1 reservoirs. Later studies would estimate the half-life of the reservoirs to be longer than the lifetime of the patients, meaning that the patients would be infected for life despite ART [211–213]. The reservoir is primarily made up of latently infected memory T cells, which can persist long-term due to homeostatic proliferation of the memory T cell pool [214, 215].

The consequences of long-lived memory T cells harboring virus is that if these long-lived cells, which are maintained for the life of the host, are ever reactivated upon

antigen recall, then the virus can begin to replicate again and progression to AIDS will

resume if ART has been halted. ART can act as a functional cure that prevents the

expansion of the viral reservoir. In the context of ART, latently infected cells can be

activated and begin producing virus again, but cannot infect new cells. The former

statement is backed up with evidence that showed that intensification of ART had no

effect on the size of the latent reservoir nor did it affect any residual viremia detected by

ultra-sensitive assays [216–220]. Because of this, several strategies for forcing

reactivation of the latent reservoir have been proposed.

One strategy known as the “shock and kill” strategy is where latency reactivated

agents (LRAs) are used to treat the patient to reactivate the latently infected cells while

the patient is also taking ART to prevent new infections [221, 222]. This forces the

latently infected cells to reactivate and most of these cells will be killed by the cytopathic

effects of the virus, but the ART treatment will prevent the virus from spreading and

establishing a new reservoir. The first clinical trial to attempt this used Vorinostat

(SAHA), a histone deacetylase (HDAC) inhibitor, and found that there was boosted

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reactivation in the patients [223]. Attempts to do this have shown, however, that not all the latently infected cells are reactivated upon treatment with LRAs [224].

Mechanisms of T cell latency

Epigenetic silencing. Regulation of gene expression in eukaryotic cells has many levels, including at the level of DNA accessibility. DNA wrapped around histone proteins allows for compacting the DNA as well as preventing access to the DNA in these regions by transcriptional machinery [225]. These highly condensed regions of transcriptionally silent DNA are referred to as heterochromatin [226]. Although HIV-1 proviral DNA preferentially integrates into areas of open chromatin [68, 227], T cells returning to a resting state may undergo chromatin remodeling that renders the integrated proviral DNA in heterochromatin and inaccessible to transcriptional machinery.

Studies have shown that many proviruses that are integrated into sites of heterochromatin in T cells are inducible by different stimuli [75]. Treatment of T cells

with HDAC inhibitor SAHA showed promise in an early proof-of-concept study [228].

Other studies in cell lines and primary CD4+ T cells showed some efficacy with HDAC inhibitors [229–231]. However, follow-up studies showed that the long-term efficacy of this approach was lacking [232–234]. This is likely due to the multifactorial nature of

HIV-1 latency. While in some cells chromatin accessibility may be the primary cause of latency, this approach alone will not reactivate the entire latent reservoir.

A 2003 study by Pion and colleagues claimed that methylation of CpG sites in the HIV-1 LTR promoter was not predictive of transcriptional activity. However, two studies in 2009 demonstrated that the HIV-1 LTR promoter can also be epigenetically silenced due to the presence of CpG islands that are methylated in the J-Lat cell line (a

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Jurkat cell model of latency) and primary CD4+ T cells [235, 236]. One of these studies

also showed that the methylation inhibitor 5-aza-2’-deoxycytidine had a synergistic

effect with NFκB inducers TNF and prostratin [236]. It is not entirely clear how much

methylation of CpG islands in the HIV-1 LTR promoter contributes to epigenetic

silencing of HIV-1 transcription, but as with chromatin remodeling, it is likely a

combination of factors depending on the cell and the genomic location of the proviral

integration.

Transcriptional interference. Another potential mechanism of latency is

transcriptional interference. This can occur due to convergent promoters, which is what

occurs when the LTR is downstream of an active host gene. Elongating RNA

polymerase II reads through the LTR pushing the transcription factors out of its way as it

transcribes the host gene [237, 238]. This can also happen if the host gene and LTR

promoter are facing in opposite direction if the RNA polymerase II collisions that could

result in premature termination of transcription [239].

Sequestration of transcription factors. Latency can also occur as a result of

sequestration of host transcription factors required for LTR promoter activity. HIV-1 has

a unique niche in that the transcription factors needed for the LTR promoter are the

same transcription factors that would be present during activation of T cells, such as

NFAT and NFκB [240–243]. NFAT and NFκB bind to the same site on the HIV-1 LTR

promoter called the κB sites, which are important for HIV-1 transcription [244, 245].

These transcription factors are required for immune activation and elimination of viruses

but are hijacked by HIV-1 LTR promoter. In a resting state, T cells sequester these

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transcription factors in the cytoplasm. This may be a major mechanism for regulating latency in CD4+ T cells.

Latency reactivating agents

Latency reversing agents (LRAs) are drugs that induce expression of latent HIV-

1. This strategy relies upon reactivation of latent virus to start producing viral proteins that will either kill the infected cell due to the cytopathic effects of the virus or will allow for enough expression of viral proteins for immune surveillance by CD8+ cytotoxic T

lymphocytes (CTLs) to kill infected cells. This strategy also requires simultaneously

treating patients with LRAs and ART to prevent the infection of new cells. An ideal LRA

will effectively reduce the size of the viral reservoir without too many adverse side

effects.

A majority of the latent reservoir is made up of memory T cells and therefore

effective LRAs will be drugs that target these cells [246]. There have been two main

approaches in developing LRAs: agents that inhibit host factors that actively silence

viral gene expression and agents that activate host factors that enforce viral gene

expression. There has been some limited success with LRAs that reverse epigenetic

silencing of proviral DNA such as HDAC inhibitors and histone methyltransferase

inhibitors that seem to work synergistically together at least in cell line models [235,

236, 247].

The basis for many other LRAs including the first ones to be used in the “shock

and kill” approach involve drugs that induce the activation of NFκB or NFAT since those

factors are required for HIV-1 transcription [240, 244]. The HIV-1 protein, Tat, recruits

the transcriptional elongation factor P-TEFb, which is required for transcription from the

LTR promoter [85, 248]. Targeting the protein kinase C (PKC) pathway that results in

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activation of NFκB and NFAT also influences the activity and availability of P-TEFb.

PKC agonists may promote upregulation of the levels of CycT1, which promotes P-

TEFb activity or release of P-TEFb from HEXIM1, which restricts P-TEFb activity [249,

250].

The PKC agonist bryostatin has shown a significant effect on reactivating HIV-1 in ex vivo models, however, in a phase I clinical trial it failed to have any effect on the transcription of latent HIV-1 [251]. Another study with bryostatin did show that its effects synergized with HDAC inhibitors in the J-Lat cell model [252]. However, the efficacy of this approach has yet to be determined in vivo. Other PKC agonists such as phorbol 12- myristate 13-acetate (PMA) and prostratin work well in vitro [253]. However, PMA is not

safe for use in vivo [254], presenting the challenge of finding PKC agonists that are

effective but still safe. Although the shock and kill strategy has shown some efficacy it

has not significantly reduced the viral reservoir in HIV-1-infected patients in clinical

trials. There is a necessity for new treatments that can complement or boost the efficacy

of the existing treatments.

The macrophage reservoir

While the “shock and kill” strategy holds promise for eliminating the T cell

reservoir, the implications on the macrophage reservoir are less clear. It is still a matter

of debate as to whether macrophages should be considered part of the viral reservoir or

not [255]. The challenge in defining the viral reservoir is that in humans the tissues

where these viral sanctuaries may exist are not accessible. While autopsies have

confirmed the existence of HIV-1 in macrophages in the brain it is unclear if this so-

called reservoir can reseed systemic infection if a successful eradication of the T cell

reservoir was achieved [255].

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Another point of controversy is whether infected macrophages are truly latent or whether there is chronic low-level replication occurring during suppressive ART [256]. If

macrophages were persistently producing new virions and infecting new macrophages it

would be apparent through analysis of viral evolution. However, studies have shown

that patients on suppressive ART do not exhibit signs of ongoing viral evolution [257,

258]. The lack of viral evolution implies that occasional low-level viremia seen in ART

suppressed patients is due to virions being produced from cells that were already

infected and not expansion of the reservoir. Given that macrophages are not typically

long-lived cells beyond a few weeks, this would suggest that macrophages do not

represent a substantial part of the reservoir that is a barrier to a cure.

However, these conclusions became contested again with new evidence from

recent discoveries. First, it was recently discovered that many tissue macrophages are

capable of self-renewal that is not dependent on the influx of blood monocytes [185–

187]. Second, a study using a myeloid-only humanized mouse model of HIV-1 infection,

where there are only macrophages but not CD4+ T cells, showed that persistent HIV-1

replication maintained the tissue reservoir despite therapy [259, 260]. Third, a study

showed persistent HIV-1 replication occurs in the tissue reservoir during therapy [261].

The implication from this study is that even during ART there may be low-level

persistent replication in lymph nodes where ART cannot gain access even though this

amount of replication may not result in detectable viremia [261]. The caveat to this study

is that there were only three patients enrolled, however, the implications of the evidence

presented are still important to consider.

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To eliminate the macrophage reservoir, a different approach or a complementary approach to “shock and kill” may be necessary. One possibility is to design drugs that can make the macrophages more susceptible to the cytopathic effects of the virus.

Another approach is to design drugs that make macrophages better at eliciting an

adaptive immune response from cytotoxic T lymphocytes.

Immune Control of HIV-1

Interferons

Discovery of interferons

Interferons (IFNs) are a class of soluble cytokines, first reported in 1957, that

were named for their ability to “interfere” with viral replication [262, 263]. Later in the

1980’s it would be realized that there was a family of type I interferon genes that would

be divided into α (alpha), β (beta), ω (omega), and κ (kappa) subfamilies [264–268].

Interestingly, these genes were found to not be spliced and were all located on the

same chromosome in humans (chromosome 9) [269]. A second class of IFN, type II

IFN, was first described in the early 1970s’ [270] and assigned to the type II family due

to its different chemical properties and the later finding that it utilizes a different receptor

[271, 272]. Interferons have now been classified into three families: Type I, Type II, and

Type III. Interferons play an important role in cell-autonomous immunity as well as in

tuning the adaptive immune response.

Type I IFN and JAK/STAT signaling

In humans, the type I IFN family consists of thirteen IFN α, one IFN β, one IFN-ε,

one IFN-κ, and one IFN ω subtypes. While these IFNs use the same receptor, they bind

with different affinities and have slightly different quantitative outcomes [273–275]. Type

I IFNs can be produced by most cell types in the body and are produced in response to

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viral ligands that signal through pattern recognition receptors. Plasmacytoid dendritic cells, however, are the primary producers of IFN-α [276]. IFNs are secreted from the cells and can act upon the heterodimeric IFN α receptor (IFNAR), which is composed of the IFN-αR1 and IFN-αR2 subunits [277–282].

The Janus Kinases (JAKs) JAK1, JAK2, and tyrosine kinase 2 (TYK2) [283–285]

are associated with the cytoplasmic tails of receptors [286–288]. After ligation of IFNAR

by type I IFN binding, JAK1 and TYK2, which are already associated with the

cytoplasmic tails of IFNAR, are activated [289–293]. Once activated, JAK1 and TYK2

phosphorylate and activate STAT1 and STAT2 [294–297], which complex with IRF9 to

form the ISGF3 complex [298–300]. This complex translocates to the nucleus [289,

301–306] and can bind to the ISRE in the promoter of hundreds of IFN-stimulated

genes (ISGs) [307–309].

ISGs are a collection of genes that respond to IFN stimulation. The protein

products of these genes have many diverse roles that help in the cell's defense against

viral and bacterial pathogens [288, 310, 311]. The gene products of ISGs can alter the

way antigens are processed and presented to the adaptive immune system, the way

mRNAs are processed and translated, receptor expression for cytokines and

chemokines, and secretion of cytokines and chemokines that recruit additional immune

cells to the site of infection. These cellular changes affect the availability of cellular

components needed for viral replication and can restrict the release of viruses that

utilize budding or multi-vesicular bodies for escaping from the host cell as well as help

to establish an antiviral state in nearby uninfected cells by making the cells harder to

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infect in the first place [288, 310, 311]. Additionally, the IFN response can help to recruit and prime adaptive immune cells [312].

HIV-1 restriction factors and evasion by HIV-1

It has been well established that type I IFNs can restrict HIV-1 replication in macrophages and T cells in vitro [313–318]. To date there have been many IFN- inducible HIV-1 restriction factors identified. HIV-1 restriction factors are host factors that restrict viral replication. Many of these factors, however, have little or no effect on wild-type virus due to evasion mechanisms employed by HIV-1 [319]. Apolipoprotein B mRNA editing enzyme, catalytic polypeptide-like 3 (APOBEC3G), Tripartite motif- containing protein 5 (TRIM5α), Sam domain and HD domain-containing protein 1

(SAMHD1), and bone marrow stromal antigen 2 (BST2/tetherin) are well-established

HIV-1 restriction factors [319–321].

APOBEC3G. APOBEC3G was first identified as a restriction factor of HIV-1 in an experiment utilizing a vif-deficient HIV-1 strain [322]. APOBEC3G is packaged into viral particles due to its ability to associate with the viral RNA [323]. When the virion infects a new cell, APOBEC3G associates with the reverse transcription complex and deaminates cytidine residues of the negative strand causing guanosine to adenosine mutations to result in the positive strand when it is copied from the deaminated negative strand [324–327]. APOBEC3G is only effective against vif-deficient strains of HIV-1 because Vif facilitates the ubiquitination of APBEC3G and its subsequent degradation by the proteasome [328–331].

TRIM5α. The restriction factor Trim5α was discovered in a screen of rhesus macaque genes that could restrict HIV-1 in human cells [332]. TRIM5 was found to restrict HIV-1 at a post-entry step that prevented it from synthesizing viral cDNA [332].

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While the rhesus macaque variant of this gene can restrict HIV-1, the human homolog cannot restrict wild-type HIV-1 strains [332]. TRIM5α, while not very effective at blocking viruses in their natural hosts, is very effective at blocking viruses from other species

[333]. TRIM5α can be made to recognize HIV-1 with a small number of amino acid changes in the V1 segment of TRIM5α [334–337]. Additional studies have shown that the PRYSPRY domain of TRIM5α determines its ability to bind to the capsids of particular viruses [338]. It is not entirely clear what the mechanism of viral restriction by

TRIM5α is, but it seems to be essential that TRIM5α binds to the capsid of incoming

viruses and may promote capsid fragmentation that inhibits formation of the reverse

transcription complex [334, 338].

SAMHD1. SAMHD1 was identified as a restriction factor that is expressed in

macrophages and dendritic cells that prevents robust HIV-1 replication [339, 340].

SAMHD1 is a deoxyribonucleoside triphosphate triphosphohydrolase (dNTPase) that

degrades and depletes the pool of dNTPs that are available for reverse transcription of

HIV-1 [341, 342]. SAMHD1 is constitutively expressed in many different cells types, but

its antiviral activity is activated by phosphorylation of SAMHD1 on Tyr592 [343, 344].

Phosphorylation of SAMHD1 is dependent on cyclin-dependent kinase 1 (CDK1), the

activity of which depends on the cell cycle [344–346]. While SAMHD1 can effectively

restrict some HIV-1 replication, it is counteracted by the accessory protein Vpx [339,

340]. However, Vpx is only found in HIV-2 and SIV. Therefore, HIV-2 and SIV can

replicate efficiently in SAMHD1 expressing cells, whereas HIV-1 cannot [347].

BST2/Tetherin. The restriction factor tetherin was discovered based on the finding that the accessory protein Vpu was required for virion release from cells [348].

