role of kinases in kaposi’s sarcoma-associated …

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ROLE OF KINASES IN KAPOSI’S SARCOMA-ASSOCIATED HERPESVIRUS PATHOGENESIS

Jason P. Wong

A dissertation submitted to the faculty at the University of North Carolina at Chapel Hill in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department

of Microbiology & Immunology in the School of Medicine.

Chapel Hill 2019

Approved by:

Blossom A. Damania

Dirk P. Dittmer

Cary Moody

Albert S. Baldwin

Nancy Raab-Traub

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© 2019 Jason P. Wong

ALL RIGHTS RESERVED

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ABSTRACT

Jason P. Wong: Role of kinases in Kaposi’s sarcoma-associated herpesvirus pathogenesis (Under the direction of Blossom A. Damania)

Kaposi’s sarcoma-associated herpesvirus (KSHV) is associated with four diseases:

Kaposi’s sarcoma (KS), primary effusion lymphoma (PEL), multicentric Castleman’s disease

(MCD), and KSHV-associated inflammatory cytokine syndrome (KICS). KS is the most

common AIDS defining malignancy, and even though long term remission is possible, KS often

relapses and there is no evidence that suggests KS can be cured. PEL is a B-cell non-Hodgkin

lymphoma (NHL) that has a poor prognosis with a median survival time of about six months.

Therefore, there is a need to develop alternative therapies for KSHV-associated malignancies.

KSHV modulates several cellular signal transduction pathways, some of which play a role in

tumorigenesis. Kinases often control the propagation and signaling within these pathways. In this

dissertation we examine how both host (Chapter 2) and viral kinases (Chapter 3) modulate

KSHV pathogenesis, which may lead to the development of targeted therapies to treat KSHV-

associated diseases. Characterization of host kinases which have upregulated activity in PEL in

comparison to healthy B-cells and other B-cell NHLs led to the discovery that the receptor

tyrosine kinase, Tyro3 is uniquely upregulated in PEL. Tyro3 drove survival in PEL cells by

upregulating the phosphoinositide-3-kinase (PI3K) pathway. We developed an inhibitor against

Tyro3, which inhibited growth and PI3K signaling in PEL cells, and resulted in a decrease in

tumor burden in a PEL xenograft mice model. We also characterized a viral kinase encoded by

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KSHV known as viral protein kinase (vPK). Since previous work in our lab has shown vPK

upregulates protein synthesis and overexpression of vPK in mice results in an increased rate of

developing lymphomas, we were interested in characterizing possible host protein interactors of

vPK that may play a role in KSHV pathogenesis. We found that vPK binds the tumor suppressor,

protein phosphatase 6 (PPP6). We also characterized vPK’s effect on mitophagy, a selective

form of autophagy that targets mitochondria for degradation.

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In memory of my mother, and to my father and brother

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ACKNOWLEDGEMENTS

I would like to acknowledge my research mentor, Blossom Damania, for her guidance

and direction throughout my graduate studies. She gave me the freedom to pursue many ideas,

while keeping me on the right path. I am also grateful that she encouraged me to pursue many

opportunities that I would have not pursued if left to my own devices. Without her mentorship, I

would not be where I am as a scientist today.

I would also like to acknowledge my committee members: Drs. Cary Moody (chair), Dirk

Dittmer, Albert Baldwin, and Nancy Raab-Traub. I am grateful for their feedback and help

throughout my graduate studies. I would also like to thank them for finding time in their

schedules to meet a tight deadline at the end. To Dr. Bob Bourret, Dixie Flannery, and Michelle

Hightower, thank you for your help during my graduate studies. To Lorrie Crammer and Gina

Donato, thank you for the wonderful TA experience. I would also like to acknowledge the

Virology training grant (and the people who run it) for providing support and training

opportunities throughout my graduate studies.

I would also like to acknowledge the Damania lab as a whole for their help and for

providing an enjoyable environment in lab. Special thanks to Aadra Bhatt and Louise Giffin,

who made my rotation wonderful; and to my fellow graduate students (past and present) in both

the Damania and Dittmer lab as we struggled together in our studies. In addition, I would like to

thank my friends, both in lab and out of lab for their support. I also thank the people of the

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Chapel Hill Bible Church with special thanks to my friend Tyler Farnsworth and to my life

group.

I also want to thank my collaborators for without them my project would not be possible.

Special thanks to Drs. Tim Stuhlmiller, Noah Sciaky, and Gary Johnson for performing the

MIB/MS analysis; and Drs. Xiaodong Wang and Stephen Frye for developing UNC3810A and

providing expertise regarding pharmacology-related questions.

Lastly, I would like to acknowledge my family. To my father, who sacrificed a lot to

raise both my brother and I; to my brother, who is gracious in dealing with me throughout life;

and in memory of my mother, who lovingly taught me during my childhood. Finally, I would

like to thank my heavenly Father, who through his love has carried me through many dark times

in life.

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TABLE OF CONTENTS

LIST OF TABLES .......................................................................................................................... x

LIST OF FIGURES ....................................................................................................................... xi

LIST OF ABBREVIATIONS ...................................................................................................... xiii

LIST OF SYMBOLS ................................................................................................................... xxi

CHAPTER 1: REGULATION OF SIGNALING PATHWAYS BY KINASES IN THE LIFE CYCLE OF KAPOSI’S SARCOMA- ASSOCIATED HERPESVIRUS .................................................................................................... 1

Kaposi’s sarcoma-associated herpesvirus (KSHV)-associated diseases ................................. 1

KSHV life-cycle ...................................................................................................................... 2

Modulation of signal transduction by kinases ......................................................................... 4

Common cancer-associated pathways dysregulated by KSHV ............................................... 6

Dysregulation of the phosphoinositide-3-kinase (PI3K)/protein kinase B (Akt)/mammalian target of rapamycin (mTOR) pathway by KSHV .................................... 10

Autophagy .............................................................................................................................. 16

Mitophagy .............................................................................................................................. 20

Conclusions............................................................................................................................ 23

CHAPTER 2: KINOME PROFILING OF NON-HODGKIN LYMPHOMA IDENTIFIES TYRO3 AS A THERAPEUTIC TARGET IN PRIMARY EFFUSION LYMPHOMA ................................................................................... 25

Introduction ............................................................................................................................ 25

Materials and methods ........................................................................................................... 26

Results.................................................................................................................................... 40

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Discussion .............................................................................................................................. 62

CHAPTER 3: VIRAL PROTEIN KINASE (VPK) MODULATION OF HOST SIGNALING PATHWAYS ........................................................................................ 66

Introduction ............................................................................................................................ 66

Materials and methods ........................................................................................................... 70

Results.................................................................................................................................... 78

Discussion .............................................................................................................................. 91

CHAPTER 4: CONCLUSIONS AND FUTURE DIRECTIONS ................................................ 95

Summary ................................................................................................................................ 95

Upregulation of several pathways in KSHV-associated PEL by Tyro3 ................................ 97

Targeting of mitochondria during the KSHV lytic cycle ................................................. 100

Elucidating the vPK and PPP6 interaction .......................................................................... 103

Conclusions.......................................................................................................................... 104

REFERENCES ........................................................................................................................... 105

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LIST OF TABLES

Table 2.1 Primers used for RT-PCR analysis ............................................................................... 32

Table 2.2 IC50 values of UNC3810A against top kinase hits from the kinome profiling. ............................................................................................................ 53

Table 3.1 Peptide counts of proteins identified to interact with vPK using mass spectrometry. ...................................................................................................... 79

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LIST OF FIGURES

Figure 1.1 Activation of the PI3K/Akt/mTOR and MAPK/ERK Pathway in KSHV Infection. ........................................................................................................ 11

Figure 1.2 Generation of the phagophore in mitophagy. .............................................................. 21

Figure 2.1 Replicates between MIB/MS runs are consistent with each other. ..................................................................................................................................... 41

Figure 2.2 The cellular kinome is more similar within subtypes than across subtypes. ..................................................................................................................... 42

Figure 2.3 A common set of kinases are activated within NHL lymphomas compared to primary B-cells. .................................................................................... 43

Figure 2.4 Tyro3 is uniquely upregulated in PEL......................................................................... 44

Figure 2.5 Trends in expression data are similar to a previous literature report.............................................................................................................................. 46

Figure 2.6 Inhibition of Tyro3 results in attenuation of PI3K signaling. ..................................... 47

Figure 2.7 Tyro3 knockdown attenuates survival and signaling in PEL cells. ...................................................................................................................................... 49

Figure 2.8 Synthesis of UNC3810A. ............................................................................................ 50

Figure 2.9 Specificity of UNC3810A to Tyro3. ........................................................................... 52

Figure 2.10 Inhibition of Tyro3 results in a decrease of cell survival and colony formation in PEL. ....................................................................................................... 55

Figure 2.11 UNC3810A treatment of PEL cells results in cell death. .......................................... 57

Figure 2.12 UNC3810A sensitizes PEL cells to treatment with PI3K inhibitors. ............................................................................................................................. 58

Figure 2.13 Pharmacokinetic analysis of UNC3810A. ................................................................. 59

Figure 2.14 UNC3810A retards PEL tumor growth in vivo. ........................................................ 60

Figure 2.15 UNC3810A treatment does not reduce tumor burden in a non-PEL lymphoma xenograft model. ................................................................................... 61

Figure 2.16 Immunohistochemistry analysis of solid tumors from BCBL1-xenograft mice. ................................................................................................................ 64

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Figure 3.1 vPK binds and phosphorylates several subunits of the PPP6 complex. .............................................................................................................................. 80

Figure 3.2 Examining the role of PPP6C in multiple pathways and cellular processes relevant in the KSHV life cycle. ...................................................................... 83

Figure 3.3 Ingenuity pathway analysis (IPA) of host interacting partners of vPK identified using mass spectrometry. ................................................................... 86

Figure 3.4 Autophagic markers are induced but not the integrated stress response (IRS) in vPK-HUVECs. ....................................................................................... 88

Figure 3.5 vPK expression promotes mitophagy. ......................................................................... 90

Figure 4.1 Unresolved questions regarding the role of Tyro3 in PEL. ......................................... 98

Figure 4.2 Potential questions regarding the role of vPK-induced mitophagy in the KSHV life cycle. ............................................................................................. 101

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LIST OF ABBREVIATIONS

4EBP1 Eukaryotic translation initiation factor 4E-binding protein 1

ABL Abelson murine leukemia viral oncogene homolog

ACSR AIDS and Cancer Specimen Resource

AIDS Acquired immunodeficiency syndrome

AMPK AMP activated protein kinase

ANKRD28 Ankyrin repeat domain 28



ANOVA Analysis of variance

AP-1 Activator protein 1

ART Antiretroviral therapy

ATF4 Activating transcription factor 4

ATG Autophagy related)

ATP Adenosine triphosphate

AURKA Aurora kinase A

BCR B-cell receptor

BCR Breakpoint cluster region (BCR)

BL Burkitt’s lymphoma

BNIP3 BCL2/Adenovirus E1B 19 kDA interacting protein 3

BTK Bruton’s tyrosine kinase

CCCP Carbonyl cyanide 3-chlorophenylhydrazone

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CDK Cyclin-dependent kinase

CFA Colony formation assays

CHOP C/EBP homologous protein

CI Combination index

CK2 Casein kinase II

CML Chronic myeloid leukemia

Co-IP Co-immunoprecipitation

CPCSEA Committee for the Purpose of Control and Supervision of Experiments on Animals (CPCSEA)

CTG CellTiter-Glo

CXCR2 C-X-C chemokine receptor type 2

DAB 3,3′-diaminobenzidine

DC-SIGN Dendritic cell-specific intercellular adhesion molecule-3-grabbing non-integrin

DDR DNA damage response

DE Delayed early

DHX9 DExH-box helicase 9

DIPEA Diisopropylethylamine

DLBCL Diffuse large B-cell lymphoma

DMSO Dimethyl sulfoxide

DNA-PK DNA-protein kinase

E. coli Escherichia coli

EBV Epstein-Barr virus

EIF2 Eukaryotic initiation factor 2

eIF2α eukaryotic initiation factor 2 subunit alpha

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eIF4B eukaryotic translation initiation factor 4B

eIF4E eukaryotic translation initiation factor 4E

ER Endoplasmic reticulum

ERK Extracellular-signal-regulated kinase

EV Empty vector

FL Follicular lymphoma

FUNDC1 Fun14 domain containing 1

GAPDH Glyceraldehyde 3-phosphate dehydrogenase

GEF Guanine nucleotide exchange factor

GFP Green fluorescent protein

GRK2 G protein-coupled receptor kinase 2

GST Glutathione S-transferase

HBV Hepatitis B virus

HCV Hepatitis C virus

HDAC Histone deacetylase

HHV8 Human herpesvirus 8

HIV Human immunodeficiency virus

HTLV Human T-lymphotropic virus

HUVEC Human umbilical vein endothelial cell

I.P. Intraperitoneal

I.V. Intravenous

IACUC Institutional Animal Care and Use Committee

IAEC Institutional Animal Ethics Committee

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IE Immediate-early

IFN Interferon

IFNAR Type 1 interferon receptor

IHC Immunohistochemistry

IKK IκB kinase

IL Interleukin

IMDM Iscove’s modified Dulbecco’s medium

IPA Ingenuity Pathway Analysis

IRAK1 Interleukin 1 receptor associated kinase 1

IRF3 Interferon regulatory factor 3

IRIS Immune reconstitution inflammatory syndrome

ISR Integrated stress response

JAK Janus kinase

JNK c-jun N-terminal kinase

KICS KSHV-associated inflammatory cytokine syndrome

KS Kaposi’s sarcoma

KSHV Kaposi’s sarcoma associated herpesvirus

LANA Latency-associated nuclear antigen

LC Liquid chromatography

LC3B Isoform B of LC3

LIR LC3 interacting region

LMP2A Latent membrane protein 2A

LRPPRC Leucine rich pentatricopeptide repeat containing

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MAPK Mitogen-activated protein kinase