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Tetherin is inducible by IFN and acts by “tethering” virions to the surface of the cell and preventing their release [349–352]. Since tetherin prevents the release of virions, the virions will accumulate in endosomes [349]. Tetherin has a transmembrane anchor at its amino terminal end with a glycophosphatidylinositol lipid anchor at its carboxyl terminus

[353]. Both anchoring domains are essential for the antiviral activity of tetherin. Mutants that lack either of these anchors fail to tether HIV-1 virions to the cell and will actually end up incorporated into the free virions instead [354]. HIV-1 evades this host mechanism of restriction by activity of the accessory protein Vpu [351, 352].

Other IFN-induced effectors. Although the above factors are the most well-

established HIV-1 restriction factors, there are many other IFN-inducible effectors that have been shown to have anti-HIV-1 activity. Schoggins and colleagues performed a high throughput screen of IFN-inducible factors in a T cell line to identify factors that could restrict HIV-1 replication [355]. This study identified TNFRSF10A, IFITM3, IRF1,

CD74, IFITM2, and MX2 as effectors that can potently restrict HIV-1 in the MT-4 T cell line [355]. Subsequent studies have confirmed the anti-HIV-1 activity of IFITM family proteins [356–360]. Other factors such as SLFN11 [361], MX2 [362–364], HERC5 [365], and ISG15 [366–368] have also been shown to have anti-HIV-1 activity. These factors are probably more appropriately called HIV-resistance factors rather than true restriction factors because HIV-1 doesn’t actively subvert these mechanisms and each of these factors cannot by itself completely restrict replication [319]. However, robust expression of many of these factors simultaneously would probably have a very strong effect on restricting HIV-1 replication.

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B cell response

It has been known since almost the discovery of HIV-1 that infected individuals could produce neutralizing antibodies to HIV-1 [369, 370]. Immune complexes form as early as 8 days after the virus becomes detectable in the blood with free plasma anti-

HIV-1 antibody against gp41 subunit of HIV-1 Env as early as 13 days after the virus becomes detectable [371]. This same study showed that anti-gp120 antibodies did not appear until 13 days after virus becomes detectable in the blood [371]. However, these antibody responses were not sufficient to control the virus. Furthermore, these antibody responses weren’t sufficient to drive evolution of the viral envelope indicating that they were not putting any selective pressure on the virus [54].

Neutralizing antibodies do develop against the founder strain months after infection but are unable to neutralize mixed viral populations derived from other infected individuals [372–380]. Furthermore, these neutralizing antibodies were the drivers of viral evolution resulting in new viral populations that were resistant to neutralization.

These viral escape mutations resulted in a changing glycan shield which prevented the binding of neutralizing antibodies to Env [376, 377, 381–384]. One of these studies also looked at antibodies from long-term non-progressors (HIV-infected individuals who do not progress to AIDS despite persistent viremia) and found that these antibodies did have the ability to broadly neutralize heterologous virus isolates [375].

While much work has been done to elucidate the role of B-cells and antibodies in

HIV-1 infection, it is still not completely understood whether neutralizing antibodies contribute to controlling viremia or not. It will be important to continue looking for an answer for aid in designing an effective preventative vaccine. If a broadly neutralizing

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antibody that worked on a wide range of heterologous virus could be developed it could lead to potentially beneficial therapies.

T cell response

HIV-1 primarily infects and depletes CD4+ T cells, however, CD8+ T cells are not

infected. To create an effective preventative vaccine, a better understanding of what

factors are required for a robust CTL response against HIV-1 is required. In HIV-1-

infected patients, strong CD8+ T cell responses have been reported [385–387]. These

CD8+ T cells were shown to be able to inhibit HIV-1 replication in vitro [388]. Although

this population of CD8+ T cells is sufficient to restrict HIV-1 in vitro, the patients still progress to AIDS indicating that it is not sufficient in vivo. Furthermore, this study showed that CTLs were not killing infected cells per se, which suggests that there is some other mechanism by which they are restricting HIV-1 replication in vitro like IFN-γ secretion [388]. Later studies confirmed that IFN-γ was produced by HIV-1-specific

CD8+ T cells [389, 390].

The importance of CTL activity in HIV-1 infection is evident by the correlation between disease outcome and the patient’s HLA class I allele [391–393]. One study showed that individuals with the HLA B*5701 allele that were infected with HIV-1 were long-term non-progressors, meaning despite infection levels being detectable, CD4+ T

cell levels didn’t drop [392]. Depletion of CD8+ T cells in an SIV infection model in

macaques have also confirmed the importance of a CTL response in controlling viremia

[394, 395]. While there is some evidence that suggests that lysis of infected cells is

important during acute infection, it is not completely clear what respective contributions

direct killing of infected cells or production of cytokines have in control of HIV-1 [396,

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397]. A better understanding of what CD8+ T cell effector functions are needed for HIV-1

control will be required for effective vaccine design.

During acute HIV-1 infection, the CTL response is usually very narrowly directed

with few epitopes targeted as the viral set-point is established [389, 398, 399]. However,

in chronic infection, there is a much broader CTL response associated [400]. This is due

in part to viral evolution as a consequence of immune selection. In chronically infected

patients, as high as 19% of total CTLs are HIV-1 specific with more than a dozen CTL epitopes targeted [401, 402]. Despite such a broad and robust CTL response, most infected persons are not able to control viremia. This may be in part due to the lack of a

CD4+ T cell response. A study performed in 2002 showed that HIV-1 specific CD4+

memory T cells are preferentially infected over other memory CD4+ T cells [158]. Upon

their reactivation, HIV-1-specific CD4+ T cells would be susceptible to the cytopathic

effects of HIV-1. This may partially explain the lack of a robust CD4+ T cell response.

There are some infected persons known as elite controllers who are able to

control viral loads without the use of ART. The prevalence of elite controllers is

estimated to make up less than 1% of the infected population [403–407]. However,

these individuals have been shown to have intact CD4+ T cell responses [408], which

may be the reason they are able to control viremia. What allows these individuals to

control viremia during acute infection early on to prevent depletion of CD4+ T cells is

unknown, although there is an association with certain HLA class I alleles [403].

Summary and Hypothesis

Current therapies for HIV-1 are able to successfully treat infection and can suppress viremia to undetectable levels. However, this treatment is required for life and is not readily available to all people with HIV. The burdens associated with a chronic

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illness and taking medication for life are difficult psychologically and economically. Non- adherence to prescribed medication results in many individuals not having adequately suppressed viremia despite access to therapy. The best hope for overcoming these problems is achieving a sterilizing or functional cure. To achieve a cure, the viral reservoirs must be eliminated. Current therapies aimed at doing so have had limited efficacy. The hypothesis of the current study is that targeting host factors that are required for HIV-1 replication such as DYRK1A and USP18 can augment current therapeutic approaches that utilize shock and kill or treatment with PEGylated IFNs.

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CHAPTER 2 DEPLETION OF USP18 ENHANCES TYPE I IFN RESPONSIVENESS AND RESTRICTS HIV-1 INFECTION IN MACROPHAGES

Introduction

It is well established that type I interferons (T1 IFNs) can restrict acute HIV-1 infection in vitro [313, 316, 317, 409, 410]. In clinical trials treating human patients with recombinant IFN-α2a, T1 IFNs can suppress viral replication in the absence of ART in some patients [411, 412]. However, this approach failed in long-term treatment when study subjects became refractory to IFN treatment and viral loads returned to previous levels. T1 IFNs are important in establishing early control of HIV-1 in an in vivo SIV rhesus macaque model [413], but chronic T1 IFN signaling correlates with chronic immune activation, immune exhaustion, elevated interferon-stimulated gene (ISG) expression, and poor long-term control of HIV-1 [414, 415].

Two recently published studies have shown that in a humanized mouse model of

HIV-1 infection that experimental blockade of T1 IFN signaling resulted in restored immune function and rescued T cell function, including HIV-1-specific T cells [416, 417].

In chronic LCMV infection, blockade of interferon α receptor (IFNAR) also had significant benefit [418, 419]. These studies highlight the importance of IFN regulation in the context of HIV-1 infection specifically and viral infections in general. However, little is known about how HIV-1 induces IFNs and why IFNs are unable to control infection in vivo.

Signaling through IFNAR results in the phosphorylation and activation of STAT proteins including STAT1 and STAT2. The consequence of STAT1/2 phosphorylation is induced expression of hundreds of ISGs including components of the ISGylation system, which have profound effects in tuning the IFN response [307–309, 420]. The

46

protein products of ISGs then target host and viral machinery as a means to restrict viral replication. However, a small subset of the ISGs expressed are negative feedback mechanisms that turn off IFN signaling so that resolution of the immune response can occur. One of these important negative regulators is ubiquitin specific proteinase 18

(USP18).

USP18

The cDNA sequence was first cloned in 1999 by Dong-Er Zhang’s group in

AML1-ETO-expressing mice [421]. The same group cloned and characterized the human homolog in 2000 [422]. It was found to have sequence homology to other ubiquitin-specific proteases and was given the name Ubp43 for ubiquitin-specific proteinase that is 43 kDa. It was later renamed USP18 when the names of the proteins in that family were standardized.

USP18 was first identified as a deISGylating enzyme in 2002 when it was observed that USP18 failed to deubiquitinate proteins in vitro [423]. The authors hypothesized that perhaps it was involved in removing one of the ubiquitin-like proteins,

such as SUMO, Nedd8, or ISG15. They confirmed in Usp18-/- mice that there was an

increase in ISGylated proteins, making USP18 the first ISG15-specific protease to be

reported. Later studies by Basters and colleagues would confirm through structural

biology that USP18 was indeed the deconjugating enzyme for ISG15 [424, 425].

It was not until 2006, however, that the role of USP18 as a negative regulator of

JAK/STAT signaling was discovered [426]. Dong-Er Zhang’s group reported that USP18 could negatively regulate JAK/STAT signaling by blocking JAK1 binding to the cytoplasmic portion of the IFNAR2 subunit. This was shown to be independent of its

47

enzymatic activity. A role for STAT2 as an adaptor for USP18-mediated suppression of type I IFN signaling would later be demonstrated [427].

Studies in Usp18-/- knockout mice showed a profound impact on the ability to

combat viruses as well as some bacteria [428, 429]. It was initially believed to be due to

increased ISGylation, however, with the discovery of USP18’s non-enzymatic role in

IFNAR signaling, it made sense why its absence would have such a profound effect on

the antiviral response. In addition, another research group showed that the phenotype

of Usp18-/- mice was not rescued with Isg15-/-[430].

Microarray analysis published in 2007 would reveal that in the absence of

USP18, murine macrophages would respond much more potently to IFN-β [431]. These knockout macrophages had an increase in expression of genes related to chemokines and cytokines, chemokine and cytokine receptors, antigen processing and presentation, and genes involved in immunity against viruses. This work begged the question, does

USP18 play the same important role in humans and what would be the consequences of USP18 deficiency in human?

In 2016, a study was published by Meuwissen and colleagues that identified loss-

of-function mutations in the USP18 gene in five patients with pseudo-TORCH syndrome

from two unrelated families [432]. Pseudo-TORCH syndrome is a condition that

resembles TORCH syndrome in the absence of any congenital infections. TORCH is an

acronym that stands for toxoplasmosis, other agents, rubella, cytomegalovirus, and

herpes simplex. The symptoms of this condition, referred to as an interferonopathy, are

the result of a congenital infection that affects fetal development due to the sustained

inflammation as a result of the infection. Pseudo-TORCH syndrome presents as

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TORCH syndrome, but in the absence of any congenital infection. In this study, it was found that five of these patients had pseudo-TORCH syndrome due to loss-of-function mutations in USP18. Without USP18 to regulate the type I IFN pathway, sustained and prolonged inflammation lead to birth defects, microcephaly, and calcium deposits in the brain [432]. Although USP18 function is necessary for proper development, selective, transient targeting of USP18 with pharmaceutical drugs could potentially be used as an immune adjuvant to treat viral infections without having profound negative effects on the patient. However, this needs to be evaluated in human cells in vitro before it is attempted in any human clinical trials.

The role of human USP18 has also been extensively studies of hepatitis C virus

(HCV) infection. In 2005, Chen and colleagues compared the gene expression profiles from liver biopsies from patients that were responders or non-responders to PEGylated

IFN-α treatment [433]. They found that along with upregulation of classical IFN-

stimulated genes like MX1, OAS3, and OAS2 that USP18 and ISG15 were upregulated

in non-responders compared to responders [433]. Given the role of USP18 and ISG15 in regulating type I IFN signaling, they hypothesized that USP18 expression may be rendering the non-responders refractory to IFN treatment.

Following up on this study, the same group used siRNA to knockdown USP18 in the hepatic cell line Huh-7.5. They found that with USP18 knockdown HCV replication was significantly reduced ISGylation of cellular proteins was increased [434]. Although they hypothesized that this defect in viral replication was due to increased ISGylation, they found that increasing an enzymatic activity deficient mutant had no effect on viral replication and that forcing increased ISGylation actually enhanced viral replication

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[435, 436]. This evidence suggested that the increased antiviral activity in the knockdown cells was due to USP18’s non-enzymatic role in regulating JAK/STAT signaling. Other studies of human USP18 in cell lines has shown that USP18 expression plays a role in limiting TRAIL-induced apoptosis and has also been shown to regulate the susceptibility of certain cancer cells to IFN-α and drug-induced apoptosis

[437, 438].

ISG15 and ISGylation

ISG15 is an interferon-stimulated gene which codes for a protein that after processing is approximately 15 kDa and has high homology to ubiquitin [439, 440].

ISG15 is a ubiquitin-like post-translational modification that gets conjugated to target proteins [441] through a process similar, but slightly different from ubiquitination [442]

that uses an E1, E2, and E3 enzyme. ISG15 has two different ubiquitin-like domains

that are connected by a hinge region.

First, ISG15 is activated by an E1-like protein, Ube1L [443], that forms a thioester

bond with the C-terminal glycine residue of ISG15. This complex is recognized and

transferred to an E2-like enzyme. There have been two E2-like enzymes identified that

can function in ISGylation, UbcH6 and UbcH8 [444, 445], although UbcH8 is the

predominant one [446]. UbcH8 and UbcH6 are also involved in the ubiquitin pathway,

so although ISGylation has a unique E1-like enzyme it shares E2 enzymes with the

ubiquitin pathway [444, 445]. The ubiquitin E3-ligases HERC5 and TRIM25 have been

identified as ISG15 E3-ligases that catalyze the covalent binding of ISG15 to lysine

residues on its target proteins [447, 448].

USP18 is an ISG15-specific deubiquitinating enzyme that removes ISG15

through thioesterase activity releasing free ISG15 back into the cell [423–425]. Other

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USP protease family members, USP2, USP5, USP13, and USP14, were identified in a screen by Catic and colleagues as able to remove ISG15 as well as ubiquitin but are not specific to ISG15 like USP18 [449]. It is important to note that ISG15, USP18, and the

E1, E2, and E3 enzymes that are involved in conjugation are all interferon-inducible proteins. Many protein products have been identified as targets of ISGylation although the specific effects of ISGylation have not been completely elucidated for all targets of

ISGylation. Many of the targets of ISGylation are interferon-inducible genes, but many constitutively expressed genes are also targeted [450].

In some cases, ISGylation has been demonstrated to stabilize expression of target proteins. ISGylation has been shown to prevent IRF3 from being ubiquitinated and targeted for proteasomal degradation, thus extending its half-life [451, 452]. This serves as positive regulation of IFN signaling since IRF3 is involved in transcription of type I IFNs. ISGylation of USP18 has also been shown to stabilize its expression by preventing USP18 from being targeted for degradation by ubiquitination, which is important in preventing over-amplification of type I IFN signaling [453, 454].

ISGylation of proteins can also serve as a mechanism of negative regulation. The host protein, PPM1B, which is a phosphatase enzyme that deactivates TAK1 and IκB

kinase is also been shown to be ISGylated. ISGylation of PPM1B results in the inability

to dephosphorylate TAK1 and IκB kinase thus acting to enhance NFκB signaling [455].

The host protein RIG-I, which plays a role in sensing of viral RNA is ISGylated as a

means to negatively regulate RIG-I expression and viral sensing [456].

ISGylation of proteins can also serve to modify the function of proteins. The

protein 4EHP, which competes with eIF4E for binding to 5’ caps of mRNA, is ISGylated,

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enhancing its affinity for 5’ cap binding. In this manner, 4EHP acts as a suppressor of cap-structure dependent translation [457]. This would interfere with translation of viral mRNAs that rely on cap-structure dependent translation, thus inhibiting viral replication.

ISGylation of protein kinase R (PKR) on lysines 69 and 159 activates it and induces the

phosphorylation of eIF2α, which also results in inhibition of translation initiation [458].

ISGylation of 4EHP and PKR can have dramatic effects on the ability of viruses to

replicate since they depend on host translation mechanisms.

ISGylation has also been shown to interfere with viral budding from the plasma membrane by ISGylation of ESCRT complex protein CHMP5 [368] and exosome

secretion by ISGylation of ESCRT complex protein TSG101 [459]. ISGylation also

interferes with ubiquitination by ISGylating other ubiquitin E2/E3 enzymes [460, 461]

and by decreasing the levels of polyubiquitinated proteins that can be degraded by the

proteasome [462, 463]. Thus, ISGylation of target proteins can have pleiotropic effects

on cell processes. ISGylation can stabilize expression of proteins, negatively regulate

proteins, and modify the function of proteins.

It has been shown that the primary targets of ISGylation are newly translated

proteins, which can serve to mark these proteins as “made during a viral infection”

[464]. It has also been shown that viral structural proteins can be ISGylated, such as

HIV-1 Gag protein [365], influenza A virus NS1 protein [465], and influenza B

nucleoprotein [466]. ISGylation of viral structural proteins may serve to disrupt the

formation of repeating capsid structures, thus disrupting the ability of the virus to

assemble [464, 466]. ISG15 can also be secreted from the cells in its unconjugated

form and it has been shown that this can stimulate NK cells to produce IFN-γ. However,

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it is not well established to what degree free ISG15 is required for an effective immune response or by what pathway it is secreted.

The importance of ISG15 is reflected by the fact several viruses have evolved countermeasures for ISGylation. Influenza B’s NS1 protein can bind to free ISG15 and

directly neutralize it [443]. The SARS and MERS viruses encode a viral deISGylating

enzyme that provides a similar function to USP18 [467, 468]. These viral mechanisms

for combating the ISGylation system reflect the importance of ISGylation in the immune

response against viruses.

Rationale and summary

Macrophages play an important role in HIV-1 as reservoirs and can contribute

directly to HIV-1 pathogenesis [469]. HIV-1 in the ART era can be seen as a chronic

disease characterized by chronic immune activation and chronic inflammation with a higher risk of non-AIDS related morbidities and mortalities. Macrophages play an important role in this process and can act as mediators of inflammation [470, 471]. We recently reported that HIV-1 replication in macrophages requires the activity of STAT1,

a protein usually associated with antiviral responses [191]. The role of STAT1 was in

the post-integration expression of HIV-1 mRNA from the LTR promoter. This

paradoxical mechanism where HIV-1 usurps antiviral pathways as a means of driving its own replication suggests that a complex host-pathogen interplay ultimately determines if

HIV-1 can efficiently replicate in macrophages. In general, the role of macrophages in the HIV-1 life cycle is important because eliminating persistently infected macrophages in addition to latently infected T cells will be necessary for a sterilizing cure to be achieved. Thus, it is critical that we understand the host-pathogen dynamics that

53

regulate HIV-1 replication in macrophages and understand how HIV-1 subverts the innate immune response including the potent antiviral T1 IFN response.

Our data show that USP18 expression is induced by HIV-1 in human macrophages. The goal of this study was to determine the role of USP18 in HIV-1 infection/replication and determine if the IFNAR-blocking effect of USP18 benefits HIV-

1. We hypothesized that USP18 suppresses antiviral pathways that would normally

restrict HIV-1 and that perturbing USP18 would restore and enhance these pathways

leading to better control of the virus. To test this, we have utilized several USP18

knockdown cell models and a novel CRISPR/Cas9 knockout induced pluripotent stem

cell (iPSC)-derived macrophage model. Our data show that experimental depletion of

USP18 in human macrophage models does restrict HIV-1 replication due to an

enhanced cellular antiviral response and increased sensitivity to the effects of T1 IFN. In

addition, we confirm previous findings by our group that low-level signaling through T1

IFN signaling pathways is required for HIV-1 replication. Thus, USP18 benefits HIV-1

replication by striking a balance between weak IFN signals required for viral replication

and strong IFN signals that would halt viral replication.

Methods and Materials

Cell culture

Monocyte-derived macrophages (MDMs). Peripheral blood mononuclear cells

(PBMCs) were prepared from leukopaks obtained from healthy donors from LifeSouth

Community Blood Center (Gainesville, FL) under approval by the Institutional Review

Board at the University of Florida. PBMCs were isolated by centrifugation on

LymphoSep® Lymphocyte Separation Medium (MP Biomedicals, Santa Ana, CA) medium. Monocytes were isolated by positive selection on LS Columns (Miltenyi Biotec,

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Bergisch Gladbach, Germany) with human CD14 MicroBeads (Miltenyi Biotec). Freshly isolated monocytes were differentiated into macrophages in MDM medium (DMEM with

4.5 g/L glucose (Corning, Corning, NY), 10% heat-inactivated human serum (Zen-Bio,

Research Triangle Park, NC), 2 mM L-glutamine (Corning), and 100 IU penicillin- streptomycin (Corning)) supplemented with 10 ng/ml M-CSF for 7-10 days. After differentiation, the medium was replenished without M-CSF and the cells rested overnight before being used for experiments.

THP-1 cells. THP-1 cells (TIB-202; ATCC, Manassas, VA) were cultured in THP-

1 medium containing RPMI 1640 with 2 mM L-glutamine (Corning), 10% heat- inactivated FBS (Sigma-Aldrich, St. Louis, MO), 100 IU penicillin-streptomycin

(Corning), 10 mM HEPES (Corning), 4500 g/L D-glucose (Sigma-Aldrich), 1 mM sodium pyruvate (Corning), and 0.05 mM 2-mercaptoethanol (Gibco, Gaithersburg, MD). Cells were differentiated into adherent macrophages by adding 100 nM phorbol 12-myristate

13-acetate (PMA) (Sigma-Aldrich). After 2 days, the PMA was removed and the cells were washed in PBS and rested overnight in THP-1 medium.

Other cell lines. Lenti-X 293T cells (Clontech, Mountain View, CA) were cultured

and maintained in 293T medium (DMEM (Corning), 10% heat-inactivated FBS (Sigma-

Aldrich), 100 IU penicillin-streptomycin (Corning), 2 mM L-glutamine (Corning), and

0.05% sodium bicarbonate (Corning)). TZM-bl cells, obtained through the NIH AIDS

Reagent Program, Division of AIDS, NIAID, NIH: TZM-bl from Dr. John C. Kappes, Dr.

Xiaoyun Wu, and Tranzyme Inc. [472–476], were cultured and maintained in 293T

medium. ACH-2 cells obtained through the NIH AIDS Reagent Program, Division of

AIDS, NIAID, NIH: ACH-2 from Dr. Thomas Folks [477, 478], were cultured and

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maintained in RPMI 1640 with 2 mM L-glutamine (Corning), 10% heat-inactivated FBS

(Sigma-Aldrich), 100 IU penicillin-streptomycin (Corning), and 10 mM HEPES (Corning).

Cytokines and antibodies

M-CSF, IL-3, and IFN-β were purchased from PeproTech (Rocky Hill, NJ).

Lipopolysaccharide from Escherichia coli O111:B4 was purchased from Sigma-Aldrich.

Anti-IFNAR2 neutralizing antibody was purchased from PBL Assay Science

(Piscataway, NJ). Ultra-LEAF purified mouse IgG2a, κ isotype control antibody was

purchased from BioLegend (San Diego, CA). USP18 [D4E7], p-STAT1-Tyr701 [58D6],

p-STAT2-Tyr690 [D3P2P], and HSP90 [C45G5] monoclonal antibodies, β-actin and

ISG15 polyclonal antibodies, and anti-mouse IgG, HRP-linked and anti-rabbit IgG, HRP- linked secondary antibodies were purchased from Cell Signaling Technology (Danvers,

MA). Anti-HIV-1 p24 Antibody [39/5.4A] was purchased from Abcam (Cambridge, MA).

CD68-PE, CD11b-FITC, CD14-APC, and isotype control flow antibodies were purchased from (eBioscience (San Diego, CA).

HIV-1 molecular clones

pNL4-3-Bal-IRES-HSA (HIVHSA), which has been previously described, was a

kind gift from Dr. Michel J Tremblay [479]. pNL(AD8) (HIVAD) was obtained through NIH

AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, NIH: pNL(AD8) HIV-1 AD8 Macrophage-Tropic R5, 11346 from Fisher BioServices [480].

HIV-1 virus production

Lenti-X 293T cells were cultured in a T75 flask and transfected with pNL4-3-Bal-

IRES-HSA using the Viromer® Red transfection reagent (Lipocalyx, Halle, Sachsen-

Anhalt, Germany) or pNL(AD8) using the FuGENE HD transfection reagent (Promega,

Madison, WI). The transfection reagent was removed after 4 hours and the medium was

56

replenished. After 48 hours, the infectious supernatants were collected, clarified, and filtered through 0.45 µM pore size Whatman PES membranes (Fisher Scientific,

Suwanee, GA). Heat-inactivated FBS (Sigma-Aldrich) was added to the virus at a concentration of 10% and stored at -80° C. After 24 hours, one aliquot of frozen virus was quickly thawed in a 37° C water bath. The virus was titered on TZM-bl cells for 48 hours and a luciferase assay was performed. The titer was determined by the

Spearman-Kärber method [481, 482] and expressed as a TCID50/ml.

TZM-bl cell luciferase assays

The medium was removed by aspiration and 100 μl of room temperature Bright-

Glo™ Luciferase Assay System (Promega) was added to the cells. After 5 minutes, the

cells were lysed and the lysate was transferred to a black Costar EIA/RIA polystyrene

half area 96-well plate (Corning). Luminescence was measured with the VICTOR™ X4

Multi-Plate Reader (PerkinElmer, Waltham, MA).

ELISA

CXCL10 was measured in cell-free supernatants by Human CXCL10/IP-10

DuoSet ELISA kit according to the manufacturer’s instructions (R&D Systems,

Minneapolis, MN). HIV-1 Gagp24 was measured in cell-free supernatants lysed with

0.5% Triton X-100 in PBS by HIV-1 Gagp24 ELISA kit according to the manufacturer’s

instructions (Sino Biological, Beijing, China).

Microarrays

HIV arrays. MDMs were infected with 500 TCID50 HIVAD for 7 days. Total RNA

was collected with RNeasy Mini Kit (Qiagen, Valencia, CA). Gene expression was

assessed with GeneChip™ Human Exon 1.0 ST Arrays (Affymetrix, Santa Clara, CA) by the Interdisciplinary Center for Biotechnology Research at the University of Florida.

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IFN arrays. MDMs were transfected with siRNA for 3 hours and treated with IFN-

β for 18 hours. Total RNA was collected with RNeasy Plus Mini Kit (Qiagen). Gene

expression was assessed with GeneChip™ Human Transcriptome Array 2.0 arrays

(Affymetrix) by the Interdisciplinary Center for Biotechnology Research at the University

of Florida.

Microarray Analysis. The analysis was performed with Partek Genomics Suite

v. 6.6 (Partek Inc., St. Louis, MO). CEL files were imported with GC correction and

intensity data were transformed to log base 2. Exons were summarized to genes using

the median method and an ANOVA with contrast was performed to determine fold

changes between control and treatment groups.

Western Blotting

Cells were washed with PBS and lysed in 1X Cell Lysis Buffer (Cell Signaling

Technology) with Protease Inhibitor Cocktail (Sigma-Aldrich) and 1 mM PMSF (Sigma-

Aldrich). Proteins were separated by size by SDS-PAGE with 4-20% Mini-PROTEAN®

TGX™ precast gels (Bio-Rad, Hercules, CA) and electroblotted onto Trans-Blot®

Turbo™ Mini PVDF membranes (Bio-Rad) using the Trans-Blot® Turbo™ Transfer

System (Bio-Rad) according to the manufacturer’s instructions. Membranes were blocked in 5% non-fat milk in TBST. The Super Signal® West Dura Extended Duration

Substrate kit was used for chemiluminescence detection of HRP-linked secondary antibodies. Blots were exposed to CL-Xposure™ Film (Thermo Scientific, Waltham,

MA). Blots were stripped by incubating with Restore™ PLUS Western Blot Stripping

Buffer (Thermo Scientific).

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shRNA Lentivirus

Human GIPZ lentiviral shRNA gene set for USP18-specific shRNA were purchased from GE Dharmacon (Lafayette, CO). The shRNA clones were screened in

HEK 293 cells and clone V3LHS_645758 (USP18-5) was determined to most efficiently knockdown USP18 (data not shown) and was used for all subsequent experiments. GIPZ plasmid DNA was co-transfected into Lenti-X 293T packaging cells cultured in 293T medium with Tet System approved FBS (tetracycline-free) using the

Lenti-X HTX Packaging System (Clontech) and Xfect Transfection Reagent (Clontech) according to the manufacturer’s instructions. The transfection reagent was removed 24 hours post-transfection. After 48 hours, supernatants containing VSV-G pseudotyped lentivirus were harvested and clarified by centrifugation at 20,000 x g for 18 hours. The supernatant was decanted and the pellet was resuspended in THP-1 medium. THP-1 cells were transduced with TransDux (System Biosciences, Mountain View, CA) by spinoculation for 1 hour at 1,200 x g at room temperature. Puromycin (1 μg/ml) was used to select for transduced cells 48 hours post-transduction. siRNA Transfections

ON-TARGETplus Non-targeting Pool control siRNA (D-001810-10-05) and ON-

TARGETplus USP18 siRNA SMARTpool (L-004236-00-0005) were purchased from GE

Dharmacon. siRNA was transfected into cells with Viromer® Blue transfection reagent

(Lipocalyx) at a concentration of 25 nM according to the manufacturer’s instructions.

RT-qPCR

Total RNA was isolated with RNeasy Plus Mini Kit (Qiagen). cDNA synthesis was carried out with High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems,

Foster City, CA) according to the manufacturer’s instructions. Pre-designed

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PrimeTime® qPCR Assays for IFIT1, IFIT2, IFIT3, IFITM1, IFITM2, USP18, CXCL9,

CXCL10, ISG15, and MX2 were purchased from Integrated DNA Technologies

(Coralville, Iowa). RT-qPCR reactions were carried out in TaqMan Universal PCR

Master Mix (Thermo Fisher Scientific, Waltham, MA) according to the manufacturer’s instructions. RT-qPCR reactions were performed on the StepOnePlus™ Real-Time

PCR System (Applied Biosystems).