MAPKK MAPK kinase

MAPKKK MAPKK kinase

MAVS Mitochondrial antiviral signaling protein

MCD Multicentric Castleman’s disease

MCL Mantle cell lymphoma

MEK Mitogen-activated protein kinase kinase

MerTK MER proto-oncogene tyrosine kinase

Met Methionine

MIB Multiplexed kinase inhibitor-conjugated bead

miRNA MicroRNA

MK2 MAPK activated protein kinase 2

MPP Matrix processing peptidase

MS Mass spectrometry

MTCO2 Mitochondrially encoded cytochrome c oxidase II

mTOR Mammalian target of rapamycin

mTORC1 mTOR complex 1

MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl-2-(4-sulfophenyl)-2H-tetrazolium)

MWF Monday/Wednesday/Friday

NBR1 Neighbor of BRCA1 gene

NDP52 Nuclear domain 10 protein 52

NF-κB Nuclear factor kappa-light-chain-enhancer of activated B-cells

NHL Non-Hodgkin lymphoma

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NIK NF-κB-inducing kinase

NIX Nip3-like protein X

NKT NK T-cells

NLS Nuclear localization signal

NSG Nod scid gamma

NTC Non-targeting control

NUSAP1 Nucleolar and spindle associated protein 1

OMM Outer mitochondrial membrane

Optn Optineurin

ORF Open reading frame

PARL PINK1/PGAM5-associated rhomboid-like protease

PARP Poly (ADP-ribose) polymerase

PAS Phagophore assembly site

PCR Polymerase chain reaction

PDK1 Phosphoinositide dependent kinase 1

PE Phosphatidylethanolamine

PEL Primary effusion lymphoma

PERK PKR-like ER kinase

PGAM5 Phosphoglycerate mutase 5

PI3K Phosphoinositide-3-kinase

PIC Pre-initiation complex

PINK1 PTEN-induced kinase 1

PIP3 Phosphatidylinositol-3,4,5-trisphosphate

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PK Pharmacokinetic

PKCδ Protein kinase C-delta

PLC Phospholipase C

pNPP p-nitrophenyl phosphate

PP2A Protein phosphatase 2A

PP4 Protein phosphatase 4

PPP6 Protein phosphatase 6

PPP6C PPP6 catalytic subunit

PPP6R1 PPP6 regulatory subunit 1



RIG-I Retinoic acid-inducible gene 1

RSK p90 S6 kinase

RT Real-time

RTA Replication and transcription activator

RTK Receptor tyrosine kinase

S6KB1 p70 S6 kinase

shRNA Short-hairpin RNA

STAT Signal transducer and activator of transcription

STING Stimulator of interferon genes

TAK1 Transforming growth factor beta-activated kinase 1

TAM Tyro3/Axl/MerTK

TAX1BP1 TAX1 binding protein

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TBK1 Tank binding kinase 1

Tbkbp1 TBK1 binding protein 1

TEM Transmission electron microscopy

TRIM28 Tripartite motif-containing protein 28

tRNA Transfer RNA

TSC2 Tuberous sclerosis complex 2

uORF Short upstream open reading frame

UTR Untranslated region

vBCL-2 Viral homolog of B-cell lymphoma 2

vCyclin Viral cyclin protein

VEGF Vascular endothelial growth factor

vFLIP Viral Fas-associated death domain-like interleukin-1β-converting enzyme-like inhibitory protein

vGPCR Viral G protein-coupled receptor

vIL-6 Viral interleukin 6

vIRF1 Viral interferon regulatory factor 1

vPK Viral protein kinase

WNV West Nile virus

WT Wild-type

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LIST OF SYMBOLS

α Alpha

β Beta

δ Delta

γ Gamma

κ Kappa

µ Mu

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CHAPTER 1: REGULATION OF SIGNALING PATHWAYS BY KINASES IN THE

LIFE CYCLE OF KAPOSI’S SARCOMA-ASSOCIATED HERPESVIRUS1

Kaposi’s sarcoma-associated herpesvirus (KSHV)-associated diseases

Kaposi’s sarcoma-associated herpesvirus (KSHV), also known as human herpesvirus 8

(HHV8), is the etiological agent of three cancers: Kaposi’s sarcoma (KS), primary effusion

lymphoma (PEL), and multicentric Castleman’s disease (MCD) (1). KS is the most common

cancer associated with KSHV, and is the most common malignancy in men in some Sub-Saharan

African countries. KS tumors are of endothelial origin and endothelial cells within the tumor are

infected with KSHV (2-4). There are four different types of KS that are characterized by the

patients who present the disease (1). Classical KS occurs in elderly men of Mediterranean and

eastern European descent, while African endemic KS presents itself in children and middle-aged

adults from sub-Saharan Africa. The other two classes, AIDS-KS and iatrogenic/post-transplant

KS, occur in the context of immunosuppression from either viral infection (HIV) or drugs (to

prevent allograft rejection). The two other KSHV-associated cancers, PEL and MCD, are of B-

cell origin. PEL is a clonal lymphomatous effusion generally present in the body cavities, while

MCD is a polyclonal B-cell hyperplasia characterized by systemic inflammatory symptoms (1,

5). PEL and MCD often exhibit rapid disease progression, and PEL patients have a median

survival time of six months post-diagnosis (6). A recently characterized KSHV-linked disease is

1A large part of this chapter previously appeared as an article in the journal Biological Chemistry. The original citation is as follows: Wong, J and Damania, B. Copyright © Biological Chemistry. 2017 Jul 26; 398(8):911-918. doi: 10.1515/hsz-2017-0101.

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Kaposi’s sarcoma-associated herpesvirus inflammatory cytokine syndrome (KICS), which has

symptoms similar to MCD (1). Patients have systemic inflammation characterized by high viral

titers, viral interleukin 6 (vIL-6), human IL-6, and human IL-10, but lack the lymphadenopathy

seen in MCD patients. KSHV-associated diseases generally occur in the context of immune

deficiency such as in AIDS, transplant, or elderly patients. Demonstrating the role the immune

system plays in KSHV malignancies, immune reconstitution due to antiretroviral therapy (ART)

in AIDS patients often result in clinical regression of AIDS-KS (1, 7). However, paradoxically

and in rare cases, the pro-inflammatory environment generated following ART initiation known

as immune reconstitution inflammatory syndrome (IRIS) can lead to the development of KS,

termed IRIS-KS (7). Current treatment regimens of all of these diseases consist of some form of

chemotherapy, radiation, and/or immunotherapy.

KSHV life-cycle

Like all herpesviruses, infection with KSHV is life-long. Transmission of the virus often

occurs through bodily fluids such as saliva and blood (1). KSHV infects a plethora of cell types

ranging from monocytes, fibroblasts, endothelial, epithelial, dendritic, and B-cells (8-10). Viral

entry is mediated by binding of different KSHV glycoproteins to a variety of host receptors. The

ubiquitous cell surface molecule heparan sulfate aids in the binding of KSHV to multiple cell

types (8, 9). There are several entry receptors that KSHV uses and these differ based on cell type.

In endothelial cells, the ephrin receptor tyrosine kinase A2, the transmembrane

cysteine/glutamine exchange transporter protein xCT, and several integrins (α3β1, αVβ3, αVβ5)

are important for KSHV entry (8). In macrophages, dendritic, and B-cells, KSHV binds the

dendritic cell-specific intercellular adhesion molecule-3-grabbing non-integrin (DC-SIGN)

instead of heparan sulfate to mediate entry (8, 9). Upon binding, the virus predominantly enters

the cell by endocytosis (8-10). The envelope of the virion fuses with the endocytic vesicle to

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release the capsid and tegument layer. The viral capsid is then transported along microtubules

and is delivered into the nucleus. Once in the nucleus, KSHV can enter into either the lytic or

latent lifecycle. KSHV infection is primarily latent with spontaneous bouts of lytic replication. In

KS tumor biopsies, ~99% of KSHV positive tumor cells remain latent (1). In contrast, MCD

biopsies display the highest degree of lytic replication.

No virions are produced during latency; instead KSHV persists within host cells utilizing

viral proteins to pass its genome to daughter cells following cell division. Only a few viral genes

are expressed during latency to prevent detection by the host immune system. The latency locus

encodes the latent proteins, ORF71 to ORF73, the kaposins, and twelve microRNAs (miRNAs)

(2, 11). During latency, lytic gene expression is suppressed; however, several proteins highly

expressed in the lytic cycle are present at low levels during latency such as K1 and vIL-6 (12,

13). The viral genome is tethered as an episome to the host chromosome by ORF73/LANA

(latency-associated nuclear antigen) and is replicated in coordination with the host genome using

host replication machinery (2). LANA also binds multiple lytic promoters, such as ORF50, to

suppress lytic gene expression. Many of the latent proteins upregulate cell proliferation and

survival to promote persistence of the infected cell. For instance, ORF71 can bind and activate

the IκB kinase (IKK), thereby activating the nuclear factor kappa-light-chain-enhancer of

activated B-cells (NF-κB) pathway to upregulate cell survival genes (2). Another latent gene,

ORF72/vCyclin is a homolog of cellular cyclin D and can bind to cyclin-dependent kinase 6

(CDK6) to phosphorylate retinoblastoma (Rb) and other targets to drive cell cycle progression

and proliferation (2). Many of the latent genes are expressed in KSHV malignancies. Indicative

of the role latent genes have in KSHV tumorigenesis, expression of the latency locus in B-cells

in C57BL/6 mice led to a significant increase of lymphomas and hyperplasia in mice (11).

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Although latency is the dominant program in the KSHV life cycle, spontaneous lytic

reactivation does take place (10). While the physiological stimuli that reactivates KSHV from

latency is not known, environmental stresses, histone deacetylase (HDAC) inhibitors, or phorbol

esters can cause KSHV to reactivate in vitro (1, 2). Expression of genes during the lytic phase is

separated into three groups: the immediate-early (IE), delayed early (DE), and late (2). During

the lytic phase, KSHV replicates its genome and produces infectious progeny to spread to other

cells. After viral progeny are produced the infected cell subsequently dies (1). Proteins encoded

during both the latent and lytic phases of the lifecycle are thought to play a role in KSHV-

associated diseases. Highlighting the importance of lytic gene expression, use of a viral

replication inhibitor, ganciclovir reduced the risk of KS by at least 75% in AIDS patients (14). In

a group of KSHV-MCD patients, treatment with zidovudine and valganciclovir resulted in a

greater than 80% clinical response (5). Zidovudine and ganciclovir (the active form of

valganciclovir) are phosphorylated into their toxic moieties by the lytic kinases, ORF21 and

ORF36, respectively. It is proposed that lytic cells can exhibit a paracrine effect on neighboring

cells by secreting inflammatory or angiogenic factors that are highly prevalent in KSHV-

associated diseases. For instance, endothelial cells treated with media harvested from viral G

protein-coupled receptor (vGPCR)/ORF74 expressing cells had activated extracellular signal-

regulated kinases 1/2 (ERK1/2), which is a kinase upregulated in KSHV malignancies (15).

Modulation of signal transduction by kinases

The ability of a cell to respond to different internal and external environmental cues is

important to the development, growth, survival, and/or function of the cell. In order to properly

respond to these environmental cues, receptors in the cell sense and then trigger a signaling

cascade upon binding to their appropriate ligand. These signaling cascades then promote a

cellular response that determines the cell’s fate. Examples of different environmental cues

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include infection by a pathogen, presence or absence of energy sources or nutrients, and crosstalk

between different cells. Improper responses either by the activation or inactivation of these

signaling cascades can result in localized or gross pathology of the cell or organism, respectively.

Therefore, signaling cascades tend to be tightly controlled to prevent improper signaling. Signal

transduction within the cell can typically be simplified into three steps: (1) recognition of an

environmental cue by the cell, (2) the initiation and amplification of a signal, and (3) a resulting

‘activated’ factor that then turns on, off, or modifies cellular processes (typically by gene

expression) (16). One of the major role players in the second step are proteins called kinases,

which are enzymes that catalyze the transfer of a phosphate group from ATP to a substrate (17).

The addition of the phosphate group to a substrate can alter its activity or functionality, which

then results in further transmission of the signaling cascade. Since kinases are enzymes, they are

able to catalyze many reactions over a short period of time, thereby amplifying the initial signal

(18). This allows the cell to quickly respond to environmental cues. The activity of a kinase is

typically regulated by its phosphorylation and its ability to access and bind its substrate. As

kinases are integral parts in signaling cascades, many kinases in cancer cells have dysregulated

activity (19). In a sequencing study of multiple cancers (which include breast, lung, colorectal,

gastric, melanoma, etc.) containing 210 samples, there were approximately 119 kinase-encoding

genes containing a driver mutation in 66 of the samples (19). Therefore, while not all cancers

contain a driver mutation within a kinase, a sizable portion of them may harbor a dysregulated

kinase that drives tumorigenesis. For example, the Philadelphia chromosome, which is present in

almost all patients with chronic myeloid leukemia (CML), results in the fusion between the

breakpoint cluster region (BCR) gene and the Abelson murine leukemia viral oncogene homolog

(ABL) gene (20). This translocation results in a fusion protein (BCR-ABL) that has constitutive

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kinase activity. Treatment of CML patients with imatinib, an ATP competitive inhibitor of the

BCR-ABL kinase, resulted in major and/or complete responses in a large portion of patients (20).

As seen in this example, targeting kinases as a therapeutic target in cancer is an attractive option.

Indeed, kinase inhibitors are the fastest growing class of FDA approved oncology drugs (21).

Common cancer-associated pathways dysregulated by KSHV

A common set of signaling pathways are frequently dysregulated in cancer. Many of

these dysregulated pathways contribute to cellular processes that are considered ‘hallmarks of

cancer’ including sustained proliferation, resistance to cell death, and evasion of immune cells

(22). A certain characteristic of herpesvirus infection is the establishment of life-long infection

within the host. Therefore, many of the proteins encoded by KSHV facilitate viral persistence

and replication. In accordance with this goal, the virus aims to keep its host cell alive. Therefore,

many pro-survival and proliferative pathways are upregulated during the KSHV life cycle, which

include the NF-κB, Janus kinase (JAK)/signal transducer and activator of transcription (STAT),

mitogen-activated protein kinase (MAPK), and the phosphoinositide 3-kinase (PI3K)/ protein

kinase B (Akt)/mammalian target of rapamycin (mTOR) pathways (6, 12, 23, 24). In the

following paragraphs, we will briefly review each of these pathways (with a more thorough

review on the PI3K/Akt/mTOR pathway), and how KSHV regulates the host cellular kinases

within them to alter their signaling.