Reprogramming CD34 to iPSCs

Donor CD34+ hematopoietic progenitor cells were isolated from peripheral blood

utilizing the EasySep Complete Kit for Human Whole Blood CD34+ Cells (Stem Cell

Technologies, Vancouver, Canada). Isolated CD34+ cells were expanded in StemSpan

SFEM II Medium plus StemSpan CD34+ Expansion supplement for one week. Following

expansion, cells were infected with Sendai viral vector SeVdp(KOSM)302L encoding 4

reprogramming factors (OCT4, SOX2, KLF4, and c-MYC) [483]. Infection was carried

out for 2 hours at 37° C at an MOI of 2. Cells were reprogrammed on Matrigel-coated

dishes using ReproTeSR (Stem Cell Technologies) following the manufacturer’s

instructions. Approximately three weeks after infection, 6 iPSC colonies were manually

isolated to generate iPSC clones. iPSC clones were maintained and expanded on

Matrigel-coated plates in mTeSR1 (Stem Cell Technologies). iPSCs were characterized by assessing morphology, expression of stem cell markers OCT4 and SSEA4 by flow cytometry analysis, and karyotyping. Pluripotency was confirmed by differentiating the cells to ectoderm, endoderm, and mesoderm lineages using the STEMdiff Trilineage

Differentiation Kit (Stem Cell Technologies).

Flow cytometry was used to examine undifferentiated iPSCs stained with OCT4-

PE, SSEA4-APC, and SSEA1-PE antibodies (BioLegend). For trilineage differentiated

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iPSCs, ectoderm was assessed by NESTIN-PE and PAX6-Alexa Fluor 647 (BD

Biosciences, San Jose, CA). Endoderm was assessed by FOXA2-PE and SOX17-Alexa

Fluor 647 (BD Biosciences). Mesoderm was assessed by Brachyury-APC (R&D

Systems) and NCAM-PE (Stem Cell Technologies).

Differentiation of iPSCs to Monocytes

Differentiation was carried out as previously described [484] with slight modifications. Briefly, iPSCs were passaged sparsely to 6-well Matrigel-coated plates and cultured for 8-10 days in mTeSR1 with daily medium changes until the colonies were approximately 1 cm in diameter. Next, embryoid bodies (EBs) were formed by gently lifting colonies using a cell lifter. Colonies were gently transferred to a 15 ml conical tube using a 10 ml serological pipette. Colonies were allowed to gravity settle for approximately 5 minutes and supernatant was aspirated. Cells were gently resuspended in mTeSR1 supplemented with 10 μM ROCK inhibitor (Y-27632) and transferred to Ultra-Low Attachment Surface 6-well plates (Corning) containing 4 mL medium per well. EBs were allowed to form for 4 days with a partial (2/3) medium change after 48 hours. On day 4, EBs were collected, washed, resuspended in X-

VIVO15 supplemented with GlutaMAX™ (Gibco) and 2-mercaptoethanol (Gibco), and transferred to adherent, cell culture treated plates. iPSC USP18 Gene knockout using CRISPR/Cas9

iPSCs were nucleofected using the Human Stem Cell Nucleofector Kit 1 (Lonza,

Basal, Switzerland) following manufacturer’s recommendations with three plasmids: one plasmid that encodes S. pyogenes Cas9, one plasmid that contains the USP18 gRNA sequence GCAAATCTGTCAGTCCATCC, and a donor plasmid with homology arms that flank the CRISPR site in USP18 exon 2 that delivers a GFP and puromycin

61

resistance gene cassette. Briefly, equal amounts of each of the three plasmids (5 μg total) were nucleofected into 8 x 105 cells using Nucleofector II (Amaxa Biosystems,

Cologne, Germany) program B-016. Cells were re-plated on a Matrigel-coated dish with mTeSR1 and allowed to recover for 3 days with daily medium changed. GFP fluorescence was observed after 24 hours. On day 3, the cells were maintained under puromycin (0.3 μg/ml) selection for 30 days. Ten clones were manually isolated and expanded in mTeSR1. Genomic DNA was isolated using DNeasy Blood & Tissue Kit

(Qiagen) for screening. Primer sequences for screening for CRISPR modifications are in Supplemental Table 1.

DQ-Ovalbumin and Phagocytosis Assays

DQ-Ovalbumin (Invitrogen, Carlsbad, CA) was added to the cells at a concentration of 50 μg/ml. NucBlue® Live ReadyProbes® Reagent (Molecular Probes,

Eugene, OR) were also added to the well and incubated at 37° C for 25-30 minutes.

The cells were washed with pre-warmed PBS and fixed in 4% paraformaldehyde. pHrodo Red Bioparticles were resuspended in Live Cell Imaging Solution (Gibco) at 1

μg/ml and sonicated for 5 minutes in a water bath sonicator. The resuspended pHrodo

Red-labeled heat-killed Escherichia coli or Staphylococcus aureus were added at a final

concentration of 1 μg/ml in Live Cell Imaging Solution to macrophages and incubated at

37° C for 1-2 hours to allow for uptake and acidification of phagosomes. NucBlue® Live

ReadyProbes® Reagent (Molecular Probes) was added before imaging. All fluorescent

images were obtained with the EVOS™ FL Cell Imaging System (Invitrogen).

Macrophage Morphology

MDMs or iMacs were washed in PBS and fixed in 4% cold paraformaldehyde for

10 minutes. The cells were washed again in PBS and permeabilized in 0.1% Triton X-

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100. The cells were washed with PBS again and then stained with either ActinGreen™

488 ReadyProbes® or ActinRed™ 555 ReadyProbes® reagent (Molecular Probes) for

30 minutes followed by two more washes in PBS. The cells were then imaged with the

EVOS™ FL Cell Imaging System (Invitrogen).

Flow Cytometry

Fc receptors were blocked with Fc Block (Miltenyi) and stained with antibodies in

PBS with 1% human serum. For intracellular staining of CD68, the cells were fixed and

permeabilized with the BD Cytofix/Cytoperm™ kit (BD Biosciences). All samples were

analyzed with the BD Accuri™ C6 Cytometer (BD Biosciences).

Integration Assay

The integration assay for detection of integrated HIV-1 proviral DNA was adapted

from methods previously described [485, 486]. Briefly, genomic DNA was isolated with

the DNeasy Blood and Tissue Kit (Qiagen). A pre-amplification step with 200 nM Alu1,

Alu2, and ULF2 primers was performed with 200 ng of genomic DNA in AmpliTaq

Gold® 360 Master Mix with 360 GC Enhancer (Applied Biosystems) and incubated in a

thermal cycler at 95° C for 10 minutes followed by 20 cycles of 95° C for 30 seconds,

55° C for 30 seconds, and 72° C for 25 seconds, and a final extension step at 72° C for

7 minutes. A second round of PCR was performed with 1/100th of the product from the

pre-amplification step with 200 nM Lambda T2 and UR2 primers and 150 nM Fam-Int-

HIV-IABkFQ TaqMan probe in TaqMan® Universal Master Mix (Applied Biosystems) at the thermal cycler conditions recommended by the manufacturer for the master mix. In parallel to the samples, a standard curve also underwent the same pre-amplification and second amplification steps.

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The standard curve was prepared from ACH-2 cells, which is a T cell line that has exactly one integrated HIV-1 proviral genome. The standard curve was prepared by making 1:10 dilutions of 2000 ng of genomic DNA from ACH-2 cells. Carrier DNA from uninfected PBMCs was added so that the total genomic DNA in each standard was

2000 ng. Primer sequences are listed in Table 3-1. Using an estimate of ~6.6 pg of

genomic DNA per cell, the number of copies of integrated proviral genomes per 10,000

cells was calculated from the standard curve.

Results

T1 IFNs restrict HIV-1 replication

It has been demonstrated previously that in vitro T1 IFNs restrict HIV-1

replication [313, 316, 317, 409, 410, 487]. To confirm these previous findings, we

pretreated TZM-bl cells with different doses of IFNs for 24 hours and infected the cells

with HIV-1. As previously reported, T1 IFN restricted HIV-1 replication in a dose- dependent manner (Figure 2-1). We tested this in MDMs and found that HIV-1 replication is potently restricted by IFN-β in a dose-dependent manner (Figure 2-2A) with an IC50 of 504.7 fg/ml (± 142.1 fg/ml).

Based on our previous work showing the importance of STAT signaling for

replication [191] we predicted that some level of IFN-induced signaling through the JAK-

STAT pathway is necessary for efficient replication. To test this, MDMs were pretreated

with α-IFNAR neutralizing antibodies and infected with HIV-1. When T1 IFN signaling was blocked, HIV-1 replication was restricted as measured by Gagp24 ELISA on supernatants (Figure 2-2B). These data demonstrate that HIV-1 must strike a balance

between some T1 IFN signaling, which is needed for producing transcription factors

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such as STAT1 and STAT3, and too much IFN signaling, which allows the cells to mount an effective antiviral response.

HIV-1 induces an interferon-like response in infected monocyte-derived macrophages (MDMs)

Since HIV-1 requires some IFN signaling to replicate efficiently, HIV-1 must be triggering an IFN response, which would be evident by expression of ISGs. To confirm this, MDMs were infected with HIV-1 for seven days and gene expression was analyzed by microarray (Figure 2-3A). We observed that all genes upregulated by HIV-1 at least

2-fold (Table 3-2) have been reported to be IFN-inducible in the Interferome (v2.01) database (Figure 2-3B).

Interestingly, HIV-1 induces expression of USP18 (Table 3-2), a negative regulator of T1 IFN signaling [426]. Western blot analysis confirmed this at the protein level with two different strains of HIV-1 (Figure 2-4A). Given that USP18 attenuates the

IFN response, we hypothesized that induction of USP18 allowed HIV-1 to achieve the balance needed to provide the necessary IFN signaling, without too strong an antiviral response.

USP18 induction by HIV-1 infection requires activity of T1 IFNs

It is unknown how HIV-1 induces USP18 expression in human macrophages.

Since USP18 is an ISG, we first pretreated MDMs with α-IFNAR neutralizing antibodies then infected the cells to see if USP18 was still induced by HIV-1 in the absence of T1

IFN signaling. The results show that USP18 is not induced by HIV-1 in the absence of

T1 IFN signaling (Figure 2-4B), providing evidence that HIV-1 does induce an IFN response in macrophages that have an autocrine effect.

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While it is known that USP18 is induced by T1 IFNs, Malakhova and colleagues showed that USP18 is induced after LPS treatment and that this was dependent on two interferon regulatory factor (IRF) binding sites: IRF-2 and IRF-3 [488]. However, this study was in mouse macrophages and did not rule out USP18 induction by T1 IFNs as a secondary response to LPS treatment. To determine if LPS could induce USP18 in human macrophages independently of T1 IFNs, human MDMs were treated with LPS in

the presence or absence of α-IFNAR neutralizing antibody. When USP18 expression

was assessed by Western blot, USP18 was induced by LPS treatment, but at much

lower levels when IFNAR was blocked by neutralizing antibodies (Figure 2-4C). These

data show that in humans, LPS-induced USP18 is secondary to T1 IFNs produced in

response to LPS and not a direct consequence of LPS stimulation.

USP18 knockdown restricts HIV-1 replication

We next wanted to determine if USP18 expression is necessary for HIV-1

replication. To test this, we used siRNA to knockdown USP18 in THP-1 cells. THP-1

cells are suspension cells that can be differentiated into adherent macrophage-like cells

with PMA treatment. They are a well-established model system for studying HIV-1

infection in macrophages [489]. THP-1 cells were differentiated with PMA first,

transfected with siRNA, and infected with HIV-1. Knockdown of USP18 worked

efficiently with this method (Figure 2-5A). However, we found that transfection with

siRNA rendered THP-1 cells refractory to HIV-1 infection, even with the non-targeting

control siRNA (data not shown).

TZM-bl cells, however, were not refractory to HIV-1 after transfection with siRNA.

Using this model, we transfected TZM-bl cells with non-targeting control or USP18

siRNA, treated the cells with IFN-β for 24 hours, and infected the cells with HIV-1.

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USP18 deficiency enhanced IFN-mediated restriction of HIV-1 (Figure 2-5B).

Transfecting MDMs with siRNA allowed for efficient knockdown of USP18 (Figure 2-

5C), but also made the cells refractory to infection (Figure 2-5D).

To overcome the limitation of siRNA, we generated THP-1 cell lines that constitutively express non-targeting control or USP18-specific shRNA (Figure 2-6A-B).

After PMA differentiation, THP-1 cells expressing non-targeting control or USP18

shRNA were infected by HIV-1. We found that HIV-1 replication was significantly

restricted in THP-1 cells with USP18 knockdown. There was significantly less p24

detected in the supernatants and in the protein lysates of USP18 knockdown cells

compared to control (Figure 2-7A-B).

USP18 deficiency enhances IFN signaling through JAK-STAT pathway

We wanted to investigate the mechanism by which USP18 deficiency results in

restricted HIV-1 replication in macrophages. USP18 is part of the normal negative

feedback response that regulates T1 IFN signaling through IFNAR and JAK-STAT

signaling. In the absence of USP18, signaling through the JAK-STAT pathway should

be enhanced with increased levels of phosphorylated STAT1 and STAT2. We treated

THP-1 cells with IFN-β for 18 hours and found increased levels of phosphorylated

STAT1 and STAT2 (Figure 2-7C) as well as increased levels of ISGylated proteins

(Figure 2-7D). While siRNA knockdown in MDMs makes them refractory to HIV-1

infection, these cells can still be used for studying T1 IFN signaling. We also measured

STAT1 and STAT2 phosphorylation in USP18 knockdown MDMs treated with IFN-β and

found similar results to THP-1 cells where STAT1 and STAT2 showed elevated

phosphorylation when USP18 was deficient (Figure 2-8A).

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USP18 deficiency enhances expression of ISGs

Increased signaling through STAT1 and STAT2 should result in enhanced expression of ISGs. To test this, microarray analysis was performed on MDMs transfected with USP18 siRNA and treated with IFN-β for 18 hours (Figure 2-8B).

Overall, the genes that were induced by IFN were induced at a higher level when

USP18 was knocked down and the genes that decreased expression during IFN treatment decreased expression more when USP18 was knocked down. The genes that were upregulated by 2-fold or greater with USP18 knockdown and IFN treatment are in listed in Figure 2-8B.

To probe more deeply into specific antiviral genes that are part of the overall ISG response, shRNA expressing THP-1 cells and MDMs transfected with USP18 siRNA were treated with IFN-β for 18 hours and gene expression of ISGs was analyzed by qPCR. In a qPCR panel looking at IFIT1, IFIT2, IFIT3, IFITM1, IFITM2, CXCL9,

CXCL10, ISG15, and MX2, all genes had increased expression when USP18 was knocked down compared to control (Figure 2-9A-B). USP18 expression was also measured to confirm knockdown of USP18 transcripts. iPSC-derived macrophages support HIV-1 replication

While THP-1 cells and MDMs with siRNA knockdown provide useful models for investigating the effects of USP18 deficiency on IFN signaling, it may not be the best model. THP-1 cells are transformed cells with an abnormal karyotype [490] and in our hands, they do not remain differentiated and adherent in culture for more than 5 days

(data not shown). While we could achieve efficient knockdown of USP18 in MDMs, the transfected cells became refractory to infection even when a non-targeting control

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siRNA was used. To overcome these obstacles, we sought to use an induced pluripotent stem cell-derived macrophage model.

iPSC-derived macrophages have been previously shown to be a suitable model to study HIV-1 [484]. First, we generated iPSCs from CD34+ cells isolated from

peripheral blood of healthy adult human donors (Figure 2-10). The CD34+ cells were

expanded in vitro and transduced with a non-integrating Sendai viral vector

(SeVdp(KOSM)302L) encoding Oct4, Sox2, Klf4, and c-Myc to reprogram them into iPSCs[483, 491]. For differentiation, we utilize an embryoid body (EB) method in which

EBs are formed and cultured in the presence of IL-3 and M-CSF to generate monocytes

[484]. The monocytes are then cultured in M-CSF alone to produce MDMs from iPSCs

(iMacs).

iMacs are CD14+, CD11b+, and CD68+ and have similar morphology to MDMs

(Figure 2-11A-B). To confirm that iMacs also function like MDMs, iMacs were treated with DQ-OVA, which is an ovalbumin that is fluorescently labeled with an intramolecular quencher. If the protein is phagocytosed and processed by acidic proteases, it will be digested freeing the fluorophore and quencher allowing for fluorescence. iMacs fed DQ-

OVA were positive for BODIPY FL dye indicating that ovalbumin was processed by acidic proteases (Figure 2-11C). To determine if iMacs can phagocytose whole bacteria and process them in acidified phagosomes, heat killed E. coli and S. aureus labeled with a pH sensitive dye were fed to iMacs. After two hours, the iMacs were RFP positive indicating that they phagocytosed the bacteria and that the phagosome was acidified

(Figure 2-11C). iMacs can also support HIV-1 replication as previously reported [484] and USP18 is induced by HIV-1 in iMacs (Figure 2-12A-B). Since USP18 is induced by

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HIV-1 in iMacs in a similar manner as MDMs, we concluded that iMacs are a better model for studying USP18 knockout/knockdown than THP-1 cells.