In humans, the NF-κB pathway involves a set of transcription factors (p50, p52, p65,

RelB, and c-Rel) that bind the κB enhancer in DNA to induce target gene expression in response

to injury or stress (25, 26). NF-κB signaling is split into two pathways: the classical and non-

canonical pathway. In the classical pathway, NF-κB is bound to IκB during unstimulated

conditions, which masks the nuclear localization signal (NLS) of NF-κB, thereby sequestering

NF-κB to the cytoplasm (26, 27). Upon stimulation (such as by cytokines), the IκB kinase (IKK)

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phosphorylates IκB, which results in the eventual proteasomal degradation of IκB, and

subsequent translocation of NF-κB to the nucleus. The NF-κB dimer then binds to its target

DNA sequences, thereby inducing gene transcription of its target genes, which have roles in cell

survival, proliferation, and inflammation (26, 28). In the non-canonical pathway, the unprocessed

form of p52 (known as p100) is bound to RelB in an inactive complex within the cytoplasm (29).

Stimulation results in the activation of NF-κB-inducing kinase (NIK), which further activates

IKKα. IKKα then phosphorylates p100, which results in the ubiquitination and proteasomal

processing of p100 to p52. The p52/RelB complexes can then translocate to the nucleus to

activate its target genes, which affect processes including cell development and the activation of

several immune cells (29). NF-κB is frequently dysregulated in cancers as it can induce

proliferation, prevent apoptosis, induce angiogenesis, and regulate cellular metabolism (30).

Highlighting the importance of the NF-κB pathway in KSHV-associated diseases, PEL cells

require activated NF-κB for survival (31-34). Two KSHV proteins, K13 and K15, can bind and

activate IKK and NIK, respectively. K13, also known as viral Fas-associated death domain-like

interleukin-1β-converting enzyme-like inhibitory protein (vFLIP), is able to bind the gamma

subunit of IKK (IKKγ – also known as NEMO) to activate classical NF-κB signaling (35-37).

On the other hand, K15 can activate non-canonical NF-κB signaling by recruiting NIK and IKKα

to its cytoplasmic tail (38). NF-κB activation by K15 is NIK-dependent as knockdown of NIK

resulted in ablation of NF-κB activation (38).

Another pathway that responds to cytokine and growth factors is the Janus kinase

(JAK)/signal transducer of activators of transcription (STAT) pathway. This pathway consists of

four JAK family members (JAK1, JAK2, JAK3, and TYK2) and seven STAT family members

(STAT1, STAT2, STAT3, STAT4, STAT5a, STAT5b, and STAT6) (39). Signaling in this

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pathway consists of three players, the aforementioned JAK and STAT proteins, and a receptor

that recognizes ligands in the extracellular environment (such as cytokines and growth factors)

(39). Since the different JAKs are associated with certain receptors, each family member tends to

respond to certain cytokines and growth factors (39). Signal transduction in this pathway begins

when a receptor engages its ligand (39). The receptor-bound JAK becomes activated, which

phosphorylates a neighboring JAK (i.e. transphosphorylation) and the cytoplasmic tail of the

receptor. This allows for the recruitment of a STAT to the complex, which recognizes and binds

the phosphorylated cytoplasmic tail of the receptor. The JAKs subsequently phosphorylate the

STATs, which result in their dimerization and translocation to the nucleus, where they can act as

a transcription factor (39). As the JAK/STAT pathway regulates proliferation, survival, and

inflammation, it is frequently dysregulated in many cancers (40). Latent KSHV infection of

endothelial cells results in constitutive activation of STAT3; and in PEL cells STAT3 and

STAT6 are constitutively active (41-44). Part of this activity is mediated by kaposin B, which is

able to bind and activate MAPK activated protein kinase 2 (MK2), which inhibits the tripartite

motif-containing protein 28 (TRIM28), a suppressor of STAT3 (44, 45). On the other hand,

KSHV seems to inhibit STAT signaling in the context of interferon induction (46, 47). The RIF

protein encoded by open reading frame 10 is able to bind JAK1 and TYK2-associated type 1

interferon receptor (also known as IFNAR) and prevent the activation of STAT1 and STAT2

upon interferon alpha (IFN-α) treatment. Furthermore, several KSHV-encoded miRNAs target

kinases (such as PKCδ and IRAK1) that can phosphorylate STAT3 (47). Nevertheless, treatment

of PEL cells with STAT3 or STAT6 inhibitors resulted in inhibition of cell proliferation and

survival (42, 43). These findings show that regulation of JAK/STAT signaling during the KSHV

9

life cycle is complex and may be dependent on the type of ligand- receptor interaction that is

activated.

Another pathway that is activated by ligands such as growth factors and chemokines is

the mitogen-activated protein kinase (MAPK) pathways. There are four canonical MAPKs: the

p38 MAPK, c-jun N-terminal kinase (JNK), ERK1/2, and ERK5 kinases (48, 49). The canonical

MAPK pathways transduce signals by a similar mechanism. A stimulus leads to the activation of

a MAPKK kinase (MAPKKK) (50). The activated MAPKKK then phosphorylates and activates

a MAPK kinase (MAPKK), which subsequently phosphorylates and activates its MAPK. One of

the most common signaling pathways dysregulated in cancer is the MAPK/ERK pathway (51).

In the MAPK/ERK pathway, there are three MAPKKKs from the Raf family, two MAPKKs,

MEK1 and MEK2, and two MAPKs, ERK1 and ERK2 which propagate the signaling cascade.

The activation of the MAPK/ERK pathway promotes cell growth and proliferation. ERK1/2

achieves this by activating the p90 S6 kinase (RSK) leading to the phosphorylation of S6 and

eIF4B, or by promoting the activation of the transcription factor activator protein 1 (AP-1) (50,

52). The signaling cascade starts by the binding of growth factors and chemokines to their

respective receptors. Some of these receptors activate Ras, which then recruits Raf to the cell

membrane to initiate the signaling cascade. The MAPK pathway is difficult to target in cancer

due to feedback within itself and crosstalk with other survival signaling pathways such as the

PI3K pathway (50, 53). In KSHV infection and in PEL cells, the MAPK/ERK, JNK, and p38

MAPK pathways are activated (Figure 1.1) (24, 48, 49, 52, 54, 55). KSHV-encoded ORF45 can

bind to RSK, thereby stabilizing the association of RSK with ERK. This complex maintains the

activation of the MAPK/ERK pathway by preventing the dephosphorylation of RSK and ERK

(52). A virally-encoded kinase ORF36, also known as viral protein kinase (vPK), is also able to

10

phosphorylate and activate MKK4 and MKK7, which are the MAPKK proteins in the JNK

signaling cascade (56). Highlighting the importance of the MAPK pathways, treatment of PEL

cells with chemical inhibitors that attenuate MAPK/ERK and p38 MAPK signaling resulted in

apoptosis (57, 58).

Dysregulation of the phosphoinositide-3-kinase (PI3K)/protein kinase B (Akt)/mammalian target of rapamycin (mTOR) pathway by KSHV

There are four different classes of PI3K kinases (10). The class IA and 1B PI3K kinases

generate phosphatidylinositol-3,4,5-trisphosphate (PIP3), which recruits phosphoinositide

dependent kinase 1 (PDK1) and Akt to the membrane, where PDK1 phosphorylates and activates

Akt (Figure 1.1). The activation of Akt leads to the upregulation of several cellular processes

including glycolysis, angiogenesis, cell survival, protein synthesis, and proliferation (10, 59, 60).

Regarding protein synthesis, Akt inhibits tuberous sclerosis complex 2 (TSC2), leading to the

activation of mTORC1. The activation of mTORC1 promotes protein synthesis by activating

several translation initiation factors. The phosphorylation of the eukaryotic translation initiation

factor 4E-binding protein 1 (4EBP1) by mTORC1 releases the eukaryotic translation initiation

factor 4E (eIF4E) leading to cap-dependent mRNA translation. The phosphorylation of p70 S6

kinase (S6KB1) by mTORC1 also leads to activation of the ribosomal protein S6 and the

eukaryotic translation initiation factor 4B (eIF4B) (50, 61). Overall the activation of mTORC1

leads to the upregulation of anabolic processes to promote cell growth. Normally, energy

deficiencies and stress signals feedback into mTORC1 thereby inhibiting cell proliferation.

In non-viral cancers many genes within the PI3K/Akt/mTOR pathway are mutated to

prevent the shutdown of this pathway, while in viral cancers, oncogenic viral proteins activate

this pathway at the protein level (10, 50). The production of new virions requires energy

11

Figure 1.1 Activation of the PI3K/Akt/mTOR and MAPK/ERK Pathway in KSHV Infection. KSHV activates the PI3K and MAPK/ERK pathways during infection, which are heavily reliant on kinases (drawn as circles) to propagate the signaling cascade. Multiple viral proteins (colored in purple) activate the PI3K/Akt/mTOR pathway (in blue) to promote protein translation by activating subunits of the translation initiation complex. The MAPK/ERK pathway (in red) is similarly activated by viral proteins to promote protein translation through RSK. Some cross-talk (shown in green) occurs between both pathways. Activation of ERK1/2 also leads to the transcription of genes regulated by AP-1. Genes under the control of AP-1 include cytokines, which after being transcribed and translated are secreted out of the cell.

generation in the cell. Since mTORC1 is the master regulator of energy metabolism, KSHV

modulates this pathway to keep key biosynthesis pathways activated during the viral life cycle.

Many KSHV proteins regulate the PI3K/Akt/mTOR pathway including vIL-6, vGPCR,

K1, K15, ORF45, and ORF36 (Figure 1.1) (10, 23, 52, 61). Interestingly, these proteins are

expressed at high levels only during the lytic cycle; while K1, K15, and vIL-6 are expressed at

12

low levels during latency. A latent miRNA, miR-K3 also regulates the PI3K/Akt pathway (4). As

mentioned before, paracrine signaling from lytic cells is thought to help drive KSHV

pathogenesis by stimulating neighboring latent cells in a paracrine fashion. In the following

paragraphs below, we describe how several viral proteins or microRNAs (miRNAs) affect the

PI3K/Akt/mTOR pathway, and how their contribution in KSHV-associated malignancies may be

dependent on paracrine and autocrine signaling.

KSHV encodes several transmembrane proteins, including a chemokine receptor vGPCR,

which is a cellular homologue of the interleukin 8 receptor (15). Two other KSHV-encoded

transmembrane proteins include K1, which mimics the B-cell receptor (BCR), and K15 (62). K1

and K15 are partial functional homologues to the latent membrane protein 2A (LMP2A) encoded

by a related gammaherpesvirus, Epstein-Barr virus (EBV). Cells infected with a mutant KSHV

virus lacking K1 had attenuated activation of Akt following reactivation (63). Both K1 and

vGPCR exhibit transformative properties when expressed in cells (10). The expression of

vGPCR immortalizes endothelial cells, which is dependent on PI3K/Akt activation as addition of

a PI3K inhibitor results in apoptosis (64). Furthermore, nude mice present vascular tumors when

injected with stably expressing vGPCR-NIH3T3 cells (54). Both vGPCR- and K1-expressing

transgenic mice develop tumors (10, 65). In vGPCR-transgenic mice, vascular KS-like tumors

express CD31, an endothelial cell marker present in KS (65). However, many of the cells within

the tumor did not express the vGPCR protein and only some tissues harvested from mice with

tumors expressed ORF74 transcripts. Knockdown of vGPCR also severely attenuated tumor

growth and secretion of VEGF in a mouse model where endothelial cells transfected with a

bacterial artificial chromosome encoding KSHV were injected into nude mice (3). These

13

observations would suggest paracrine signaling by vGPCR-expressing cells to neighboring cells

play a role in the development of KSHV malignancies.

KSHV also encodes a cellular homologue of IL-6 known as vIL-6; however unlike IL-6

which requires binding to the gp130 and IL-6 receptor, vIL-6 can transduce a signal by binding

to gp130 alone (10, 12, 66). Similar to human IL-6, vIL-6 activates the JAK/STAT, MAPK, and

PI3K/Akt pathways upon binding to gp130. In endothelial cells, vIL-6 activates gp130 leading to

the upregulation of several lymphatic markers including podoplanin in a Akt- and STAT3-

dependent manner (10). Stably-expressing vIL-6 NIH3T3 cells can also induce tumor formation

in nude mice and transgenic mice expressing vIL-6 exhibit MCD-like disease (10, 66).

However, expressing vIL-6 in IL-6 deficient mice resulted in no MCD-like disease suggesting

that human IL-6 is important for MCD pathogenesis (10). Recently it was shown that the HIV-1

Nef protein synergistically augments the angiogenesis of vIL-6 in an Akt-dependent manner

(66). Endothelial cells stably expressing vIL-6 showed a significant increase in cell growth,

angiogenesis, and levels of activated Akt upon addition of soluble Nef. Highlighting the

importance of PI3K signaling, nude mice injected with NIH3T3 cells co-expressing Nef and vIL-

6 failed to develop any tumor growth upon treatment with a PI3K inhibitor. These results suggest

a novel mechanism on how HIV-infection can play a role in the development of KSHV

malignancies besides immune suppression.

KSHV ORF36 encodes a serine/threonine viral protein kinase (vPK) conserved among

the herpesviruses, and has been shown to activate JNK, a MAPK family member (56, 61, 67).

Though vPK is classified as a late lytic gene, it can be expressed in hypoxic conditions. Knock

down of vPK negatively affects lytic replication and attenuates expression of several late genes

(67). A ~3 to 10 fold reduction in viral progeny production and infection was observed in

14

deficient or kinase-dead vPK viruses. We recently reported that vPK can mimic S6KB1 within

the mTOR pathway and residues lining the catalytic pocket of vPK are conserved to S6KB1 (61).

Similar to S6KB1, vPK can phosphorylate S6, which was inhibited upon addition of a S6KB1

specific inhibitor, but not a JNK or Aurora kinase inhibitor. Overexpression of vPK in cells

augmented anchorage-independent growth, angiogenesis, and cell proliferation as well.

Indicative of vPK’s possible role in KSHV-associated malignancies, transgenic vPK mice in a

C57BL/6 background resulted in an eight-fold higher incidence of B-cell lymphomas in aged

mice (68). Lymphomas harvested from these mice were of germinal center and post-germinal

center origin, and exhibited activation of S6, which is downstream of mTORC1. Furthermore,

IL-1β and IL-12 p40 were elevated in the serum of vPK transgenic mice in comparison to the

wild-type (WT) control mice. Both of these inflammatory cytokines have been implicated in

different cancers and may contribute to the development of these lymphomas in the vPK

transgenic mice (69-72).

The microRNAs (miRNA) encoded by KSHV are transcribed from the latent

Kaposin/K12 promoter and are highly expressed in KSHV-associated malignancies (4). One

miRNA, miR-K3 was shown to activate Akt by downregulating the G protein-coupled receptor

kinase 2 (GRK2) which suppresses expression of an IL-8 receptor, C-X-C chemokine receptor

type 2 (CXCR2) (4). Knockdown of CXCR2 led to decreased activation of Akt in endothelial

cells similar to what was shown previously in skin keratinocytes. Furthermore, GRK2 can

interact and inhibit Akt signaling in CXCR2-independent manner. The expression of miR-K3

resulted in enhanced cell migration and invasion of endothelial cells in an Akt-dependent

manner.

15

As seen by the many proteins encoded by KSHV that regulate the PI3K/Akt/mTOR

pathway, this pathway plays an important role in the KSHV life cycle. Indeed, the

PI3K/Akt/mTOR pathway is important for cell survival in KSHV-infected cells. Treatment of

PEL cells with rapamycin, an inhibitor of mTORC1, inhibited cell growth in vitro and

significantly attenuated tumor growth in mice (73). Low doses of NVP-BEZ235, a dual inhibitor

that targets both PI3K and mTOR resulted in apoptosis in PEL cells (6). Inhibition of the

PI3K/Akt/mTOR pathway also attenuated cytokine secretion from PEL cells as well (6, 73). In

renal-transplant patients, KS biopsies at the time of diagnosis showed elevated levels of activated

Akt and S6KB1, and increased vascular endothelial growth factor (VEGF) expression (74).

Treatment of these patients with Sirolimus (rapamycin) resulted in remission of KS in all patients

by six months. Therefore, inhibiting PI3K signaling has shown promise in a variety of KSHV-

associated diseases.

In this dissertation (Chapter 2), we expand on how PEL cells have upregulated Tyro3, a

host receptor tyrosine kinase. Tyro3 is a member of the TAM receptor family consisting of

Tyro3, Axl, and MerTK, whose activation can lead to the upregulation of several signaling

pathways including the PI3K/Akt/mTOR pathway (75). The TAM receptors have been shown to

be upregulated in many different cancers, thereby promoting cell survival and chemoresistance

(76-78). A group previously showed that Axl was upregulated in Kaposi’s sarcoma (KS), but not

MerTK or Tyro3 (79). The expression of Axl in KS promoted cellular proliferation by

upregulating PI3K signaling. Furthermore, in a KS xenograft mouse model, treatment of mice

with a monoclonal antibody against Axl reduced tumor growth. We utilized multiplexed

inhibitor beads (MIB) coupled with mass spectrometry (MS) to profile the kinome of primary B-

cells and several different subtypes of non-Hodgkin lymphoma (NHL) including PEL. From our

16

MIB/MS screen, we identified Tyro3, but not Axl or MerTK, as being uniquely upregulated in

PEL. Knockdown of Tyro3 resulted in an attenuation of PI3K signaling, cell survival, and colony

formation in PEL cells. Furthermore, treatment of PEL cells with a Tyro3 inhibitor, UNC3810A,

resulted in a decrease in tumor burden in a PEL xenograft mouse model. Overall, this work

identifies a new mechanism by which the PI3K pathway is modulated in PEL, and that targeting

Tyro3 may provide clinical benefits in the treatment of patients with PEL.

Autophagy

As mentioned above, there are four classes of PI3K kinases. There is one class III PI3K

kinase in humans known as VPS34 (encoded by the PIK3C3 gene), whose role is to regulate

macroautophagy (80). Autophagy is the process by which proteins and organelles in cells can be

degraded into their building blocks, which can then be recycled to build other proteins and

organelles. Macroautophagy is a non-selective process by which bulk cargo in the cell is

degraded. The converse is selective autophagy, where different organelles and proteins are

selectively targeted for degradation. Types of selective autophagy include mitophagy

(degradation of mitochondria), pexophagy (degradation of peroxisomes), ribophagy (degradation

of ribosomes), and reticulophagy (degradation of the endoplasmic reticulum) (81). Proper

induction of autophagy is important to maintain a healthy cell by preventing the buildup of old or

defective proteins and organelles. In humans, autophagy is an important cellular process by

which the cell can respond to stress such as nutrient starvation, hypoxia, ER stress, microbial

infection, and low energy levels (82). The maturation of the phagosome to engulf cellular cargo

can be defined by four steps: (1) induction and nucleation of the phagophore assembly site

(PAS), (2) expansion of the phagophore, (3) completion of the phagosome and fusion with a

lysosome, and (4) degradation of the cargo in the autolysosome (82).

17

The most well-studied mechanism by which macroautophagy (henceforth called

autophagy) is induced is during nutrient starvation (83). The Autophagy related 1 (Atg1)

complex, which consists of Atg1, Atg13, Atg17, Atg29, and Atg31 controls the recruitment of

other Atg proteins to the phagophore assembly site (PAS) in a process known as nucleation to

initiate autophagy (82). The mammalian homologues to Atg1 are the ULK proteins - with the

main family members being ULK1 and ULK2 (83). AMP activated protein kinase (AMPK) and

mTORC1 are the main regulators of the ULK1 complex (83). Under normal conditions,

mTORC1 phosphorylates ULK1 resulting in its inactivation (84). However under nutrient

starvation, AMPK phosphorylates and activates ULK1 (84). The activation of the ULK1

complex in turn allows the recruitment and activation of the class III PI3K complex, which

includes VPS34, Beclin-1 (the mammalian homolog of Atg6), and Atg14L (82, 83). The

activation of this complex promotes nucleation, and thus the formation of the autophagosome.

The expansion of the phagophore is dependent on the recruitment of Atg8 (also known as LC3 in

mammals), Atg9, and Atg12 to the PAS (82). Both LC3 and Atg12 undergo ubiquitin-like

conjugation reactions (in a reminiscent manner similar to the E1-, E2-, and E3-like enzyme chain

reaction as seen in the priming and attachment of ubiquitin to substrates). Atg12 is conjugated to

Atg5 by Atg7 and Atg10 (81). Conjugated Atg5-Atg12 forms a complex with Atg16, which is

recruited to the phagophore. The Atg5-Atg12-Atg16 complex helps recruit the lipidated form of

LC3 (known as LC3-II) to the phagophore (81). The generation of LC3-II from LC3 also

involves a similar ubiquitin-like conjugation system. LC3 is first cleaved by Atg4 on the C-

terminus in the cytoplasm generating LC3-I (82). The conjugation of LC3-I to

phosphatidylethanolamine (PE) is then performed by Atg3 and Atg7 generating LC3-II (81, 82).

Recruitment of LC3-II to the phagophore results in elongation and eventual formation of the

18

autophagosome. The fusion of a lysosome to the autophagosome results in an autolysosome,

where the cargo originally enclosed in the autophagosome is degraded.

The expression of several Atg proteins is upregulated in response to stress. A mechanism

by which this occurs is the translation of a differential set of mRNAs under stress conditions.

One of the central proteins in the integrated stress response (ISR) is the eukaryotic initiation

factor 2 subunit alpha (eIF2α) (85, 86). The eukaryotic initiation factor 2 (eIF2) complex -

consisting of eIF2α, eIF2β, and eIF2γ - initiates mRNA translation by recognizing the AUG start

codon, and by forming the 43S pre-initiation complex (PIC) with other translation initiation

factors (85). The 43S PIC is then recruited to mRNA containing a 5’ N7 methylated guanosine

cap (known as the 5’ cap) (87). The loading of the Met-tRNA to the AUG start codon results in

the hydrolysis of GTP to GDP on eIF2. After placement of the methionine (Met)-transfer RNA

(tRNA), the 43S PIC disassociates, and the complexes are recycled to initiate another round of

translation. The association of GTP with eIF2 determines its ability to associate with the 43S PIC

(85). Therefore after a round of translation initiation, the guanine nucleotide exchange factor

(GEF), eIF2B, must exchange GDP with GTP on eIF2. However, under stress conditions, eIF2α

becomes phosphorylated on the serine 51 residue, which inhibits the ability of eIF2B to

exchange out GDP to GTP (85). This prevents the formation of the 43S PIC and ultimately shuts

down cap-dependent translation in the cell. On the other hand, the translation of mRNAs

containing a short upstream open reading frame (uORF) in the 5’ untranslated region (UTR)

occurs instead as they are not dependent on 5’ cap-dependent translation (85). A couple of these

mRNAs include the transcription factors, ATF4 and CHOP (88). Synthesis of these transcription

factors results in the production of stress-related proteins including some of the Atg proteins like

19

isoform B of LC3 (LC3B) (89, 90). Therefore, regulation of autophagy is two-fold, and involves

the production of the autophagy machinery, and the maturation of the autophagosome.

Due to the importance of autophagy in maintaining cell homeostasis, dysregulation of

autophagy has been tied to several neurodegenerative diseases that are characterized by the

buildup of toxic proteins such as in Alzheimer’s and Parkinson’s disease (91). Autophagy has

also been linked to cancer, but its role in tumor development is less clear (92). Autophagy seems

to confer a more tumor suppressive phenotype during early stages of tumor progression, but is

more protective during late stages of tumor progression (92). The role of autophagy in the

herpesvirus life cycle is complex with many different viral proteins encoded by several

herpesvirus either inhibiting or inducing autophagy (which is reviewed in depth by Lussignol

and Esclatine, see reference (93)). Many of the herpesvirus induce autophagy early on in the lytic

cycle, but block the process in later stages of the lytic cycle (93). Several of the herpesviruses

block the formation of the autolysosome, and utilize the autophagosomes generated as a

precursor for the generation of viral particles (93-97). KSHV upon reactivation from latency

induces autophagy early on in the lytic cycle (97). Interestingly, the autophagic process during

the lytic cycle may be independent of mTORC1 regulation as treatment with the mTOR

inhibitor, Torin, did not affect autophagic flux or induction of autophagy markers (98).

Expression of KSHV-encoded RTA, the main protein which controls reactivation from latency,

is able to increase autophagic vesicle formation as seen by transmission electron microscopy

(TEM) (99). In reactivated BC3 PEL cells, viral particles were also observed in autophagic

vesicles (97). However when examining autophagic flux in reactivated BC3 cells at 36 hours

post-infection, autophagic flux was inhibited (97). Measuring autophagic flux is important as it

examines if the induction of autophagy results in actual degradation of the cargo (100). The

20

upregulation of autophagy markers, yet the inhibition of autophagic flux indicate that the

autophagic process is initiated, but the cargo is not degraded. KSHV-encoded K7 protein binds

Rubicon, which is a cellular protein that inhibits autophagosome maturation (101). Expression of

K7 did not alter autophagosome formation, but decreased the amount of autolysosomes.

Furthermore, KSHV encodes a viral homolog of B-cell lymphoma 2 (vBCL-2), which can bind

and inhibit Beclin-1 similar to cellular Bcl-2 to inhibit nucleation (102-104). On the other hand,

KSHV inhibits autophagy during latency. While, the viral cyclin protein (vCyclin) induces

autophagy due to activating the DNA damage response (DDR), co-expression of vCyclin with

vFLIP, demonstrated that vFLIP was able to inhibit vCyclin-mediated autophagy (105-107).

Highlighting the potential role of autophagy in herpesvirus-associated cancers, the EBV-encoded

latent membrane protein 2A (LMP2A) is able to induce autophagy in nonmalignant breast

epithelial cells to prevent anoikis, a type of cell death that occurs when adherent cells become

detached to the extracellular matrix (108).

Mitophagy

A selective form of autophagy, which degrades mitochondria, is known as mitophagy

(109). The recycling of old and defective mitochondria is important for the health and proper

energy homeostasis in the cell. Highlighting the importance of maintaining healthy

mitochondria, 1 in 2,000 individuals are affected with diseases related to mitochondria-

dysfunction (110). Mitophagy occurs under basal conditions, but can be upregulated under

stress-conditions such as in hypoxia conditions or nutrient starvation (111). The central step in

mitophagy is the recruitment of mitophagy-associated receptors, which helps trigger the

recruitment of the autophagy machinery including ULK1 (109). These receptors have an LC3

interacting region (LIR), which help recruit LC3 to the mitochondria. While there are many

mitophagy receptors, the two main processes of receptor recruitment are characterized by PTEN-

21

induced kinase 1 (PINK1)-Parkin ubiquitin-dependent or hypoxia-mediated mitophagy (Figure

1.2).

Figure 1.2 Generation of the phagophore in mitophagy. The generation of the phagophore during mitophagy primarily occurs through either hypoxia-dependent mitophagy or ubiquitin-dependent mitophagy. During hypoxia-dependent mitophagy, hypoxic conditions lead to the upregulation of certain mitophagy receptors that localize to the mitochondria to recruit LC3. In contrast, ubiquitin-dependent mitophagy requires the stabilization of PINK1 in the mitochondrial membrane. Stabilized PINK1 then activates Parkin, which ubiquitinates outer mitochondrial membrane (OMM) proteins. Ubiquitin acts as docking sites for ubiquitin-dependent mitophagy receptors, which upon localizing to the mitochondria recruit LC3. In both cases, the recruitment of LC3 results in the development of the phagophore. Figure 1.2 is modeled from Figure 1 from Palikaras et. al., 2018 (111).