CRISPR knockout of USP18 in iMacs

To test the effects of USP18 deficiency in iMacs, we utilized CRISPR/Cas9 gene editing to knockout USP18 in iPSCs before differentiation to the myeloid lineage. The advantage of knocking out USP18 in iPSCs is that once knockout is achieved we have

a self-renewing iPSC line that can be a continuous source of new EBs for making

iMacs.

To knockout USP18, iPSCs were transfected with three plasmids that delivered a

USP18 targeting single-guide RNA (sgRNA), Cas9, and a donor plasmid with homology arms to the region where the sgRNA targets USP18 in exon 2 (Figure 2-13A). The donor DNA contains a GFP and puromycin resistance gene allowing for selection of successfully modified cells. Puromycin-resistant clones were screened by PCR and

Western blot for successful knockout of USP18 (Figure 2-13B-C). Two knockout clones

(GFP+ clone 5 and GFP- clone 2) were obtained and used in subsequent experiments.

To determine if USP18 knockout (USP18-/-) iMacs have a similar phenotype to

USP18 knockdown THP-1 and MDMs, USP18-/- iMacs were treated with IFN-β and

levels of phosphorylated STAT1 and STAT2 were measured by Western blot. USP18-/-

iMacs had increased and sustained phosphorylated STAT1 and phosphorylated STAT2

(Figure 2-14A). Since USP18-/- iMacs also had increased STAT1 and STAT2 signaling,

we wanted to determine if USP18-/- iMacs also had enhanced ISG expression after IFN-

β treatment. Indeed, after treatment with IFN-β for 18 hours, USP18-/- iMacs had

increased expression of ISGs compared to USP18 sufficient cells (Figure 2-14B). These

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findings are consistent with the results from THP-1 cells and MDMs with USP18 knockdown.

HIV-1 replication is restricted in USP18 knockout iMacs

Next, we wanted to determine if USP18 knockout influenced HIV-1 replication.

USP18 knockout results in an enhanced response to IFN stimulation in macrophages.

Therefore, we hypothesized that HIV-1 replication would be restricted in USP18-/- iMacs

due to enhanced IFN signaling in response to infection. iMacs were infected with HIV-1

and there was a significant reduction in supernatant p24 and intracellular p24 as

determined by ELISA and Western blot in USP18-/- iMacs at days four and eight post-

infection (Figure 2-15A-B).

We also wanted to determine if the restriction in HIV-1 replication was due to

fewer cells becoming infected during the first round of infection or if it was due to

decreased spreading. To test this, iMacs were infected and treated with Darunavir, an

HIV-1 protease inhibitor, to prevent second rounds of infection. After 24 hours, genomic

DNA was collected and a qPCR assay developed to measure integrated proviral

genomes showed that there were on average 3,800 genomes per 10,000 cells in

USP18+/+ iMacs and on average 4,900 integrated genomes per 10,000 cells in USP18-/- iMacs treated with Darunavir 24 hours post-infection. This suggests that the decrease in viral replication seen over time at day 4 and 8 is due to restricted production and spreading of the virus and not due to decreased or inhibited entry into cells.

Discussion

In this study, we have demonstrated that depletion of USP18 in human macrophages restricts HIV-1 replication. We determined that this was due to an enhanced IFN response from a lack of feedback inhibition of the JAK/STAT pathway

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resulting in increased expression of ISGs that have antiviral effects. Previous work done by our group has shown that STAT1 and STAT3 are necessary for efficient HIV-1 replication. Blocking phosphorylation of either of these two molecules resulted in significant reduction in viral replication [191]. However, the current study shows that increasing the amount of phosphorylated STAT1 and STAT2 also results in a significant reduction in viral replication. Thus, we propose that USP18 acts as a tuning mechanism that prevents a robust IFN response that would restrict viral replication (Figure 2-16).

T1 IFN secretion is induced by HIV-1, but at very low levels not detectable by microarray after 7 days of productive HIV-1 infection of macrophages. Clearly, the relatively small amounts of IFN produced early in infection have a meaningful effect on virus replication. When IFNAR blocking antibody was used, HIV-1 infection was reduced rather than enhanced, demonstrating how low levels of IFN can benefit the virus. At the same time, USP18 is not induced by HIV-1 when anti-IFNAR is included, demonstrating that low-level T1 IFNs being produced are responsible for USP18 induction as well as the IFN-like transcriptional response. By providing exogenous IFN-β, we were able to demonstrate that lack of USP18 makes cells more sensitive to the effects of IFN.

Previous work in murine macrophages suggests that USP18 can be induced by LPS through IRF-3 alone, but did not rule out the possibility that T1 IFNs were responsible for inducing USP18 [488]. Here we show that in human macrophages USP18 can be induced by LPS, but that the effect is significantly reduced in the presence of IFNAR neutralizing antibody.

The strategy used by HIV-1 for transcription from its LTR promoter utilizes transcription factors such as NFκB, NFAT, and IRFs, [492, 493] that would normally be

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present in a cell that has detected a viral infection and has initiated an antiviral response. By utilizing this transcriptional machinery, HIV-1 has been able to hijack normal immune processes to improve its own fitness, however, in vitro experiments

have long demonstrated that inflammatory activators like LPS or IFNs can potently

restrict HIV-1 replication. Thus, the physiological state that supports HIV-1 replication is

a window of low-level immune activation that is due, at least in part, to balanced positive

and negative regulators of antiviral immunity (Figure 2-16). Infected cells eventually

become refractory to T1 IFN signaling due to induction of negative regulators such as

USP18. Other negative regulators are also induced by T1 IFNs including SOCS1,

SOCS3, and PIAS [288]. We focused on USP18 for this study due to its additional

function as a deISGylating enzyme and its appearance in gene expression profiling

experiments following HIV infection.

It is unknown from our study exactly how HIV-1 is inducing production of T1

IFNs. The IFI16 and cGas-STING pathways can sense HIV-1, which could lead to the

production of T1 IFNs and promote pyroptosis [165–167]. It has also been

demonstrated in monocyte-derived macrophages and monocyte-derived dendritic cells

that HIV-1 Vpr can differentially regulate expression of ISGs, including USP18, MX1,

MX2, ISG15, ISG20, IFIT1, IFIT2, IFIT3, IFI27, IFI44L, and TNFSF10 [494, 495]. Many

of these genes were induced by HIV-1 in MDMs in our current study (Table 3-2). So,

while there may be multiple mechanisms by which HIV-1 prevents a fully robust IFN

response, induction of negative regulators such as USP18 play an important role in this

process. If HIV-1 accessory protein Vpr alone can induce a T1 IFN response then it

demonstrates the importance of T1 IFN signaling for HIV-1 replication.

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Our work demonstrates that knockdown of USP18 by shRNA restricts HIV-1 replication. Previous work has shown that Usp18-/- mice have increased protection

against intracerebral infection by LCMV and VSV [428]. These mice were shown to

have an increased ability to restrict Salmonella typhimurium replication as well as

increased survival. However, these mice are hypersensitive to LPS due to an overactive

T1 IFN response [429]. These studies showed that USP18 deficiency resulted in increased ISGylation of proteins and enhanced phosphorylated STAT1 levels in response to LPS treatment or viral infection.

Human USP18 functions similarly to mouse Usp18. However, human USP18 is

ISGylated, which promotes stability of USP18. USP18 and ISG15 knockout cells have

similar enhanced ISG responses in humans due to the fact that in ISG15 deficiency,

USP18 is not stable [453, 454]. Infants born with naturally occurring USP18 deficiency

have pseudo-TORCH syndrome due to aberrant IFN responses in the absence of

congenital infections [432].

Our current work shows that USP18 deficient human macrophages also have

increased ISGylation and enhanced levels of phosphorylated STAT1 and

phosphorylated STAT2 after treatment with IFN-β. We expected that if there was enhanced signaling through IFNAR and STAT1/STAT2 that there would be increased expression of ISGs. Indeed, we found that in USP18 knockdown cells expression of

ISGs was enhanced after treatment with IFN-β. This is consistent with work done in murine macrophages treated with IFN-β that demonstrated an increase in genes involved in antigen processing and presentation, cytokines and chemokines, as well as genes involved in responses to viral infections [431]. That study demonstrated not only

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increased but also sustained expression of genes involved in responses against viruses. We demonstrate in the current study that human macrophages with USP18 knockdown and USP18-/- iMacs have a similar phenotype.

ISG15 and ISGylation of proteins have been previously shown to be important in

restricting many different viruses including VSV, HIV-1, HCV, Chikungunya, Influenza A,

Influenza B, LCMV, and Sindbis viruses [496, 497]. Previous studies have shown that

ISGylation is important for restricting HIV-1 in 293T, U1.1, HOS-CD4/CXCR4, HeLa,

U2OS, and 293E cell lines [365, 366, 368]. However, we are unaware of any previous

studies showing that relevance of ISGylation in primary cells or non-immortalized cells

such as human MDMs, human T cells, or a model like iMacs. The current study does

not prove that ISGylation plays a role in HIV-1 restriction in macrophages, however, it

does demonstrate that in the absence of USP18 ISGylation is enhanced. In addition to

previous work done in cell lines, it is likely that the enhanced ISGylation response

shown in IFN-β treated cells with USP18 deficiency contributes to the restriction of HIV-

1. Moving forward, a key area of focus should be determining which functional domains

of USP18 are required for HIV-1 replication to dissect the relative roles of deISGylation

and IFNAR blocking in this process.

We have demonstrated in USP18 deficient cells that the first round of infection is

unaffected in its ability to enter the cells and integrate into the host chromosome. After

multiple rounds of infection, there is a difference in integration (data not shown). This

may be because during the first round of infection the IFN pathway has not yet been

activated. After sensing of viral DNA in the infected cells, however, IFNs are produced

and secreted from the cells. T1 IFNs can then signal back to the same cell in an

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autocrine manner or to neighboring cells in a paracrine manner. Signaling back to already infected cells and activating ISGs can shut down machinery that is utilized by the virus for replication. This effect may simply be enhanced in USP18 deficient cells due to increased JAK/STAT signaling.

Uninfected cells that have received paracrine signals from T1 IFNs, in addition to shutting down machinery used by the virus for transcription and translation of viral proteins, may also upregulate factors that prevent cells from binding such as the IFITM family of proteins. The IFITM family members have been shown to inhibit HIV-1, by interfering with viral fusion or binding [356, 357, 360]. This could potentially result in fewer cells being infected. This effect may simply be enhanced in the absence of

USP18.

iPSC-derived cell models have previously been used to study other diseases in a variety of different cell types, including vascular smooth muscle cells, cardiomyocytes, and neurons [498–501]. iPSC-derived macrophages have previously been shown to be a useful model for HIV-1 infection [484] and we have confirmed those findings in the

current study. Our iMac model supports very robust HIV-1 replication and shows a similar induction of USP18 in response to infection as what occurs in MDMs.

The use of iPSC-derived cells has several advantages over conventional cell models. First, iPSC technologies can be used to derive cells that may not be obtainable from living human donors, such as endothelial cells. Second, iPSCs can be grown indefinitely in culture allowing for repeated experiments from the same original donor.

This helps to eliminate donor variability or the difficulty recalling previous donors. Third, iPSC-derived cells can be genetically modified by use of gene editing technologies such

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as CRISPR/Cas9 and allow for a renewable source of edited cells. This eliminates the need to constantly treat primary cells like MDMs with siRNA and has the advantage of being able to have a knockout and an unedited cell line derived from the same donor.

In conclusion, we have shown in a novel iPSC-derived macrophage model that

USP18 is required for HIV-1 replication in vitro. We propose that durable and enhanced

IFN signaling can restrict HIV-1 replication in vivo, but only if certain elements of the

ISG response are limited. Negative regulators serve an important cellular function by

limiting the magnitude of an initial response or helping to resolve a response following

early signaling events. Here we show how HIV-1 capitalized on the biologic effects of

one negative regulator, USP18, as a means of tuning the T1 IFN response for increased

viral fitness. Many studies have focused on the antiviral effects of ISGs, but

understanding how host cells regulate antiviral pathways will unveil more complete

biological mechanisms. Many infections or hereditable diseases are not easily modeled

in rodents, so new platforms are required to answer key questions about host

gene/proteins including negative regulators of signaling. We have demonstrated that

iPSC-derived cell models when paired with genome editing technologies, are a powerful

tool for studying not only HIV-1 infection, but can also be used to study other infectious

disease models, autoimmunity, and other general biological problems that lack good

conventional cell line models.

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Figures

Figure 2-1. Type I IFN restricts HIV-1 replication in TZM-bl cells. TZM-bl cells were plated in 96-well plates (10,000 cells/well) and allowed to adhere overnight. The following day, the cells were pre-treated with IFN-β for 24 hours and then infected with 100 TCID50 HIVHSA. After 48 hours, the amount of HIV-1 replication was measured by luciferase assay. The dashed line indicates the level of luciferase activity in cells that were not pretreated with IFN.

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Figure 2-2. IFNAR blocking inhibits HIV-1 replication. (A) MDMs were pretreated with IFN-β for 24 hours and infected with HIVHSA for 7 days. Supernatant Gagp24 was measured by ELISA. (B) MDMs were infected with 1000 TCID50 HIVAD in the presence of α-IFNAR2 neutralizing antibody or IgG2a isotype control antibody. Supernatants were harvested on days 4 and 8 post-infection and supernatant Gagp24 was measured by ELISA.

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Figure 2-3. HIV-1 induces an IFN-like response in MDMs. MDMs were infected with HIVAD for 7 days and gene expression was assessed by Affymetrix arrays. (A) Heat map showing expression of genes induced by HIVAD infection. (B) Venn diagram showing genes that were induced by HIVAD by 2-fold or greater are also inducible by IFNs. Information based on Interferome (v2.01) database.

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Figure 2-4. USP18 expression is induced by HIV-1 and is dependent on type I IFN signaling. (A) MDMs were infected with 5000 TCID50 HIVHSA or HIVAD for 7 days. USP18 and Gagp24 expression was assessed by Western blot. (B) MDMs were infected with 1000 TCID50 HIVAD in the presence of α-IFNAR2 neutralizing antibody or IgG2a isotype control antibody. Expression of USP18 and Gagp24 on day 4 post-infection was measured by Western blot. The experiment was performed on 3 donors with similar results. A representative blot is shown. (C) MDMs were pretreated with α-IFNAR2 neutralizing antibody or IgG2a isotype control antibody for 1 hour and then treated with LPS (10 ng/ml) for 24 hours. USP18 expression was measured by Western blot in three donors.