During normal conditions, the matrix processing peptidase (MPP) processes PINK1,

which is then further cleaved by the inner membrane protease PINK1/PGAM5-associated

rhomboid-like protease (PARL) (109, 112). Truncated PINK1 translocates to the cytosol, where

22

it is degraded. However, under mitochondrial membrane depolarization, PINK1 is stabilized and

becomes activated (109). Afterwards, PINK1 phosphorylates ubiquitin present on the

mitochondria on residue serine 65 (112). Parkin, an E3 ubiquitin ligase, is able to bind phospho-

ubiquitin on the mitochondria, and becomes phosphorylated by PINK1. This results in the

activation of Parkin, which leads to Parkin’s ubiquitination of mitochondrial proteins (109). The

ubiquitinated mitochondrial proteins, then act as docking sites for the mitophagy receptors. The

five main receptors are optineurin (Optn), sequestosome-1 (p62), nuclear domain 10 protein 52

(NDP52), TAX1 binding protein (TAX1BP1), and neighbor of BRCA1 gene (NBR1) (112). Of

the five, Optn and NDP52 are the primary receptors as cells with all five receptors knocked out

were only rescued by Optn and NDP52 (113). After receptors bind to the mitochondria, the

autophagy machinery is recruited to begin phagophore formation, which leads to the eventual

degradation of the mitochondrion (113). In hypoxia-mediated mitophagy, the receptors are

regulated by hypoxia (112). These receptors are BCL2/Adenovirus E1B 19 kDA interacting

protein 3 (BNIP3), Nip3-like protein X (NIX – also known as BNIP3L), and Fun14 domain

containing 1 (FUNDC1) (112). Both BNIP3 and NIX expression levels increase in hypoxic

conditions (109). BNIP3 is able to stabilize PINK1 to promote mitophagy (114). On the other

hand, FUNDC1 is phosphorylated by Src and casein kinase II (CK2), which prevents its

interaction with LC3 (112). In hypoxic conditions, phosphoglycerate mutase 5 (PGAM5)

dephosphorylates FUNDC1, thereby relieving this suppression (112).

The role of mitophagy is not well-studied in herpesviruses. Recently, a group showed that

mitophagy is induced during lytic reactivation in KSHV in a viral interferon regulatory factor 1

(vIRF1)-dependent manner (115). Mitochondrial content was decreased and increased

localization of mitochondria to the lysosomes was seen upon reactivation in PEL cells. The

23

induction of mitophagy was shown to be dependent on vIRF1-mediated activation of NIX.

Inhibiting mitophagy resulted in an attenuation of viral protein expression and replication.

Interestingly, depletion of NIX had little effect on viral replication and ORF45 protein

expression. This suggests there may be alternative mechanisms that KSHV uses to regulate

mitophagy. In this dissertation, we examine how vPK, a kinase encoded by ORF36 in KSHV,

may regulate mitophagy (Chapter 3). Expression of vPK in human umbilical vein endothelial

cells (HUVECs) results in an increase in phosphorylated-eIF2α and LC3B-II expression.

However, protein expression of alternative transcripts such as ATF4 and CHOP was not induced.

A group that pulled down all the open reading frames (ORFs) in KSHV to identify host cellular

interacting partners to KSHV proteins noted that vPK heavily interacted with mitochondrial

proteins (116). When fractionating mitochondria, we found that vPK was present in

mitochondria, and that mitochondrial content decreased in vPK-expressing cells. Our work

suggests that vPK may induce mitophagy in the KSHV life cycle.

Conclusions

The modulation of signaling pathways by viral proteins is critical to the survival and

persistence of KSHV in the human host. These pathways rely on kinases to induce a signaling

cascade by phosphorylating proteins to recruit and/or alter activity of target substrates. Inhibition

of these pathways by biochemical means or drug treatment often attenuates viral replication

and/or induces cell death in KSHV-infected cells (6, 24, 73). Therefore, kinase inhibitors have

represented an attractive target in treating KSHV malignancies (74, 117). Currently there are

clinical trials to test the efficacy of Selumetinib, an inhibitor of the dual specificity mitogen-

activated protein kinase kinase 1/2 (MEK1/2) in KS patients, and Sirolimus (rapamycin), an

mTOR complex 1 (mTORC1) inhibitor in HIV-positive MCD patients (1). Already several

kinase inhibitors such as Sirolimus and Imatinib have had partial success in treating KS patients

24

(74, 117). Active drug discovery aimed at cellular kinases comprises a large proportion of

pharmaceutical drug development. Therefore, novel specific kinase inhibitors identified by drug

discovery programs may be viable treatment options for KSHV associated diseases. In this

dissertation, we aim to further characterize how both host (Chapter 2) and virally-encoded

(Chapter 3) kinases play a role in KSHV-infection and pathogenesis. This work allows us to gain

a better understanding of how kinases modulate pathways that are important in the KSHV life-

cycle.

25

CHAPTER 2: KINOME PROFILING OF NON-HODGKIN LYMPHOMA IDENTIFIES

TYRO3 AS A THERAPEUTIC TARGET IN PRIMARY EFFUSION LYMPHOMA2

Introduction

Non-Hodgkin lymphoma (NHL) consists of many subtypes covering a wide range of

characteristics that include differences in cell-of-origin, aggressiveness, and viral association.

One of these NHL subtypes is primary effusion lymphoma (PEL), which is associated with

Kaposi’s sarcoma-associated herpesvirus (KSHV). Patients with PEL have a median survival

time of 6 months after diagnosis, as standard chemotherapy is generally ineffective in treating

PEL (118, 119). Several clinical studies have taken different approaches to treat PEL, and while

some have shown promise, there is still no effective treatment for PEL (120). For example,

treatment of PEL with bortezomib, a proteasome inhibitor that inhibits NF-κB activity, has

shown variable results in clinical trials (121-123). Therefore, there is a need to identify new

targeted therapies to treat PEL.

Signaling pathways are often dysregulated in cancer, and kinases that control the

propagation of these signaling cascades frequently have aberrant activity. The use of kinase

inhibitors to treat cancers has shown clinical efficacy as a single agent or in the adjuvant setting.

For example, imatinib, an inhibitor of the BCR-ABL kinase, is efficacious in treating chronic

2This chapter previously appeared as an article in the journal PNAS. The original citation is as follows: Wong JP, Stuhlmiller TJ, Giffin LC, Lin C, Bigi R, Zhao J, Zhang W, Bravo Cruz AG, Park SI, Earp HS, Dittmer DP, Frye SV, Wang X, Johnson GL, and Damania B. Copyright © PNAS. 2019 Aug 13; 116(33):16541-16550. doi: 10.1073/pnas.1903991116.

26

myelogenous leukemia (20). We were interested in identifying kinases that were uniquely

activated in PEL in comparison to other NHL subtypes. A kinase with unique or significantly

higher activity in PEL compared to other NHL subtypes may indicate that PEL is dependent on

the signaling activity of that particular kinase compared to other NHL subtypes.

We decided to compare PEL to four other B cell NHL subtypes: follicular lymphoma

(FL), diffuse large B-cell lymphoma (DLBCL), mantle cell lymphoma (MCL), and Burkitt’s

lymphoma (BL). These subtypes are derived from different stages of B-cell development that

include the mantle zone, germinal center, and post-germinal center (124-129). As different B-cell

NHL subtypes may arise during different stages of normal B-cell development, there are likely

differences in the expressed kinome (130). Two of the subtypes (BL and PEL) are commonly

associated with viral infection. BL is associated with Epstein-Barr virus (EBV) infection, while

PEL is distinguished by Kaposi’s sarcoma-associated herpesvirus (KSHV) infection (131-133).

Co-infection with EBV is also common in PEL (134-136). By examining an array of NHL

subtypes that are derived from different stages of B-cell development, and being virally-

associated or not, we intended to capture a wide enough spread in cellular phenotypes to identify

distinct kinases which are activated or repressed in PEL in comparison to other B-cell NHL

subtypes.

Materials and methods

Mice

Nod scid gamma (NSG) mice used in the PEL and BJAB xenograft studies were

maintained in pathogen-free conditions and the studies were approved by the Institutional

Animal Care and Use Committee (IACUC) at the University of North Carolina at Chapel Hill.

Four to five-week old NSG female mice weighing approximately 20 grams used in the xenograft

studies were purchased from The Jackson Laboratory (005557). The pharmacokinetic (PK)

27

analysis was completed by Sai Life Sciences Limited. The study was conducted in accordance to

guidelines provided by the Committee for the Purpose of Control and Supervision of

Experiments on Animals (CPCSEA) as published in The Gazette of India (1998) and approved

by the Institutional Animal Ethics Committee (IAEC). Eight to twelve-week old Swiss Albino

male mice weighing approximately 25 to 35 grams used in the PK study was purchased from

Global, India.

Cell Culture

All cell lines are grown at 37oC at 5% CO2 levels. NHL cell lines except for OCl-Ly1,

OCl-Ly3, and PEL cell lines were grown in RPMI 1640 medium supplemented with 10% FBS,

1% penicillin-streptomycin, and 1% L-glutamine. PEL lines were grown in the same complete

RPMI media as described above supplemented additionally with 0.075% sodium bicarbonate

(Gibco, 25080-094) and 0.05 mM β-mercaptoethanol (Gibco, 21985-023). OCl-Ly1 and OCl-

Ly3 were grown in Iscove’s modified Dulbecco’s medium (IMDM), 10% human AB serum

(Sigma-Aldrich, H4522), 1% penicillin-streptomycin, and 75 µM β-mercaptoethanol (Gibco,

21985-023). 293FT and B16-F10 cells were grown in DMEM supplemented with 10% FBS, 1%

penicillin-streptomycin, and 1% L-glutamine. 500 µg/ml of G418 (Gibco, 10131-027) was also

added for selection in 293FT cells. HT-29 cells were grown in McCoy’s 5a medium

supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% L-glutamine. Human

umbilical vein endothelial cells (HUVEC) were grown in endothelial cell basal medium 2

(PromoCell, C-22211) supplemented with 1% L-glutamine, 10% FBS, 1% penicillin-

streptomycin, and the growth medium 2 supplement pack (PromoCell, C-39211) excluding the

addition of heparin and ascorbic acid from the pack. Cell lines knocked down for Tyro3 also had

4 µg/ml of puromycin (Corning, 61-385-RA) added for selection. Trex-RTA Luc BCBL1 cells

28

were grown in RPMI 1640 medium supplemented with 10% Tet-free FBS (Clontech, 631107),

1% penicillin-streptomycin, 1% L-glutamine, 20 µg/ml hygromycin B (Corning, 30-240-CR),

and 1.25 µg/ml puromycin (Corning, 61-385-RA). All cell lines were tested free of mycoplasma

contamination using the LookOut Mycoplasma PCR Detection kit. All PEL cell lines were

authenticated by STR profiling using the GlobalFiler STR panel.

Lentiviral infection of cell lines

To generate Tyro3 knockdown cell lines, plasmids containing MISSION pLKO.1-puro

control (Sigma, SHC002), TRC2 pLKO.5-puro control (Sigma, SHC202), or the following

Tyro3 shRNA (Sigma, SHCLNG-NM_006293): #1524 (Clone NM_006293.2-1664s21c1),

#1527 (Clone NM_006293.2-2755s21c1), #9716 (Clone NM_006293.2-1718s21c1), #2178

(Clone NM_006293.x-2490s1c1), and #2179 (Clone NM_006293.x-743s1c1) were used. Clones

#2178 and #2179 contain the pLKO.1 backbone; clones #1524, #1527, and #9716 contain the

pLKO.5 backbone. Lentiviruses containing these plasmids were then made in 293FT using

ViraPower (Invitrogen, K497500) according to the manufacturer’s instructions. B-cells were

centrifuged with lentiviruses containing either the control plasmid or shTyro3 for 90 minutes at

2,500 rpm at 30oC in serum-free medium containing 10 µg/ml polybrene (Sigma, 107689). After

centrifugation, cells were incubated overnight. The next day, cells were spun down and fresh

complete media was added. Two days after infection, 1 µg/ml puromycin (Corning, 61-385-RA)

selection was added. The concentration of puromycin was doubled after each subsequent split to

a final concentration of 4 µg/ml. Trex-RTA Luc BCBL1 cells were made from Trex-RTA

BCBL1 cells (obtained as a gift from J. Jung) that were infected with RediFectTM Luc

lentiviruses (PerkinElmer, CLS960002) according to the manufacturer’s instructions (137).

29

Primary B-cell isolation

Human B-cells were isolated from whole human blood purchased from Gulf Coast

Regional Blood Center (E5318). Blood was tested by Gulf Coast for human immunodeficiency

virus 1 (HIV-1) RNA, HIV-1 Group M RNA, HIV-1 Group O RNA, HIV-2 RNA, hepatitis C

virus (HCV) RNA, hepatitis B virus (HBV) DNA, West Nile virus (WNV) RNA, Zika RNA,

hepatitis B surface antigen, and for antibodies to HIV-1/2+O, hepatitis B core antigen, HCV,

human T-lymphotropic virus I/III (HTLV-I/III), CMV, and syphilis. Trypanosoma cruzi was also

tested for if patient history indicated an exposure risk since previous testing, otherwise noted as

historical non-reactive. B-cells were only used from blood that tested non-reactive or negative to

all of the above. Briefly, 20 mL of blood was transferred to a 50 mL conical tube and diluted

with 20 mL of 2 mM EDTA-PBS. The diluted blood was then layered slowly over 15 mL of

Ficoll-Paque (GE Healthcare, 17-5442-02) in SepMate-50 tubes (StemCell Technologies,

85450). The tubes were centrifuged at 1500 rpm for 20 minutes, after which the top layer

containing PBMCs was isolated. Cells were washed with 2 mM EDTA-PBS, and pelleted at

1500 rpm at 4oC for 5 minutes. Remaining blood cells were lysed by resuspending the cell pellet

in ACK lysing buffer (Gibco, A10492-01) and was incubated for 10 to 15 minutes at room

temperature. The suspension was further washed with 2 mM EDTA-PBS and resuspended in 2

mM EDTA, 0.5% BSA (Sigma-Aldrich, A7906) in PBS. B-cells were isolated with the B-cell

isolation kit II (Miltenyi Biotec, 130-091-151) according to manufacturer’s instructions. Cells

were stained with CD19 (Miltenyi Biotec, 130-091-247) and ran on a MACSQuant VYB

cytometer to determine purity. Analysis was performed with FlowJo v10.2 software. B-cell

isolations containing ≥95% CD19+ cells by flow analysis were used in further experiments.

30

MIB/MS affinity chromatography

Suspension cells were pelleted and washed three times in cold PBS. Pellets were lysed on

ice in MIB lysis buffer [50 mM HEPES (N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid)

pH 7.5, 0.5% Triton X-100, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 10 mM sodium

fluoride, 2.5 mM sodium orthovanadate, 1X Roche protease inhibitor cocktail (Sigma-Aldrich,

11836145001), and 1% each of phosphatase inhibitor cocktail 2 (Sigma-Aldrich, P5726) and 3

(Sigma-Aldrich, P0044)]. Cell lysates were sonicated 3×10 seconds on ice and centrifuged at

16,000×g in microfuge tubes for 10 minutes at 4°C. The supernatant was syringe-filtered through

a 0.2 µm SFCA membrane. Protein concentration was determined by Bradford assay (Bio-Rad,

500-0006) and equal amounts of protein, around 2.5 mg, were used for each sample.

The filtered lysate was brought to 1 M NaCl and passed through a column of multiplexed

kinase inhibitor-conjugated beads (MIBs) consisting of Sepharose-conjugated Purvalanol B,

PP58, CTx-0294885, VI-16832, and two custom-synthesized pan-kinase inhibitor compounds,

UNC-8088A and UNC-2142A (53). The MIBs were washed with 5 mL of high-salt buffer and 5

mL of low-salt buffer [50 mM HEPES (pH 7.5), 0.5% Triton X-100, 1 mM EDTA, 1 mM

EGTA, and 10 mM sodium fluoride, and 1 M NaCl or 150 mM NaCl, respectively]. The

columns were washed a final time with 1 mL 0.1% SDS before elution in 1 mL of 0.5% SDS

(100°C, 5 min). Eluted kinases were reduced (dithiothreitol) and alkylated (iodoacetamide) prior

to being concentrated with Amicon Ultra centrifugal filters (Millipore, UFC801096) and

detergent was removed from the concentrated eluate by chloroform/methanol extraction. Protein

pellets were resuspended in 50 mM HEPES (pH 8.0) and were digested overnight with

sequencing grade modified trypsin (Promega, V5111). Peptides were dried down in a speed-vac

31

and cleaned with C-18 spin columns (Thermo Scientific, 89870) according to the manufacturer

instructions.

Peptides were resuspended in 5% acetonitrile and 0.1% formic acid, and 25-50% injected

onto a Thermo Easy-Spray 75uM x 25cm C-18 column via an Easy nanoLC-1000. Peptides were

separated as a single fraction on a 300 minute gradient (5-40% acetonitrile) and identified by a

Q-Exactive mass spectrometer. Parameters: 3e6 AGC MS1, 80ms MS1 max inject time, 1e5

AGC MS2, 100ms MS2 max inject time, 20 loop count, 1.8 m/z isolation window, 45s dynamic

exclusion. Spectra were identified using MaxQuant software and the Uniprot/Swiss-Prot

database (138). Peptide abundance was calculated using label-free quantification (LFQ). Mass

spectrometry runs were normalized by natural log-transformed and median-centering LFQ values

on a common-captured set of 151 kinases. Heat maps were generated using the Euclidian

hierarchical clustering program from GENE-E (https://software.broadinstitute.org/GENE-E/).

RNA-sequencing (seq)

RNA was harvested from cell lines using the RNeasy Plus Mini Kit (Qiagen, 74136)

according to manufacturer’s instructions. For sequencing of the NHL cell lines, mRNA-Seq

libraries were generated using the Stranded mRNA-Seq kit (KAPA Biosystems, KK8421) and

multiplexed with Illumina TruSeq adapters. Samples were run on two 75-cycle single-end

sequencing runs with an Illumina NextSeq-500. QC-passed reads were aligned to hg19 using

MapSplice (139). Picard Tools v1.64 was used to determine the alignment profile. Aligned reads

were sorted and indexed using SAMtools and translated to transcriptome coordinates and filtered

for indels, large inserts, and zero mapping quality using UBU v1.0 (140). Transcript abundance

estimates were performed using an Expectation-Maximization algorithm (141).

32

For analyzing the RNA-seq data from Journo et. al., 2018 (142), the RNA-SEQ Analysis

tool from the CLC Genomics Workbench (v. 11.0) program was used. The defaults set in the

program were used for analysis (with the exception of strand specificity being set as reverse).

RNA isolation and real-time (RT) PCR

RNA was harvested from cell lines and primary B-cells using the RNeasy Plus Mini Kit

(Qiagen, 74136) following the manufacturer’s protocol. Pleural effusions from patients with PEL

were obtained through the AIDS and Cancer Specimen Resource (ACSR). RNA was harvested

from the pleural effusion from patients with PEL using TRIzol (Invitrogen, 15596018) following

the manufacturer’s instructions. RNA was then reverse-transcribed using the iScriptTM gDNA

Clear cDNA Synthesis kit (Bio-Rad, 1725035), and RT-PCR was performed using the Bioline

SYBR® Lo-ROX kit (BIO-94020) on an Applied Biosystems Quantstudio 6 Flex RT-PCR

system. Primers used are listed below and had been previously reported (143-146).

Table 2.1 Primers used for RT-PCR analysis

Immunoblotting

Cells were pelleted and washed once in cold PBS before being lysed in either NP-40 [50

mM Tris (tris(hydroxymethyl)aminomethane)-HCl (pH 8), 150 mM NaCl, and 0.1% NP-40] or

RIPA lysis buffer [50 mM Tris-HCl (pH 8), 150 mM NaCl, 1% NP-40, 0.1% SDS, 0.5% sodium

deoxycholate] containing phosphatase inhibitors [30 mM β-glycerophosphate, 50 mM sodium

fluoride, and 1 mM sodium orthovanadate] and 1x Roche protease inhibitor tablet (Sigma-

33

Aldrich, 11836145001). When probing for hard to detect or membrane-bound proteins (such as

p85 PI3K and AKT), samples were lysed in RIPA buffer; otherwise NP-40 was used to lyse

cells. Lysates in RIPA buffer were also sonicated using a Branson Digital Sonifier for 10 seconds

at an amplitude of 15% prior to centrifugation. Lysates were clarified of cellular debris, and

protein quantification was performed afterwards using either a Bradford assay (Bio-Rad, 500-

0006) for lysates in NP-40 or a BCA assay (Thermo Scientific, 23227) for lysates in RIPA.

Lysates were loaded with 5x urea buffer [10% w/v SDS, 10 mM β-mercaptoethanol, 20% v/v

glycerol, 0.2 M Tris-HCl (pH 6.8), 0.05% w/v bromophenol blue, and 8M urea] and proteins

were resolved by SDS-PAGE and then transferred onto a nitrocellulose membrane. Primary

antibodies purchased from Cell Signaling Technology were used to detect Tyro3 (1:1000,

5585S), α-tubulin (1:5000, 9099S), p-p85 PI3K Y458 (1:1000, 4228), total p85 PI3K (1:1000,

4292), p-AKT S473 (1:1000, 4060), total AKT (1:1000, 9272), p-S6 (1:10000, 4858), total S6

(1:5000, 2217), caspase-3 (1:1000, 9665), cleaved-PARP (1:1000, 5625), total PARP (1:1000,

9542), MerTK (1:1000, 4319), and Axl (1:2500, 8661). Primary antibody used to detect GAPDH

(1:5000, sc-25778) was purchased from Santa Cruz. Blots that were probed for phosphorylated

proteins were stripped with RestoreTM Stripping Buffer (Thermo Scientific, 46430) and reprobed

with the total antibody. Blots were developed with Clarity Western ECL substrate (Bio-Rad,

1705060) or ECL Prime (GE Healthcare, RPN2232). Blots were imaged using a ChemiDoc

imaging system and viewed with Image Lab v5.2.1 software.

For mice tumors harvested from the BCBL1-xenograft model, a pea-sized tumor was cut

and lysed in T-PER Tissue Protein Extraction Reagent (Thermo Scientific, 78510) according to

the manufacturer’s instructions.

34

Synthesis of UNC3810A (1)

A solution of 2-ethylbutan-1-amine (10.1 g, 100 mmol) and trans-4-(5-bromo-2-chloro-

7H-pyrrolo[2,3-d]pyrimidin-7-yl)cyclohexan-1-ol (3) (16.5 g, 50 mmol) in iPrOH (50 mL) was

added to potassium carbonate (K2CO3) powder (6.91 g, 50.0 mmol) and diisopropylethylamine

(DIPEA) (100 mmol, 17.5 mL). The mixture was heated at 120 oC for 4 days in a sealed tube

under N2, then quenched with water, and extracted with ethyl acetate (EtOAc) (3X). The

combined organic layer was washed with water and brine, then dried (Na2SO4). The residue was

purified by recrystallization in a mixture of hexane/EtOAc to yield the desired trans-4-(5-bromo-

2-((2-ethylbutyl)amino)-7H-pyrrolo[2,3-d]pyrimidin-7-yl)cyclohexan-1-ol (4) as a pale yellow

solid. The mother liquid was concentrated and purified by ISCO (0~60% EtOAc in petroleum

ether) to afford additional 4 (combined: 18 g, 91%).

To a solution of 4 (11.9 g, 30.0 mmol) in ethanol (200 mL) and water (50 mL) was added

K2CO3 (12.4 g, 90.0 mmol), tetrakis(triphenylphosphine)palladium (Pd(PPh3)4) (0.867 g, 0.750

mmol), and 1-methyl-4-(4-(4,4,5,5-tetramethyl-1,3,2-dioxaborolan-2-yl)benzyl)piperazine (10.4

g, 33.0 mmol). The reaction mixture was heated at 60oC under N2 for 5 hours (LC-MS indicated

consumption of the starting material), then was concentrated and extracted by EtOAc (3X). The

combined organic layer was washed with water and brine, then dried (Na2SO4). The residue was

purified by ISCO (methanol/dichloromethane with 1% NH3, 0~15%) to afford a red foam after

concentration. The red foam was further washed with a 1:1 mixture of petroleum ether/EtOAc

(20 mL) and recrystallized from a mixture of hexane/EtOAc to afford the desired compound

trans-4-(2-((2-ethylbutyl)amino)-5-(4-((4-methylpiperazin-1-yl)methyl)phenyl)-7H-pyrrolo[2,3-

d]pyrimidin-7-yl)cyclohexan-1-ol as a pale yellow solid (free base). Water (15 mL) and HCl (6.0

M, 30 mL) was gradually added to the solid at 0 oC. The mixture was stirred for 0.5 h and

35

concentrated. The resulting solid was dissolved in a mixture of H2O and acetonitrile (30 mL,

100:1, v/v), and lyophilized to yield UNC3810A (1) as a fine yellow powder (11.0 g, 60%).

Phospho-RTK arrays

Phospho-RTK profiling was performed using the proteome profiler human phospho-RTK

array kit (R&D Systems, ARY001B). Assay was performed according to manufacturer’s

instructions with the addition of 1 mM sodium orthovanadate (Sigma-Aldrich, S6508) to the

lysis buffer.

Trypan blue exclusion assay

PEL cells were plated in increasing doses of UNC3810A at a density of 200,000 cells per

mL for 72 hours. Afterwards, live cells were counted using trypan blue with a hemocytometer.

Saline solution (0.9% NaCl) was used as the vehicle control with a final concentration of 0.1%.

Caspase-3 activity assay

Caspase-3 activity was measured using the ApoAlertTM Caspase-3 Fluorescent Assay Kit

(Takara Bio, 630215). Assay was performed according to manufacturer’s instructions.

CellTiter-Glo (CTG) cell viability assay

Cell viability was measured using the CellTiter-Glo luminescent cell viability assay

(Promega, G7570). Assay was performed according to manufacturer’s instructions. Leniolisib

(HY-17635) and duvelisib (HY-17044) was purchased from MedChem Express.

AQueous One solution cell proliferation assay, MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl-2-(4-sulfophenyl)-2H-tetrazolium) assay

Cell proliferation was measured using the CellTiter 96® AQueous One solution cell

proliferation assay (Promega, G3581). Assay was performed according to manufacturer’s

instructions.

36

Soft-agar colony formation assays (CFA)

Colony formation assays were performed using using 1x RPMI 1640 (Gibco, 31800-105)

media containing a final concentration of 1% methyl cellulose (Sigma-Aldrich, M0512) except

for colony formation in BCBL1 control or shTyro3 cells which were performed instead using

1.4% methyl cellulose. The cells (and if appropriate, drug inhibitor or vehicle control) was added

to soft-agar medium in Falcon tubes. For experiments regarding Tyro3 knocked-down cell lines,

0.5 µg/ml of puromycin (Corning, 61-385-RA) was added to the soft-agar medium as well. The

tubes were briefly vortexed, and then allowed to sit for at least 10 minutes to allow air bubbles to

rise. Air bubbles were carefully aspirated and the cell suspension was transferred in triplicate to

35 mm cell culture dishes (StemCell Technologies, 27116) using a 5 mL syringe with a 16 gauge

blunt-end needle (StemCell Technologies, 28110). Colonies were visualized after 3 weeks with a

Leica MZ6 light microscope and pictures were taken with an attached Leica MC170 HD camera.

Colonies were quantified using ImageJ v1.50i.