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Figure 2-5. USP18 knockdown by siRNA makes THP-1 cells and MDMs, but not TZM- bl cells, refractory to HIV-1 infection. (A) PMA activated THP-1 cells were transfected with control or USP18 siRNA or transfection reagent only (VB Only) followed by IFN-β treatment for 18 hours. USP18 expression was measured by Western blot. (B) TZM-bl cells were transfected with control or USP18 siRNA for three hours followed by IFN-β treatment at different doses for 24 hours. TZM-bl cells were infected with 150 TCID50 HIVHSA for 48 hours and HIV-1 replication was measured by Luciferase assay. (C) MDMs were transfected with control or USP18 siRNA for three hours followed by IFN-β treatment for 18 hours. USP18 expression was measured by Western blot to confirm knockdown. (D) MDMs treated with siRNA for six hours were infected with 1000 TCID50 HIVHSA for four days. Supernatant Gagp24 was measured by ELISA.

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Figure 2-6. USP18 knockdown by shRNA in THP-1 cells. THP-1 cells were transduced with GIPZ lentiviral vectors that express shRNA and GFP from the same transcript. (A) After PMA activation, the CMV promoter becomes very active and GFP is readily detectable by epifluorescence microscopy. (B) USP18 expression was measured by Western blot to confirm successful knockdown by shRNA.

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Figure 2-7. shRNA knockdown of USP18 inhibits HIV-1 replication in THP-1 cells and enhances IFN-β-induced STAT activation. THP-1 cells expressing control or USP18-5 shRNA were activated with PMA. (A) THP-1 cells were infected with 5000 TCID50 HIVAD for 5 days and supernatant Gagp24 was measured by ELISA. (B) THP-1 cells were infected with 5000 TCID50 HIVHSA for 5 days and intracellular Gagp24 expression was measured by Western blot. (C) Undifferentiated and PMA activated THP-1 cells expressing control or USP18-5 shRNA were treated with IFN-β for 18 hours. Expression of p- STAT1, p-STAT2, and USP18 was measured by Western blot. (D) ISGylated proteins of various molecular weights and free ISG15 were measured by Western blot. A paired T test was used to determine statistical significance. N.D. = not detected, *** p<0.001.

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Figure 2-8. siRNA knockdown of USP18 enhances STAT activation and modulates the transcriptome in IFN-β-treated MDMs. MDMs from six donors were transfected with non-targeting (NT) control or USP18 siRNA for three hours followed by IFN-β treatment for 18 hours. (A) Expression of p-STAT1, p- STAT2, and USP18 was measured by Western blot. (B) Heat map showing transcriptional profiles assessed by Affymetrix arrays. (C) Gene ontology enrichment of USP18 siRNA + IFN-β vs. NT control siRNA + IFN-β.

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Figure 2-9. ISG expression is enhanced in USP18 deficient THP-1 Cells and MDMs. (A) PMA activated THP-1 cells expressing control or USP18-5 shRNA were treated with IFN-β for 18 hours. Gene expression of ISGs was determined by RT-qPCR (n=4). (B) MDMs from six donors were transfected with control or USP18 siRNA for three hours followed by IFN-β treatment for 18 hours. Gene expression of ISGs was determined by RT-qPCR (n=6). A paired T test was used to determine statistical significance. ns = not significant, * p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001.

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Figure 2-10. Workflow for generating iPSC-derived monocytes. CD34+ cells are isolated from the peripheral blood of healthy human donors (1-3). The CD34+ cells are expanded in vitro and transduced with a Sendai virus that delivers Oct3/4, Sox2, Klf4, and c-Myc (4). These factors induce the CD34+ cells to become pluripotent stem cells (6). These cells can now be modified by CRISPR/Cas9 gene editing or the colonies are scraped from the plate and form embryoid bodies. The embryoid bodies are cultured in the presence of IL-3 and M-CSF and will begin to produce CD14+ monocytes (7). The monocytes are then cultured in M-CSF alone and will differentiate into macrophages. Some images were adapted from Servier Medical Art under a creative commons attribution 3.0 unported license.

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Figure 2-11. Characterization of iMacs. (A) iMacs are CD14+, CD11b+ and CD68+ as measured by flow cytometry. (B) The nuclei (Blue) and actin (red or green) of iMacs and MDMs were stained showing similar morphology. (C) iMacs fed DQ-OVA were able to take up the protein and digest the protein with acidic proteases as indicated by GFP fluorescence (top right panel). iMacs were also fed heat-killed pHrodo red-labeled Escherichia coli and Staphylococcus aureus. The pH-sensitive dye fluoresces red when the phagosome acidifies (bottom panels).

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Figure 2-12. iMacs support HIV-1 replication. iMacs were infected with 1000 TCID50 HIVHSA for 8 days. (A) Supernatant Gagp24 was measured by ELISA. (B) Intracellular Gagp24 and USP18 expression was measured by Western blot.

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Figure 2-13. CRISPR/Cas9 knockout of USP18 in iPSCs. (A) Schematic showing the CRISPR gRNA target site in Exon 2 of USP18. Homology regions flanking the CRISPR site were placed in the donor plasmid with an EF-1α promoter driving expression of a copGFP-T2A-PuroR transcript. After a double- stranded break, homology-directed repair in the presence of the donor plasmid will allow for integration of selection cassette at CRISPR site disrupting expression of USP18. (B) Clones were screened by PCR with three primer sets. Primer set A (P1 and P2) would amplify a 1466 bp product if there was no modification of the USP18 locus. Primer sets B (P1 and P5) and C (P2 and P6) would amplify 1313 bp and 1097 bp products, respectively, if the resistance cassette was integrated at the USP18 locus. Clones that had integration in only one allele had the wild-type allele sequenced to detect INDELs. One lane which had another marker ladder was omitted from the image in the bottom panel. (C) Clones that had INDELs resulting in frameshift or clones that had an insertion of the selectable cassette in both alleles were treated with IFN-β for 18 hours and USP18 expression was measured by Western blot.

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Figure 2-14. USP18-/- iMacs have enhanced STAT phosphorylation. (A) iMacs and MDMs were treated with IFN-β and p-STAT1 and p-STAT2 expression was measured by Western blot 1, 4, and 12 hours after treatment. (B) iMacs were treated with IFN-β for 18 hours and gene expression of ISGs was measured by RT-qPCR (n=3). The average RQ of the USP18-/- clones divided by the average RQ of the USP18+/+ clones is expressed as fold change.

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Figure 2-15. Knockout of USP18 restricts HIV-1 replication in iMacs. (A) USP18+/+ and -/- USP18 iMacs were infected with 1000 TCID50 HIVHSA for 8 days. Supernatant Gagp24 was measured by ELISA (n=5). (B) Intracellular Gagp24 and USP18 were measured by Western blot. Representative blot of three experiments is shown. A paired ratio T test was used to determine statistical significance. ** p<0.01.

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Figure 2-16. USP18 tunes the IFN response to allow for efficient HIV-1 replication. A low-level IFN response in the presence of IFNAR neutralizing antibodies characterized by low levels of phosphorylated STAT1/2 and low expression of ISGs limits HIV-1 replication. In the absence of USP18, a strong IFN response characterized by high levels of phosphorylated STAT1/2 and high expression of ISGs also limits HIV-1 replication. In the presence of USP18, the optimal levels of phosphorylated STAT1/2 and ISG expression allows for robust HIV-1 replication.

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Tables

Table 2-1. Primer Sequences

Primer Name Sequence

P1 5’- TTGAGTCACACCCCCAGAAT -3’ P2 5’- GAGCACATTGCTTCCAGACA -3’ P5 5’- GATGCGGCACTCGATCTC -3’ P6 5’- GCAACCTCCCCTTCTACGAG -3’ 5'- ATGCCACGTAAGCGAAACTCTGTACTGGGTCTCTCTG - ULF2 3' Alu1 5'- TCCCAGCTACTGGGGAGGCTGAGG -3' Alu2 5’- GCCTCCCAAAGTGCTGGGATTACAG -3’ Lambda T2 5'- ATGCCACGTAAGCGAAACTCTGT -3' UR2 5’- CTGAGGGATCTCTAGTTACC -3’ FAM-Int_HIV-IABkFQ 5’- CACTCAAGGCAAGCTTTATTGAGGC -3’

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Table 2-2. HIV-1-Induced Gene Expression Fold-Change Gene Symbol (HIV vs. Untreated)

CCL7 6.06824 IFI44L 5.89781 IFIT1 5.53147 CXCL11 5.01958 RSAD2 4.75459 IFIT2 4.64842 CCL8 4.58559 TNFSF10 4.42962 CXCL10 4.01804 APOBEC3A_B 3.74453 IFITM1 3.33806 USP18 2.98445 IFI44 2.73129 SIGLEC1 2.51668 IFITM2 2.48532 OAS3 2.4681 GMPR 2.46577 EPSTI1 2.46526 SERPING1 2.40793 HERC5 2.38334 SP110 2.34157 HERC6 2.30317 DDX60 2.28238 OAS2 2.25047 IFI35 2.2397 MX1 2.22373 MX2 2.20131 TLR3 2.17625 MFAP5 2.16209 NT5C3A 2.14975 IFIT3 2.14181 JUP 2.13316 IFI27 2.04006 FBXO6 2.02068

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CHAPTER 3 HARMINE BOOSTS REACTIVATION OF LATENT HIV-1 BY LATENCY REVERSING AGENTS

Introduction

HIV-1 infection remains incurable because of the ability of this virus to permanently integrate its genetic material into the host genome of infected cells.

Integration is not a key feature of all latent viruses. For example, herpes viruses maintain latency as a non-integrated viral DNA episome [502, 503]. However, the ssRNA genome of HIV-1 requires a stable intermediate in the form of integrated proviral

DNA to complete its life cycle. This integrated DNA is subject to the same regulatory

mechanisms as host genes including requirements for active transcription factors as

well as epigenetic control of chromatin structure and accessibility [504].

Like host genes, the HIV-1 genome may enter a state of non-expression wherein

viral mRNA and proteins are not expressed [505]. Whether this non-expression is

through neglect (lack of stimuli required to drive the HIV-1 LTR promoter) or active

suppression (binding of inhibitory proteins to the LTR or epigenetic silencing of the

locus), the outcome is the same – HIV-1 is not produced and the virus remains hidden

from host immunity. In this state, HIV-1 endures as long as the cell that it infects.

Reactivation of latent HIV-1 requires availability of transcription factors, reversal of

epigenetic silencing, and active transcriptional machinery.

P-TEFb and HIV Tat

During the first rounds of early transcription, basal levels of transcription factors

such as NFκB and Sp1 bind to the HIV-1 LTR promoter, which promotes initiation of

transcription by RNA Pol II. However, due to the activity of negative elongation factor

(NELF) and DRB-sensitive inducing factor (DSIF), which are associated with RNA Pol II,

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transcription is paused [506–508]. Approximately 87% of the transcripts that are initiated will terminate prematurely between positions +55 to +59 [84]. However, a small number of transcripts will reach full length allowing for the translation of HIV-1-encoded Tat.

HIV-1 Tat binds to the transactivation-responsive element (TAR) on the nascent

HIV-1 mRNA, which recruits and interacts with the P-TEFb complex comprised of

CyclinT1 and Cdk9 [85, 88, 89, 248, 509]. After recruitment by Tat, Cdk9 in the P-TEFb complex phosphorylates NELF and DSIF, which causes disassociation of NELF with

RNA Pol II and DSIF to become a positive elongation factor that stays associated with

RNA Pol II [508, 510, 511]. The C-terminal domain (CTD) of RNA Pol II is also phosphorylated at serine-2, which promotes processivity of the enzyme allowing elongation of transcript [512]. Thus, when Tat levels are high approximately 99% of the transcripts reach full length [84].

Control of P-TEFb is accomplished by regulated expression of its subunits and its association with different inhibitory complexes that determine the specific activity (or inactivity) of P-TEFb. A 7SK snRNP complex containing HEXIM1 and a 7SK snRNA binds to P-TEFb inhibiting the kinase activity of CDK9 [513–516]. A key role for HIV-1

Tat protein is to release CDK9/CycT1 P-TEFb from 7SK snRNA so that it may drive

HIV-1 mRNA transcription [517]. Alternatively, CDK9/CykT1 can be bound by Brd4 and this drives expression of host genes such as c-Myc [518–521]. However, Brd4 competes with Tat for P-TEFb [522]. It has been shown that the Brd4 inhibitor JQ1 enhances tat-induced HIV transcription and reversal of latency [523, 524]. Thus, the P-

TEFb transcription complex acts as a three-position switch relative to HIV and host gene mRNA expression. HEXIM-containing complexes are in the off position for all P-

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TEFb-dependent genes. Brd4-containing complexes are in the off position for HIV transcription but the on position for host genes such as c-Myc. Finally, tat-containing complexes are in the on position for HIV transcription.

HIV-1 transcription factors

The LTR promoter of HIV-1 has binding sites for the general transcription factors such as a TATA sequence for binding of TFIID and associated TAFs and a highly active start site region for directing RNA Pol II binding [525–527]. The HIV-1 core promoter also contains three tandem Sp1 binding sites [528]. The HIV-1 LTR also contains an enhancer region with two binding sites for NFκB to which NFκB or NFAT may bind [240,

245, 529]. Since the NFκB and NFAT binding sequences overlap with each other, NFκB and NFAT compete for binding to these sites [530–532]. Binding of Sp1 to the Sp1 binding sites enhances the binding of NFκB to the NFκB binding sites [533].

Chen and colleagues demonstrated that mutation of the NFκB binding sites does not result in a significant reduction in HIV-1 replication in transformed cell lines [534].

However, signaling through the viral enhancer by NFκB or NFAT is required in primary

T cells for transcription as well as reactivation of HIV-1 from latency [535, 536]. These

TFs are normally sequestered in the cytoplasm as inactive proteins. Numerous early studies of HIV latency have focused on treatments that stimulate T cell activation including cytokines (IL-2 and IL-7), ligation of surface proteins (PHA, anti-CD3) or chemical stimulators of signaling pathways (PKC agonists) [537–545]. NFκB, in particular, is a potent inducer of HIV mRNA expression [546]. In vitro stimulation with

TNF was able to reactivate latent HIV-1 in cells by activating NFκB [243, 477, 505].

Several PKC agonists have been validated for their ability to elicit reactivation of latent

HIV. These treatments are even more potent when paired with complementary drugs

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that target HDACs or Brd4 [253, 254]. Other transcription factors such as STATs and

IRFs play roles in regulating the HIV-1 LTR [547].

We became interested in the NFAT family of transcription factors because of the presence of NFAT TF binding sites in the HIV LTR and evidence that NFAT can enhance HIV mRNA expression. NFAT, like NFκB, is sequestered as an inactive protein in the cytoplasm unless activated by specific upstream signals. However, unlike NFκB,

NFAT is hyperphosphorylated in its inactive state [548]. This hyperphosphorylation is driven by a dual specificity kinase DYRK1A [549, 550] and NFAT only becomes de- phosphorylated following calcium-dependent calmodulin signaling [548].

DYRK1A inhibitors

Harmine. Harmine (Figure 3-1) is a naturally-occurring tricyclic β-carboline alkaloid with hallucinogenic properties that is derived from the hallucinogenic plant

Banisteriopsis caapi as well as others [551]. Harmine has been shown to be an inhibitor of DYRK1A [552–554]. Harmine’s primary target is monoamine oxidase A (MAO-A)

[555–558], but a well-reported role for inhibition of DYRK1A leading to enhanced NFAT activity has been reported [552–554, 559]. The crystal structure of harmine complexed with DYRK1A has been solved confirming its binding to the ATP-binding pocket of

DYRK1A [560].

INDY. Inhibitor of DYRK1A (INDY) was discovered through a screen of chemical library to identify inhibitors of DYRK1A [560]. It was shown through a competitive binding assay as well as through x-ray crystallography that INDY inhibits DYRK1A through competitive binding to the ATP-binding pocket of DYRK1A [560]. INDY and harmine make hydrogen bonds with the same residues (Leu241 and Glu203) [560].