In vitro kinome selectivity and IC50 kinome profiling of UNC3810A

In vitro kinome profiling and determining the IC50 for several kinases to UNC3810A was

performed by Carna Biosciences, Inc. Kinome profiling was performed for UNC3810A at a

concentration of 1.0 µM. Data was graphed using KinMap (147). To determine the IC50 of

several kinases to UNC3810A, concentrations ranging from 0.3 nM to 10 µM were tested to

generate curves. UNC3810A was dissolved in DMSO and then further diluted with assay buffer

[20 mM HEPES, 0.01% Triton X-100, 1 mM DTT (pH 7.5)] to make a 4x test compound

solution. The 4x compound solution was then added to a well with a kinase reaction solution

containing substrate, ATP, and an appropriate metal ligand. After 1 to 5 hours of incubation at

room temperature (dependent on the kinase), termination buffer was added. The ratio of the

37

product to the substrate was quantified. The IC50 was calculated by fitting the points to a four

parameter logistic curve.

Pharmacokinetic analysis (PK) of UNC3810A

PK analysis was completed by Sai Life Sciences Limited. Plasma pharmacokinetics was

determined in 8-12 weeks old male Swiss Albino mice administered with a single dose of 3, 10,

or 25 mg/kg UNC3810A intraperitoneally (i.p.), 3 mg/kg orally (p.o.), or 3 mg/kg intravenously

(i.v.). Blood samples were collected from the retro-orbital plexus of three mice at each time point

over a series of nine time points ranging from pre-dosing to 24 hours after dosing. Plasma

samples were separated by centrifugation at 4000 rpm for 10 minutes at 4oC in a micro

centrifuge tube containing K2EDTA as a coagulant, and protein was precipitated using

acetonitrile. Samples were then analyzed using fit for purpose LC/MS/MS method.

Pharmacokinetic analysis was performed using the non-compartmental analysis tool of Phoenix

WinNonlin v6.3.

Trex-RTA Luc BCBL1 xenograft mouse model

Four to five-week old NOD scid gamma (NSG) female mice were injected with 100,000

Trex-RTA Luc BCBL1 cells intraperitoneally (i.p.). Eight mice were sorted into each arm such

that total luminescent counts were equal between both groups before beginning treatment. Drug

treatment began three days after cell injection. The vehicle used for administering UNC3810A

was saline (0.9% NaCl). Mice were administered with either saline vehicle control or 25 mg/kg

UNC3810A three times a week – Monday, Wednesday, and Friday (MWF). Imaging (prior to

drug treatment) using the Xenogen IVIS-Lumina System (Caliper Life Sciences) was performed

weekly by administering through i.p. injection 10 µl/gbw of luciferin (PerkinElmer, 122799).

Images were analyzed using Living Image v4.4. Mice were harvested when their body score

38

index or tumor burden reached IACUC limits. All mice were harvested when either group

reached IACUC limits.

BJAB xenograft mouse model

Four to five-week old NOD scid gamma (NSG) female mice were injected with 7.5 x 106

BJAB cells subcutaneously. Tumor volume was measured MWF using calipers. Drug treatment

began when tumors reached palpable size. Six mice were sorted into each arm (in an unblinded

fashion) such that average tumor volume was equal between both groups before beginning

treatment. The vehicle used for administering UNC3810A was saline (0.9% NaCl). Mice were

administered either with either saline vehicle control or 25 mg/kg UNC3810A intraperitoneally

(i.p.) three times a week (MWF) until tumor volumes reached IACUC limits of 2000 mm3.

Immunohistochemistry (IHC)

Solid tumors were fixed in 10% formalin containers (Fisher Scientific, 23-032059) for 3

days from which 4 µM paraffin-embedded sections were prepared on slides. Slides were

deparaffinized using Histochoice Clearing Agent (Sigma-Aldrich, H2779) and rehydrated

through an ethanol gradient (100%, 95%, and 70%). Antigen retrieval was performed using the

Agilent Dako PT Link with citrate buffer pH 6.0 (Sigma-Aldrich, W302600). Slides were

washed afterwards with EnVision FLEX wash buffer (Agilent, K8007) and distilled H2O before

being incubated in a 3% hydrogen peroxide and 10% methanol solution to quench endogenous

peroxidases. Sections were incubated in primary antibodies from Cell Signaling Technology

either against p-AKT T308 (1:100, 13038) or p-S6 (1:1000, 4858) in antibody diluent (Cell

Signaling, 8112) overnight. The following day after washing, Rabbit SignalStain Boost IHC

Detection Reagent (Cell Signaling, 8114) was added for 30 minutes. After washing, sections

were developed using DAB (3,3′-diaminobenzidine) (Vector Laboratories, SK-4100) and then

39

counter-stained with hematoxylin (Vector Laboratories, H-3404). After washing and dehydration

through an alcohol gradient, slides were mounted with Cytoseal XYL (Thermo Scientific, 8312-

4). Slides were imaged using a Leica microscope (DMLS) and camera (DMC2900), and acquired

using the Leica Application Suite v4.8 software.

CompuSyn analysis

To determine whether the drug combination of UNC3810A with leniolisib or duvelisib

was synergistic, additive, or antagonistic, the combination index was calculated at the 0.5 effect

level (Fa = 0.5) using CompuSyn (http://www.combosyn.com/) (148). The program was run as

directed according to the manual on the website.

Statistical analysis

A one-way ANOVA with a Dunnett’s multiple comparisons test for adjustment was used

to determine statistical significance (P< .05) between MIB binding of cell-cycle related kinases

from primary B-cells compared to B-cell NHL subtypes using GraphPad Prism 7.02. The

reported significance value in Figure 2.2B is the highest P value of all comparisons between

primary B-cells and each of the B-cell NHL subtypes. To test for statistical significance (P < .05)

of a few kinases (Figure 2.2C-F) a one-way ANOVA with a Turkey’s multiple comparisons test

for adjustment was used instead. An unpaired Mann-Whitney test (P<.05) with a two-tailed

distribution was used to determine statistical significance for tumor growth and effusion volume

using GraphPad. An unpaired, multiple t-test using the Holm-Sidak method (P<.05) assuming

same scatter across populations was used to determine significance of control compared to 2 µM

UNC3810A treated cell lines using GraphPad. A one-way, unpaired ANOVA test (P<.05) was

used to determine statistical significance for comparisons in colony formation with increasing

concentrations of UNC3810A using GraphPad. Colony formation in Tyro3 knocked-down cells

40

was analyzed instead with an unpaired, two-tailed student’s t-test (P<.05) using GraphPad. An

ANOVA test was used to determine statistical significance in gene expression data between

primary B-cells and effusions from patients with PEL using R.

Data Availability

The RNA-seq data reported in this study is deposited in the GEO database under the

accession number GSE114791. The mass spectrometry proteomics data from the MIB/MS

profiling is deposited to the ProteomeXchange Consortium by the PRIDE (149) partner

repository with the dataset identifier PXD012087.

Results

The functional cellular kinome is more similar within a NHL subtype than across subtypes.

To identify potential vulnerabilities within subtypes of lymphoma, we performed

Multiplexed Inhibitor Bead-mass spectrometry (MIB/MS) kinome profiling on 24 individual cell

lines in biological triplicate as well as in primary B-cells from healthy donors (53). The cellular

lysate was run through a column containing beads attached with different kinase inhibitors,

which bind and capture functionally expressed kinases. The bound kinases are eluted off, and

then identified and quantified using mass spectrometry. Replicate MIB/MS runs were consistent,

and were averaged and compared to freshly harvested primary B-cells from five donors (Figure

2.1). From all MIB/MS experiments combined, 386 kinases were identified in at least one cell

line, and 151 kinases were commonly captured from all cell lines. Kinase names are derived

from the Uniprot naming nomenclature. Euclidian hierarchical clustering of all MS data

demonstrated that kinome profiles of lymphoma cell lines generally clustered them according to

subtype, with the primary effusion lymphoma (PEL) subtype being the most distinct (Figure

2.2A). Though FL is generally indolent, it can progress, and in some cases transform into

41

Figure 2.1 Replicates between MIB/MS runs are consistent with each other. Euclidian hierarchical clustering of lymphoma cell lines from four subtypes that were subject to MIB/MS. Individual replicates are presented for most DLBCL and mantle cell lymphoma cell lines, while the average of two replicates are shown for BJAB, Burkitt’s and follicular lymphoma cell lines. Kinases and their corresponding family are listed below the heatmap. Rep indicates replicate; avg, average; min, minimum; and max, maximum.

DLBCL (150, 151). Notably, there were several cell lines (SuDHL4, Karpas422, and WSU-

NHL), which have been characterized as either FL or DLBCL (152-157). Since WSU-NHL

clustered with the other DLBCLs, and SuDHL4 and Karpas422 with the other FL, we

categorized them as such respectively (Figure 2.2A). Three kinases were captured in all cancer

cell lines but were absent from primary B-cells: STK6 (Aurora kinase A), NEK2, and MK06

(ERK3) (Figure 2.2B). Other kinases with greater MIB-binding in NHL compared to primary B-

cells were associated with cell cycle regulation (CDK1, CHK1, PLK1, PMYT1, and AURKB)

42

Figure 2.2 The cellular kinome is more similar within subtypes than across subtypes. (A) Euclidian hierarchical clustering of 24 lymphoma cell lines from five subtypes and primary B-cells that were subject to MIB/MS. Data presented is the average of two biological replicate MIB/MS runs for each cell line and average of five replicate MIB/MS runs for primary B cells. Gray color indicates a kinase that was not captured in that cell line. (B) A select set of kinases having greater MIB-binding in all cell lines relative to primary B cells are graphed. Points within each lymphoma subtype represent a single cell line. Points for primary B-cells are from individual mass spectrometry runs. Data presented is the mean ± SEM with significance (ANOVA, *P<.05, **P<.01, ***P<.001, df=24) tested between primary B-cells and each NHL subtype. Reported significance is the highest P value of all comparisons between each NHL subtype to the primary B-cells. (C-F) A select set of kinases: (C) KS6B1 (D) ATR (E) LIMK2 and (F) PKN1 displaying MIB-binding bias across subtypes. Each point represents a single cell line. Data presented is the mean ± SEM (ANOVA, *P<.05, **P<.01, ***P<.001, df=19). LFQ indicates label free quantification.

(Figure 2.2B). Comparing subtypes to one another, some kinases displayed a bias towards having

greater or less MIB-binding in a particular subtype versus others. For example, KS6B1 (p70

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S6K) showed the greatest MIB-binding in PEL (Figure 2.2C). Conversely, ATR displayed less

MIB-binding in MCL and DLBCL versus other subtypes (Figure 2.2D). LIMK2 also displayed

the greatest MIB-binding in DLBCL (Figure 2.2E), while PKN1 was captured the least in the

follicular subtype (Figure 2.2F). Euclidean hierarchical clustering analysis of kinases commonly

activated in different NHL subtypes compared to primary B-cells demonstrate PEL in general

cluster together, while having lower functional MIB binding of RIOK2 and CHKA, but higher

MIB binding of CDK16 compared to other NHL subtypes (Figure 2.3).

Figure 2.3 A common set of kinases are activated within NHL lymphomas compared to primary B-cells. Euclidian hierarchical clustering of kinases having statistically higher or lower MIB-binding in NHL as compared to primary B-cells. Data presented is the average of two biological replicate MIB/MS runs for each cell line, while the data for primary B-cells are MIB/MS runs pooled from different donors. Min indicates minimum; and max, maximum.

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Figure 2.4 Tyro3 is uniquely upregulated in PEL. (A) Euclidian hierarchical clustering of kinases having statistically higher or lower MIB-binding in PEL as compared to all other samples combined. Data presented is average of two biological replicate MIB/MS runs for each cell line and an average of five replicate MIB/MS runs for primary B-cells. Orange asterisk indicates the row containing Tyro3. (B) Tyro3 MIB-binding levels in each individual subtype. Each point represents one cell line. Data presented is the mean ± SEM. (C-E) mRNA expression

45

of (C) TYRO3 (D) AXL or (E) MERTK from RNA-seq in each individual subtype. Each point represents one cell line. Data presented is the mean ± SEM. (F) Western blot analysis of Tyro3 protein levels from different NHL subtypes. Representative of n = 3 independent experiments. (G,H) RT-qPCR analysis of (G) PROS1 or (H) GAS6 mRNA from PEL cell lines or primary B-cells isolated from three different donors. Graph is representative of n=3 independent experiments with different primary B-cell donors used for each experiment. Bar graph represents the mean ± SD (n=3). (I) RT-qPCR analysis of TYRO3, PROS1, or GAS6 from primary B-cells isolated from healthy donors (n=7) or effusion from patients with PEL (n=6). Replicates are shown as the same color for each primary B-cell donor or PEL effusion sample. Data points are presented as the delta CT value between the gene of interest and actin for each individual replicate. Box plot represents the median with the 25th and 75th percentiles, respectively (ANOVA, P value listed below each graph). Lymph indicates lymphoma and LFQ indicates label free quantification.

Tyro3 is uniquely upregulated in PEL

Large clusters of kinases in the PEL subtype displayed significantly greater or less MIB-

binding compared to all other NHL subtypes and primary B-cells (Figure 2.4A). Kinases

significantly downregulated in PEL include BTK, GSK3B, and TGFR1. This corresponds well to

a previous report that PEL is insensitive to BTK inhibition unlike DLBCL (158). Kinases

upregulated in PEL include KS6A1 (p90 S6K), KS6B1, and AKT2. KS6B1 and AKT2 are part

of the phosphoinositide 3-kinase (PI3K)/protein kinase B (AKT)/mammalian target of rapamycin

(mTOR) pathway, which has been shown to be important for PEL survival (6, 73, 159).

Interestingly, Tyro3 was consistently captured by MIBs from all six PEL cell lines but absent

from all other cell lines and primary B-cells (Figure 2.4B). Tyro3 is a member of the

Tyro3/Axl/MerTK (TAM) receptor tyrosine kinase (RTK) family, whose overexpression

promotes proliferation, survival, and chemoresistance in solid cancers such as in melanoma,

hepatocellular carcinoma, and colon cancer (146, 160-162). MerTK, but not Axl or Tyro3, has

also been shown to promote proliferation in multiple myeloma, which is a malignancy of

plasmacytic origin (163). Transcriptional profiling by RNA-sequencing (seq) demonstrated

TYRO3 was uniquely transcriptionally upregulated in PEL compared to all other subtypes

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Figure 2.5 Trends in expression data are similar to a previous literature report. (A,B) Expression of TAM receptors and their ligands as determined from (A) our and (B) Journo et. al., 2018 RNA-seq analysis (142). TPM indicates transcripts per million.