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However, INDY does not inhibit MAO-A [560]tg00. This mechanism is similar to how

harmine inhibits DYRK1A, but the structures are chemically distinct (Figure 3-1).

TG003. TG003 is an inhibitor that has a very similar structure to INDY and was

first identified as an inhibitor of Cdc2-like kinase (Clk) [561, 562]. It has been shown to

also have inhibitory activity against DYRK1A, but is not as potent as INDY with INDY

being about three-fold more potent [560, 563]. TG003 and INDY both share the same

basic benzothiol structure with TG003 having a methoxy group in place of a hydroxyl

group on the benzyne ring (Figure 3-1).

Rationale and summary

In the absence of antiviral drugs, reactivation most frequently leads to resumption

of the HIV/AIDS clinical progression [41]. Elimination of the latent reservoir, or

significant reduction of the size of the reservoir, are seen as the only real hope for a

sterilizing or functional cure, respectively. A large body of work has now been focused

on pharmacologic strategies to “purge” the latent reservoir. The most widely studied

approach is known as “shock and kill” [221, 222]. The goal is to simultaneously

reactivate all (or most) replication-competent latent HIV while maintaining ART. Ideally,

upon re-expression of viral mRNA and proteins, the reservoir cells will die through either

cytopathic effects of the virus or through immune mechanisms (e.g. cytotoxic T

lymphocytes [CTL] or natural killer [NK] cells).

There has been some limited success with the HDAC inhibitor Vorinostat

(Chapter 1) for reactivating the latent reservoir. However, not all the latently infected

cells are reactivated and in vitro studies have shown that not all reactivated CD4+ T cells are killed by the cytopathic effects of the virus [564]. This presents two problems:

1) better pharmacologic methods are needed to reactivate all cells latently infected with

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replication-competent HIV-1; 2) better methods for augmented CTL killing of reactivated cells are also needed.

Because of its ability to augment NFAT signaling, we undertook a study of harmine and other DYRK1A inhibitors to determine if these compounds could enhance

HIV-1 reactivation alone or in combination with other known LRAs. Here we report that harmine and other DYRK1A inhibitors INDY, TG003 and L41 boost HIV-1 reactivation by PKC agonists, Vorinostat, and JQ1. Interestingly, the effect was independent of

NFAT activity because harmine was effective at boosting LRAs even when no evidence of NFAT activity was detected. Instead, we found by whole genome microarray that harmine modulates expression of key P-TEFb components. HEXIM expression is significantly down-regulated by harmine whereas the cyclin CCNT2 was up-regulated.

We conclude that DYRK1A inhibitors regulate P-TEFb complex heterogeneity and establish an environment that is more conducive to HIV-1 reactivation. Combination treatments with harmine and LRAs may prove efficacious in vivo.

Methods and Materials

Cell culture

J-Lat 5A8 cells have been previously described [565] and were a kind gift from

Warner Greene. 5A8 cells were cultured in RPMI 1640 with 2 mM L-glutamine (Corning) supplemented with 10% heat-inactivated FBS (Sigma-Aldrich, St. Louis, MO) and 100

IU penicillin-streptomycin.

Flow cytometry

Cells were fixed in 2% paraformaldehyde and GFP fluorescence was measured using the BD Accuri™ C6 flow cytometer (BD Biosciences).

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Reagents

Harmine, INDY, TG003, bryostatin, JQ1, SAHA, and ingenol were purchased

from Tocris Bioscience (Bristol, UK). Prostratin was purchased from Cayman Chemical

(Ann Arbor, MI). C305 was purchased from Millipore (Darmstadt, Germany). PMA and

PHA were purchased from Sigma-Aldrich. Recombinant TNF was purchased from

PeproTech (Rocky Hill, NJ). DMSO was purchased from Fisher Scientific (Suwanee,

GA). Anti-HIV-1 p24 Antibody [39/5.4A] was purchased from Abcam (Cambridge, MA) and HSP90 [C45G5] and phospho-p44/42 MAPK (ERK1-Tyr204/ERK2-Tyr187)

[D1H6G] antibodies and the MEK1/2 inhibitor, U0126, were purchased from Cell

Signaling Technology (Danvers, MA).

Western blot

Cells were washed with PBS and lysed in 1X Cell Lysis Buffer (Cell Signaling

Technology) with Protease Inhibitor Cocktail (Sigma-Aldrich) and 1 mM PMSF (Sigma-

Aldrich). Proteins were separated by size by SDS-PAGE with 4-20% Mini-PROTEAN®

TGX™ precast gels (Bio-Rad, Hercules, CA) and electroblotted onto Trans-Blot®

Turbo™ Mini PVDF membranes (Bio-Rad) using the Trans-Blot® Turbo™ Transfer

System (Bio-Rad) according to the manufacturer’s instructions. Membranes were blocked in 5% non-fat milk in TBST. The Super Signal® West Dura Extended Duration

Substrate kit was used for chemiluminescence detection of HRP-linked secondary antibodies. Blots were exposed to CL-Xposure™ Film (Thermo Scientific, Waltham,

MA). Blots were stripped by incubating with Restore™ PLUS Western Blot Stripping

Buffer (Thermo Scientific).

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RT-qPCR

Total RNA was isolated with RNeasy Plus Mini Kit (Qiagen). cDNA synthesis was

carried out with High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems,

Foster City, CA) according to the manufacturer’s instructions. RT-qPCR reactions were

carried out in SYBR™ Select Master Mix (Thermo Fisher Scientific, Waltham, MA)

according to the manufacturer’s instructions. RT-qPCR reactions were performed on the

StepOnePlus™ Real-Time PCR System (Applied Biosystems). Gag primers used were

FWD: 5’- GAGCTAGAACGATTCGCAGTTA-3’; REV: 5’-

CTGTCTGAAGGGATGGTTGTAG-3’.

Luciferase Assay

Reporter cell lines were created by transducing Jurkat T cells with Cignal Lenti

NFAT reporter (Cat# CLS-015L), Cignal Lenti NFκB reporter (Cat# CLS-013L), or

Cignal Lenti Negative Control (CLS-NCL) lentiviral particles purchased from Qiagen. An

equal volume of Bright-Glo™ Luciferase Assay System (Promega) was added to an

equal volume of cells. After 5 minutes, the cells were lysed and the lysate was

transferred to a black Costar EIA/RIA polystyrene half area 96-well plate (Corning).

Luminescence was measured with the VICTOR™ X4 Multi-Plate Reader (PerkinElmer).

Microarrays

J-Lat 5A8 cells were treated with DMSO alone, harmine alone, PMA alone, or

PMA + harmine overnight. Total RNA was collected with RNeasy Plus Mini Kit (Qiagen).

Gene expression was assessed with GeneChip™ Human Transcriptome Array 2.0

arrays (Affymetrix) by the Interdisciplinary Center for Biotechnology Research at the

University of Florida. The analysis was performed with Partek Genomics Suite v. 6.6

(Partek Inc., St. Louis, MO). CEL files were imported with GC correction and intensity

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data were transformed to log base 2. A two-way-ANOVA with contrast was performed to

determine fold changes between control and treatment groups.

Results

DYRK1A inhibitors enhance efficacy of T cell activating LRAs

DYRK1A inhibitors harmine or INDY were added to J-Lat 5A8 cells alone or in

combination with the PKC agonists PMA, the T cell activating plant lectin PHA, TNF, or

the IgM antibody C305 that crosslinks the Jurkat TCR. Both harmine and INDY failed to

cause any measurable HIV reactivation when used alone at any dose. However,

harmine and INDY boosted the reactivation effects of PMA, PHA, C305, and TNF

(Figure 3-2A-H). Harmine was more potent for PMA whereas INDY was more potent

when paired with PHA, C305, or TNF. Next, the panel of PKC agonists was expanded

to include more clinically relevant compounds prostratin, bryostatin, and ingenol. Along with PMA, each of these PKC agonists was markedly more potent when combined with harmine (Figure 3-3A-B). Bryostatin was then paired with harmine or INDY as well as two additional DYRK1A inhibitors TG003 and L41. All four DYRK1A inhibitors boosted the efficacy of bryostatin with the greatest effect coming from harmine (Figure 3-3C).

When analyzing flow cytometry data, we recognized a unique phenomenon in the

J-Lat reactivation experiments. Not only was harmine increasing the frequency of cells

that became GFP+, but it also appeared that the brightness of each GFP+ cell was also

increased when harmine was included (Figure 3-3C). J-Lat cells were then activated

with increasing doses of PMA, prostratin or bryostatin in the presence of DMSO,

harmine, INDY or TG003. When gating on only the GFP+ cells, it became clear that

harmine increases the amount of GFP expressed in each reactivated cell (Figure 3-4).

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By contrast, INDY and TG003 actually decrease the brightness of GFP+ cells compared to DMSO. This finding suggests that harmine works through a mechanism that is unique from other DYRK1A inhibitors. It also seems to indicate that the boosting effect of harmine is working through at least two biological pathways – one that increases the sensitivity of cells to activating stimuli (reduced the dose of stimulus required to activate

HIV) and a second pathway that enhances the magnitude of HIV LTR activity in cells where reactivation occurs.

J-Lat cells express GFP in place of Nef as an indicator of LTR activity. For a more direct measure of HIV-encoded genes/proteins, J-Lat cells were activated with bryostatin in the presence or absence of harmine. Here, like GFP, Gag protein was induced by bryostatin + DMSO and the expression was markedly increased with bryostatin + harmine. Harmine alone failed to induce Gag expression (Figure 3-5). Next,

RT-qPCR was performed to measure gag mRNA. For three different stimuli, bryostatin,

SAHA, and JQ1, harmine increased gag mRNA levels (Figure 3-6). We did not anticipate that harmine would enhance HIV reactivation by SAHA because our proposed mechanism (enhanced NFAT activity) would require an activating stimulus to have a meaningful effect. To investigate this further, we analyzed GFP expression in J-

Lat cells treated with SAHA in combination with DYRK1A inhibitors. Consistent with our expectations, we observed no difference in the frequency of GFP+ cells when DYRK1A inhibitors were included. However, analysis of the mean fluorescence intensity of those cells that became GFP+ showed that harmine enhanced the magnitude of GFP expression while INDY and TG003 did not (Figure 3-7). These findings indicate that harmine is mediating two distinct effects on HIV latency. One effect increases the

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sensitivity of cells to activating LRAs whereas a second effect increases the expression

level of HIV genes/proteins. This finding leads to two key questions: 1) Is DYRK1A the

target of harmine in HIV latency models?; and 2) Does harmine enhance HIV

reactivation through enhanced NFAT activity?

DYRK1A expression is required for harmine’s positive effects on HIV-1 reactivation

A J-Lat cell line lacking DYRK1A expression was derived by CRISPR/Cas9 gene

targeting (Figure 3-8). Because harmine is an inhibitor of DYRK1A, we anticipated that

J-Lat cells lacking DYRK1A would behave similarly to harmine treated cells with

increased GFP expression in response to activating LRAs. In contrast to this

expectation, we found that absence of DYRK1A expression actually reduced the frequency of GFP+ cells and the MFI of GFP in positive cells was the same as those

expressing DYRK1A (Figure 3-9). Thus, harmine either acts through a different pathway

altogether or the active site binding of harmine to DYRK1A directs the protein into a

different pathway such as ERK1/2 signaling.

There is evidence to support crosstalk between DYRK1A and MAPK in murine

models of brain disease where DYRK1A enhances ERK phosphorylation and harmine

inhibits this effect [566]. In our model, where natively expressed DYRK1A is inhibited by

harmine and ERK is activated by an extrinsic stimulus, we find the opposite – that

harmine enhances ERK phosphorylation. The difference could be due to a number of

factors including cell type, species (our study is human), overexpression versus native

expression of DYRK1A, or the presence of an LRA ERK agonist in our study. To test

this, J-Lat cells cultured with PMA +/- harmine were analyzed for phospho-ERK1/2

levels. We found that harmine markedly boosted phospho-ERK1/2 induced by PMA

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(Figure 3-10). Furthermore, the inclusion of an ERK inhibitor, U0126, resulted in dose- dependent suppression of PMA-induced GFP expression. Importantly, while the inhibitor markedly reduced the frequency of GFP+ cells, it had no effect on the MFI of

GFP+ cells (Figure 3-11). The harmine + PMA treated cells were still significantly brighter, even at the highest concentration of inhibitor tested. Thus, it appears that harmine boosts the frequency of GFP+ cells by enhancing sensitivity to activating LRAs and that a second mechanism is increasing the magnitude of LTR activity independent of a strong activating signal.

Harmine boosts HIV-1 reactivation independent of effects on NFAT

DYRK1A negatively regulates NFAT by hyperphosphorylation which excludes

NFAT from the nucleus. Only through calcium-dependent calmodulin activation is NFAT de-phosphorylated resulting in nuclear translocation and activation of NFAT dependent genes such as IL-2. Because the HIV LTR contains at least two NFAT binding motifs and because harmine is a known inhibitor of DYRK1A, we wanted to know if enhanced

NFAT activity contributes to harmine’s anti-latency effects. First, we wanted to determine if harmine affects NFAT activity in T cells. Jurkat cells were transduced with lentiviral luciferase reporter constructs for NFAT, NFκB or a negative control virus with luciferase gene but no promoter. When reporter cells were treated with PMA, prostratin, or ingenol A alone, the NFAT reporter cells failed to express luciferase, but the NFκB cells showed a strong response to PMA, prostratin, and ingenol A (Figure 3-12). This finding is meaningful because harmine does potently enhance the LRA effects of PMA and other PKC agonists. Thus, NFAT is dispensable for the boosting effects of harmine.

To confirm that the NFAT reporter cells functioned properly, the cells were treated with ionomycin, which induces a calcium flux known to result in NFAT activation. The NFAT

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reporter cells responded as expected with strong NFAT induction in response to ionomycin (Figure 3-13). All of the agonists used to reactivate latent HIV-1 in this study that were enhanced by harmine do not induce NFAT, but NFκB, suggesting that the mechanism of action of harmine is not by inhibiting DYRK1A from phosphorylating

NFAT.

Harmine downregulates HEXIM1 in PMA treated 5A8 cells

We next wanted to determine what effect harmine treatment had on gene expression in PMA activated 5A8 cells. To determine effects harmine has on the whole transcriptome, 5A8 cells were treated with DMSO, harmine, PMA, and PMA + harmine and gene expression was assessed using Affymetrix arrays (Figure 3-14). The most downregulated gene with harmine + PMA was HEXIM1, which is a protein in the inhibitory P-TEFb complex (Table 3-1). Interestingly, the most upregulated gene was

CyclinT2 (Table 3-1), which may also be a part of the P-TEFb complex. Thus, harmine boosts reactivation of latent HIV-1 by changing the expression of proteins that are part of the P-TEFb complex required for elongation of HIV-1 transcripts.

Discussion

In the current study, we demonstrate that the plant-derived compound harmine can boost the efficacy of latent HIV-1 reactivation mediated by LRAs. Our data show that harmine has two effects: 1.) it increases the number of cells that are reactivated by

LRAs, and 2.) it increases LTR promoter activity in reactivated cells. Harmine has been shown to inhibit DYRK1A, a negative regulator of NFAT signaling [552–554, 559].

However, in the current study, we demonstrate that the boosting effect is independent of

NFAT. We found that harmine treatment increases PMA-induced phosphor-ERK1/2 levels.

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Previous studies have demonstrated that PKC agonists such as prostratin, bryostatin, and ingenol can reactivate latent HIV-1 and that this mechanism is through

NFκB activation [252, 543, 567–572]. These studies have also demonstrated that reactivation of latent HIV-1 through PKC agonists is synergistic with latency reversal with HDAC inhibitors or JQ1 [252, 543, 567–572]. This synergistic effect is because two different mechanisms for latency are being targeted: availability of transcription factors and either epigenetic modifications or availability of P-TEFb complexes. In our current study, we demonstrate that the drug harmine has a synergistic effect with PKC agonists,

HDAC inhibitors, and JQ1.

A recent study by Booiman and colleagues demonstrated that using the DYRK1A inhibitor, INDY, that reactivation of latently infected J-Lat cells could occur without the use of a PKC agonist [573]. In our hands, treatment of J-Lat with DYRK1A inhibitors alone was not sufficient to reactivate latently infected cells. Treatment with DYRK1A inhibitors only had an effect on J-Lat cells that were stimulated with PKC agonists,

SAHA, or JQ1. The study by Booiman and colleagues used J-Lat clones 8.4 and A1, whereas we used clone 5A8 [573].

While we initially expected that the mechanism of harmine boosting HIV-1 reactivation was due to inhibiting DYRK1A and increasing the availability of activated

NFAT, we found that there was no NFAT activity when using PKC agonists. Instead, we found that NFκB activation was boosted by harmine. While it is not surprising that the mechanism of action is through NFκB for the PKC agonists used, it is surprising that harmine would boost this pathway.

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We also provide evidence that harmine may be boosting signaling through the

MEK/ERK pathway. Indeed, inhibition of MEK resulted in abrogation of the boosting effect of harmine with PMA stimulation. When used in conjunction with PKC agonists our study demonstrated that DYRK1A inhibitor harmine boosted the response differently than DYRK1A inhibitors INDY and TG003. While all DYRK1A inhibitors boosted the frequency of reactivated cells after treatment with PKC agonists, only harmine boosted the MFI of the reactivated cells. This suggests that harmine may be having a secondary effect on DYRK1A or another unidentified cellular target.

Microarray data demonstrated that with harmine treatment that levels of HEXIM were reduced. HEXIM, when part of the P-TEFb complex, is inhibitory for HIV-1 replication [515–517]. This would suggest that the mechanism of boosting by harmine is through increasing the availability of active P-TEFb complexes for recruitment by Tat.

The microarray also revealed increased expression of CyclinT2. While CyclinT1 is the kinase normally associated with the PTEF-b complex CyclinT2 has also been reported to associate with CDK9 in the PTEF-b complex. Unlike CyclinT1, which promotes Tat activity, CyclinT2 has been reported to be inhibitory for HIV-1 transcription [574].

In our current study, we treated J-Lat 5A8 cells with PMA, PHA, C305, and TNF in the presence or absence of DYRK1A inhibitors harmine and INDY. In a dose- dependent manner, the percentage of GFP positive cells increased with DYRK1A inhibitors and LRAs compared to just LRAs alone. This is interesting since PMA and

TNF treatments are not expected to induce NFAT phosphorylation. If DYRK1A inhibitors were boosting reactivation due to lack of negative regulation of NFAT, then the effect should not be seen with PMA and TNF.

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In the J-Lat model, GFP acts as a surrogate measure for LTR activity. We also wanted to measure gag expression by Western blot and RT-qPCR to see if LTR promoter activity was increased. Bryostatin-induced gag expression was dramatically increased with the addition of harmine. Increased expression of viral proteins will also make the “kill” phase of the shock and kill approach more effective as it is necessary for targeting by immune cells. Signaling through the TCR activates NFκB and NFAT. It has been previously reported that TCR signaling enhances transcriptional elongation of latent HIV-1 by activating P-TEFb through an ERK-dependent pathway [546].

Interestingly, we found that treatment with PMA leads to ERK-1/2 phosphorylation and that this effect was enhanced by treatment with harmine. This may be the mechanism by which harmine treatment enhances LTR promoter activity through P-TEFb.

Thus, harmine increases PMA induced ERK1/2 signaling which leads to downregulation of HEXIM1 and increased transcriptional elongation through P-TEFb.

Further, harmine also enhances the effects of PKC agonist HIV-1 reactivation through enhanced NFκB signaling and not through NFAT signaling. We propose that harmine can be used in combination with existing LRAs to increase their efficacy in vivo.

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Figures

Figure 3-1. Chemical structure of DYRK1A inhibitors.

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Figure 3-2. DYRK1A inhibitors boost LRA-induced HIV-1 reactivation in 5A8 cells. 5A8 cells were treated with DYRK1A inhibitors or DMSO and GFP fluorescence was measured by flow cytometry and the fold change calculated relative to DMSO after treatment with PMA (A-B), PHA (C-D), C305 (E-F), or TNF (G-H).

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Figure 3-3. Harmine boosts clinically-relevant LRA-induced HIV-1 reactivation in 5A8 cells. Clinically relevant LRAs prostratin, bryostatin, and ingenol were used to activate cells in the presence or absence of harmine. GFP fluorescence was measured by flow cytometry (A) and the percentage of GFP+ cells was calculated (B). 5A8 cells treated with bryostatin were treated with one of four DYRK1A inhibitors and the fold change of GFP compared to control was calculated (C).

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Figure 3-4. Harmine boosts the MFI of GFP induced by LRAs. J-Lat 5A8 cells were treated with different doses of PMA, prostratin, or bryostatin. A) Gating strategy to target GFP positive cells. B) GFP positive cells treated with 20 nM PMA. C) Dose titration of PMA. D) GFP positive cells treated with 10 μM prostratin. E) Dose titration of prostratin. F) GFP positive cells treated with 12.5 μM bryostatin. G) Dose titration of bryostatin.

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Figure 3-5. Harmine boosts bryostatin-induced gag expression in J-Lat cells. J-Lat 5A8 cells were treated with bryostatin in the presence of harmine or DMSO. (A) GFP expression representing HIV reactivation was measured by flow cytometry and (B) HIV-1 Gagp24 expression was measured by Western blot.

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Figure 3-6. Harmine boosts LRA-induced gag expression in J-Lat cells. J-Lat 5A8 cells were treated with bryostatin, SAHA, or JQ1 in the presence of harmine or DMSO. Expression of gag mRNA was measured by RT-qPCR.

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Figure 3-7. Harmine boosts the magnitude of SAHA reactivation, but not the frequency of reactivated cells. J-Lat 5A8 cells were treated with different doses of SAHA in the presence of DYRK1A inhibitors. A) The percentage of GFP positive cells was measured by flow cytometry. B) The mean fluorescence intensity was measured by flow cytometry.

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Figure 3-8. DYRK1A CRISPR knockout in 5A8 cells. J-Lat 5A8 cells were transfected with plasmids containing a DYRK1A specific sgRNA, Cas9, and a selectable marker cassette. After selection with puromycin, the expression of DYRK1A was measured by Western blot.

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Figure 3-9. DYRK1A knockout in 5A8 cells does not boost PMA-induced reactivation of latent HIV-1. J-Lat 5A8 cells with and without knockout by CRISPR/Cas9 gene editing were treated with 5 nM PMA for 24 hours and GFP expression was measured by flow cytometry.

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Figure 3-10. Harmine boosts PMA-induced phosphor-ERK1/2 expression. J-Lat 5A8 cells were treated with DMSO or harmine with or without PMA treatment. Phospho-ERK1/2 expression was measured by Western blot.

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Figure 3-11. MEK/ERK inhibitor abrogates effect of harmine in PMA treated 5A8 cells. J-Lat 5A8 cells were activated with PMA in the presence of DMSO or harmine and different concentrations of MEK/ERK inhibitor U0126. GFP fluorescence was measured by flow cytometry.

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Figure 3-12. Jurkat T cells that were transduced with lentivirus NFκB or NFAT luciferase reporters or a negative control lentivirus were treated with different doses of PKC agonists overnight. The dashed line represents no agonist treatment.

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Figure 3-13. Ionomycin induces NFAT activation in Jurkat luciferase reporter cell line. The dashed line indicates luciferase reagent with no cells added (blank).

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Figure 3-14. Heat map of gene expression in 5A8 cells treated with PMA with or without harmine. Genes that were significantly induced (p<0.001; FDR 0.05) at least 2-fold up or down in the PMA + harmine vs. PMA alone treatment are shown. Significance was calculated by two-way-ANOVA with contrast using Partek Genomics Suite 6.6. A list of genes can be found in Table 3-1.

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Tables

Table 3-1. PMA-Induced Gene Expression with Harmine Fold-Change Gene Symbol (PMA + harmine vs. PMA alone)

CCNT2 2.2142 SOX30 2.10846 TMEM2 -2.0027 RAB23 -2.03377 ANKRD50 -2.12436 USP27X -2.13768 IL27RA -2.1761 RASA2 -2.3009 SLC7A11 -2.37159 LRRC8B -2.3876 SKIL -2.4415 USP27X -2.44667 CHAC1 -2.52968 HEXIM1 -3.29431

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CHAPTER 4 CONCLUDING REMARKS

The main barrier to a cure for HIV-1 is viral reservoirs that persist for the life of

the host. For a sterilizing cure to be achieved these viral reservoirs must be eliminated.

Although existing antiretroviral therapies have been life-saving for many HIV patients, it

requires life-long treatment and places a high economic burden on the patient. Many

patients outside of the developed world do not even have access to these life-saving

medications. New therapies that eliminate persistently infected cells must be developed.

The HIV-1 field has advanced considerably in the last three decades since its

discovery. The strong social movement from patients and patient advocate groups in

the 1980s and 1990s helped push the field to develop FDA approved antiretroviral

drugs that have saved many lives since their implementation. There has also been a

considerable amount of research funds from the federal government in the United

States of America as well as in other countries that have allowed for extensive research

of HIV-1. While the basic biology of HIV-1 is very well understood there are still many aspects of HIV-1’s interactions with host cells that we still do not understand.

In recent years, clinical trials utilizing LRAs have sought to reactivate the pool of latently infected cells in patients. Although this approach has had some efficacy in vitro and in vivo, it has not been able to eliminate all infected cells. Even a few latently infected cells remaining can be reactivated naturally and allow the course of infection to resume.

One approach to solving this problem is to develop reagents that are better at reactivating T cells. However, the problem with this approach is that these reagents do not specifically activate infected cells only, but actually reactivate all resting T cells.

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Finding reagents that do this better may reactivate all latently infected cells, but may not be safe for the patient. Too much immune activation can lead to a cytokine storm that can kill the patient.

A second approach is to develop therapies that are better at targeting HIV-1 reactivation without strongly and non-specifically reactivating all T cells. To develop a therapy such as this a better understanding of how the components involved in HIV-1 transcription and that govern latency is required. In the current study, we use harmine, a naturally occurring plant compound, to boost the efficacy of existing LRAs.

Harmine was found to boost the efficacy of LRAs that stimulate HIV-1 transcription through NFκB activation. Treatment with harmine boosted the amount of activated NFκB present in the cells, which acts to tune the magnitude of reactivation.

However, there seems to be a second effect where harmine lowers the threshold for reactivation leading to not only a higher magnitude of reactivation in our cell model but also a higher frequency of reactivated cells.

Since these LRAs such as prostratin, bryostatin, and ingenol are already FDA approved drugs, it is possible that complementing these treatments with harmine may prove to be a more efficacious way to achieve latency reversal in vivo. Although harmine is a hallucinogen, there are no known damaging side effects of using this naturally-occurring compound. Our data suggest that harmine may act through the

MEK/ERK1/2 pathway and decreases the availability of HEXIM1 a negative regulator of

HIV-1 transcription. Although HEXIM1 also regulates the ability of P-TEFb to promote elongation of host genes, the subset of genes affected may be small. If this is the case,

132

this may be a safer alternative for boosting current therapies without activating all T

cells and causing a cytokine storm.

While this approach may be helpful in augmenting existing latency reversing

strategies, it is unclear whether this would have any effect on infected macrophages.

There are conflicting reports in the literature as to whether true latency of HIV-1 occurs

in macrophages during ART or if there is just low-level replication in the tissues that is

not detected by laboratory tests on serum. The goal with the “shock and kill” strategy is

that if the virus is actively replicating again in cells that were previously latently infected,

then the cells will either be killed by the cytopathic effects of the virus or they will be

susceptible to immune surveillance. However, macrophages are resistant to the

cytopathic effects of the virus. This suggests a need for a therapy that specifically

targets macrophages that allows them to either become susceptible to the cytopathic

effects of the virus, gives them the ability to intrinsically suppress the virus, or make

them more susceptible to immune surveillance.

In microarrays looking at what genes are induced by HIV-1 in macrophages,

USP18 was one of the genes induced with the highest fold change by HIV-1. USP18

negatively regulates T1 IFN signaling which may be making the cells more permissible

for HIV-1 infection. Indeed, studies have shown that HIV-1 is restricted by T1 IFNs in

vitro and in vivo. However, in vivo therapies only have efficacy for a short period of days

before the patients become refractory to IFN treatment. This is likely due to the

induction of negative regulators of IFN signaling.

Negative regulators of IFN signaling as with any signaling pathway are essential for shutting down immune responses when they are no longer needed. If this regulation

133

did not exist there would be excess inflammation which would damage and kill the host.

However, if viruses can induce these negative regulators without inducing a strong enough response to eliminate the virus, then the negative regulators may be allowing the virus to persist.

To test this, we utilized in vitro knockout models with cells that lacked USP18.

Our results showed that the cells were better able to suppress viral replication through the enhanced IFN signaling and enhanced expression of antiviral genes. Targeting

USP18 in conjunction with recombinant IFN therapies may make the therapies more efficacious. However, care must be taken that the response isn’t too strong and damaging to the host.

Targeting USP18 may also make the infected macrophages more susceptible to immune surveillance and killing by CTLs. Microarray data and qPCR data in my studies have shown that genes involved in antigen processing and presentation are expressed at higher levels when USP18 is deficient. This may mean that antigen is presented to

CTLs and co-activators such as B7-1 are expressed higher making the macrophages better at eliciting killing by CTLs. The limit of this study is that it is in vitro with macrophages alone. The next logical step would be to test this in co-culture experiments with HIV-1-specific T cells.

In summary, this study shows that a strategy of targeting negative regulators of immune signaling pathways may help to boost the efficacy of existing HIV-1 therapies.

This approach is novel in that current therapies primarily target viral proteins. However, viruses are obligate organisms that depend on host genes as much as they depend on virally encoded proteins. Many studies have focused on host proteins that negatively

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regulate HIV-1 replication. In these studies, we demonstrate that targeting host cell factors that positively regulate HIV-1 replication, such as DYRK1A and USP18 may prove to be an effective strategy. The findings here may have applications outside of the

HIV-1 field and may help to develop better therapies for a variety of infectious diseases including other viruses and bacteria. Targeting USP18 may prove to be a useful adjuvant for vaccines as well.

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BIOGRAPHICAL SKETCH

Jared Taylor first began his science career at Millersville University of

Pennsylvania doing research in the laboratory of Dr. Ryan Wagner. Here, Jared researched polyphenol oxidase in Ailanthus altissima as an inducible chemical defense against herbivory. While attending college, Jared worked several jobs including working in quality control at Turkey Hill Dairy, an ice cream and flavored drink manufacturing plant. In 2012 Jared received his Bachelor of Science degree in biology with a minor in biochemistry.

Jared matriculated to the Interdisciplinary Program in Biomedical Sciences (IDP) in the College of Medicine at the University of Florida in 2012. After laboratory rotations,

Jared joined the laboratory of Dr. Mark Wallet in 2013. Here his main research focused on host-pathogen interactions of HIV-1. During his tenure in Dr. Wallet’s lab, Jared has presented his work at several domestic and one international research conference. He co-authored a paper that was published in 2017 in the Journal of AIDS Research and

Human Retroviruses. He has also written and submitted a first-author manuscript to the

Journal of Leukocyte Biology that is pending review. Jared graduated with his Doctor of

Philosophy degree in medical Sciences with a concentration in immunology and microbiology from the University of Florida in December 2017.

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