(Figure 2.4C). In contrast, its related family members AXL and MERTK displayed a similar low

level of expression across subtypes (Figure 2.4D, E). To confirm our expression data, we also

examined expression levels of the TAM receptors and their ligands in a recent literature report

by Journo et. al. (142) that performed RNA-seq on BJAB, BCBL1, and BC3 cell lines. Trends in

expression levels of these genes between cell lines were similar in both studies (Figure 2.5). PEL

lines also exhibited high levels of Tyro3 protein expression in comparison to some other non-

PEL NHL, which showed little to none (Figure 2.4F). BC3, which had the lowest protein and

mRNA levels of Tyro3 within the PEL subtype, nevertheless had greater MIB-binding/activity in

comparison to the cell lines from other subtypes. TAM receptors achieve maximal signaling

when bound by the ligands Pros1 and/or Gas6 (164). Therefore, we examined if PEL also exhibit

upregulated expression of Pros1 or Gas6. PEL in general exhibited higher transcription levels of

PROS1, but not GAS6 when compared to primary B cells (Figure 2.4G, H). To verify that the

transcriptome obtained from PEL cell lines reflected the primary tumor, we harvested RNA from

primary PEL patient samples to confirm if upregulation of TYRO3 transcripts held true in

primary human samples. We observed significantly increased levels of TYRO3, PROS1, and

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GAS6 mRNA transcripts in the PEL effusion (n=6) in comparison to healthy primary B cells

(n=7) (Figure 2.4I). Overall, our data demonstrate that Tyro3 and its activating ligand Pros1 are

upregulated in PEL cells.

Figure 2.6 Inhibition of Tyro3 results in attenuation of PI3K signaling. (A) Phospho-receptor tyrosine kinase (RTK) assay examining tyrosine phosphorylation of different RTKs. Reference spots are located at all corners except for the bottom left. (B) Western blot analysis of Burkitt's lymphoma or PEL cell lines stably knocked down with non-template control or Tyro3 shRNA. All cell lines were run on the same gel, but the blot was split in half for clarity due to different exposure times required to detect PI3K in Burkitt's compared to PEL cell lines. The split occurs on the right-side of BL41 and to the left-side of BCBL1. The red asterisk indicates the location of a non-specific band and the red arrow indicates the p85 PI3K band. Representative of n = 3 independent experiments. (C) Structure of UNC3810A with IC50 values to the TAM family receptors. (D) Representative blot (n = 2 independent experiments) of a phospho-RTK assay with BCBL1 cells treated for 24 hours with increasing concentrations of UNC3810A. Reference spots are located at all corners except for the bottom left. DMSO was used as the vehicle control. (E) Western blot analysis of the PI3K/Akt/mTOR signaling pathway

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in BCBL1 cells treated with increasing concentrations of UNC3810A for 24 hours. DMSO was used as the vehicle control. Representative of n = 3 independent experiments.

Inhibition of Tyro3 attenuates PI3K signaling

During activation of an RTK, phosphorylation of tyrosine residues within the kinase

domain results in propagation of a signaling cascade. We examined if Tyro3 was phosphorylated

on its tyrosine residues, which would indicate the activation of Tyro3. Phospho-RTK arrays,

which contain antibodies against a panel of RTKs including Tyro3, confirmed that Tyro3 was

phosphorylated on tyrosine residues in PEL cells (BC1 and BCP1) in comparison to a DLBCL

(BJAB), which exhibited none (Figure 2.6A). In contrast one kinase, ROR2, was commonly

phosphorylated across all three cell lines. To confirm whether or not Tyro3 expression affects the

activity of a downstream substrate in PEL cells, we examined if phosphorylation of the p85

regulatory subunit of PI3K, a known substrate of Tyro3, was affected upon knockdown of Tyro3

(165). We tested several short-hairpin (sh) RNA constructs against Tyro3 in PEL (BCBL1 and

BC1) and DLBCL BJAB cells (Figure 2.7). Since shRNA clone #1527 negatively affected cell

viability in PEL BCBL1 cells (Figure 2.7B), but had similar cell viability compared to the non-

template control in DLBCL BJAB cells (Figure 2.7D), we proceeded with shRNA clone #1527

for further studies. Knockdown of Tyro3 in PEL, but not in BL, resulted in a decrease in tyrosine

phosphorylation of p85 PI3K, consistent with a decrease in PI3K activity (Figure 2.6B).

UNC3810A is a selective kinase inhibitor against Tyro3

The inclusion of TAM inhibitors to conventional therapies to allow chemotherapy to have

a durable response may be helpful in treating cancers that often develop resistance to treatment.

Currently there are several small molecule inhibitors that target the TAM receptors, but many of

the compounds are pan-inhibitors of the Tyro3/Axl/MerTK family and frequently have a broad

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Figure 2.7 Tyro3 knockdown attenuates survival and signaling in PEL cells. (A,B) PEL BCBL1, (C,D) non-PEL BJAB, or (E) PEL BC1 cells were infected with lentivirus containing different constructs of shRNA against Tyro3 for 72 hours. The shRNAs 2178 and 2179 contain a pLKO.1 backbone, while the rest of the Tyro3 shRNAs contain a pLKO.5 backbone. Cells infected with the pLKO.1 control shRNA and the pLKO.5 control shRNA are denoted as NTC 1 or NTC 5, respectively. (A,C,E) Western blot analysis of (A) PEL BCBL1, (C) non-PEL BJAB or (E) PEL BC1 cells knocked down with non-template control or different Tyro3 shRNA constructs. BC1 and HUVEC are positive controls expressing Tyro3, or Axl and MerTK, respectively. (B,D) Cell-Titer Glo analysis of (B) BCBL1 or (D) BJAB cells knocked down with

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non-template control or Tyro3 shRNA in serum-starved conditions over a course of 72 hours. Luminescence was normalized to the reading at 0 hours for each condition. Data is presented as mean ± SD (n=3 independent wells) and is representative of n=3 independent experiments.

spectrum that includes MET and other RTKs (77, 166). We created a small molecule inhibitor

with specificity towards Tyro3. Structure-activity studies of our MerTK inhibitors versus the

TAM family led to UNC3810A (1), a pyrrolopyrimidine-based specific small molecule inhibitor

against Tyro3 (Figure 2.6C and Figure 2.8) (167). To determine the specificity of UNC3810A

for Tyro3, we performed in vitro kinome selectivity profiling of UNC3810A against a wide array

of kinases at 1 µM (Figure 2.9A) and determined the IC50 of UNC3810A versus the TAM

kinases, FLT3, and several additional targets that were inhibited by >80% at 1 µM (Table 2.2).

Figure 2.8 Synthesis of UNC3810A. Starting material (denoted as 3) is trans-4-(5-bromo-2-chloro-7H-pyrrolo[2,3-d]pyrimidin-7-yl)cyclohexan-1-ol and the intermediary product (denoted as 4) is trans-4-(5-bromo-2-((2-ethylbutyl)amino)-7H-pyrrolo[2,3-d]pyrimidin-7-yl)cyclohexan-1-ol. NMR and MS (ESI) data for (4): 1H NMR (400 MHz, CD3OD) δ 8.26 (s, 1H), 7.11 (s, 1H), 4.52 – 4.42 (m, 1H), 3.69 – 3.60 (m, 1H), 3.34 (d, J = 6.3 Hz, 2H), 2.11 – 2.01 (m, 2H), 2.01 – 1.82 (m, 4H), 1.58 – 1.34 (m, 7H), 0.95 (t, J = 7.4 Hz, 6H); 13C NMR (100 MHz, CD3OD) δ 160.89, 153.23, 150.12, 123.14, 112.18, 88.91, 70.18, 54.17, 45.20, 42.28, 35.33, 31.40, 25.05, 11.57; MS (ESI) for [M+H]+ (C18H28BrN4O+): calculated m/z 395.14; found m/z 395.15; LC-MS: >95% purity. NMR and MS (ESI) data for UNC3810A (1): 1H NMR (400 MHz, CD3OD) δ 8.85 (s, 1H), 7.97 (s, 1H), 7.82 – 7.72 (m, 4H), 4.67 – 4.58 (m, 1H), 4.55 (s, 2H), 3.91 – 3.59 (m, 9H), 3.51 (d, J = 6.3 Hz, 2H), 3.02 (s, 3H), 2.19 – 2.02 (m, 6H), 1.70 – 1.60 (m, 1H), 1.59 – 1.40 (m, 6H), 1.01 (t, J = 7.4 Hz, 6H); 13C NMR (100 MHz, CD3OD) δ 155.95, 152.66, 140.22, 135.44, 133.59 (3C), 128.46 (3C), 118.17, 111.25, 69.90 (2C), 60.84, 55.23, 51.30 (2C), 49.14 (overlapped), 45.16, 43.33, 42.07, 35.15 (2C), 30.94 (2C), 24.94 (2C), 11.47 (2C); MS (ESI) for [M+H]+ (C30H45N6O+): calculated m/z 505.36; found m/z 505.40; LC-MS: >95% purity. K2CO3 indicates potassium carbonate; DIPEA, N,N-diisopropylethylamine; PrOH, isopropyl alcohol; and Pd(PPh3)4, tetrakis(triphenylphosphine)palladium.

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Figure 2.9 Specificity of UNC3810A to Tyro3. (A) Kinome tree graphed using KinMapbeta (kinhub.org/kinmap/) depicting targets of UNC3810A at 1 µM. Percentage of inhibition represents the ratio of signal conversion of the substrate by the kinase with or without UNC3810A present. Positive or negative inhibitions refer to a decrease or increase in signal, respectively, upon addition of UNC3810A compared to the control. Refer to Table 2.2 for IC50 values of UNC3810A against several kinases. (B-D) Cell-Titer Glo analysis of non-PEL or PEL cell lines treated with (B-D) UNC3810A or (D) UNC2369A, a negative control of UNC3810A, for (B,C) 72 hours or (D) 48 hours. Luminescence was normalized to saline control. Data is presented as mean ± SD (n=3 independent wells) and is representative of n=3 independent experiments. Non-PEL lines are BJAB, Ramos, and BL41, while PEL lines are BCBL1, BC1, BC3, VG1, JSC1, and BCP1. IC50 values are (B) BC3 – 2.13 µM, VG1 – 2.78 µM, JSC1 – 1.36 µM, and BCP1 – 2.20 µM, (C) BJAB - 3.23 µM [3.16, 3.30], Ramos – 4.77 µM [4.53, 5.03], BL41 – 3.20 µM [3.13, 3.27], BCBL1 – 0.93 µM [0.76, 1.13], and BC1 – 1.57 µM [1.46, 1.70], and (D) BJAB 3810A – 5.32 µM [5.10, 5.54], BJAB 2369A – 5.65 µM [5.29, 6.04], BCBL1 3810A – 1.04 µM [0.61, 1.77], and BCBL1 2369A – 3.07 µM [2.73, 3.46] with the upper and lower limits of the 95% confidence interval presented in brackets. (E) Western blot analysis of Tyro3 expression. Representative of n=2 independent experiments. (F) Cell-Titer Glo analysis of cell lines with differing expression levels of Tyro3 treated with UNC3810A. Luminescence was normalized to saline control. Data is presented as mean ± SD (n=3 independent wells) and is representative of n=2 independent experiments. IC50 values are BJAB – 4.75 µM, BCBL1 – 1.23 µM, HT-29 – 1.83 µM, and B16-F10 – 1.44 µM. (G) Western blot analysis of caspase-3 activation in PEL cells treated with UNC3810A for 72 hours. Representative of n=2 independent experiments. AGC is a kinase group containing PKA, PKG, PKC family kinases; and CMGC contains CDK, MAPK, GSK3, CLK family kinases. CAMK indicates calcium/calmodulin-dependent protein kinase; CK1, casein kinase 1; STE, homologs of yeast Sterile 7, Sterile 11, Sterile 20 kinases; TK, tyrosine kinase; TKL, tyrosine-kinase like.

UNC3810A was most potent against Tyro3 (1.2 nM) and showed at least 10-fold selectivity

versus all other kinases beyond MerTK, the most closely related TAM member, and TRKA.

When examining the sensitivity of PEL cells to UNC3810A, most PEL lines exhibit an IC50

between 0.9 – 2.8 µM (Figure 2.9B, C). Furthermore, PEL cell lines (BCBL1 and BC1)

exhibited more sensitivity to UNC3810A in comparison to non-PEL cell lines (BJAB, Ramos,

and BL41), which do not highly express Tyro3 (Figure 2.9C). Comparison of UNC3810A to a

structurally related analogue, which does not target Tyro3, UNC2369A, revealed that BCBL1

cells are more sensitive to UNC3810A compared to UNC2369A (Figure 2.9D) (167).

UNC2369A is another pyrrolopyrimidine-based compound that is structurally similar to

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UNC3810A with an IC50 of 11 µM against Tyro3, 3.9 µM against Axl, and 400 nM against

MerTK (see compound 20 from Zhang et. al., 2014 (167)). To ensure these effects are not cell-

Table 2.2 IC50 values of UNC3810A against top kinase hits from the kinome profiling. The IC50 to UNC3810A was determined for the top 10 kinase hits from the kinome profiling, the TAM receptors, and FLT3. Fold selectivity of Tyro3 versus other kinases to UNC3810A is listed in the last column.

Kinase IC50

(nM) Fold selectivity TYRO3

TYRO3 1.2 1 MERTK 7.3 6.1 TRKA 11.4 9.5 AXL 23.2 19.3

TRKC 33.1 27.6 FLT3 34.3 28.6

MELK 41.5 34.6 DDR1 61.1 50.9 TRKB 62.0 51.7 YES 129.5 107.9 IRR 131.8 109.8

DDR2 209.8 174.8 MAP4K2 216.9 180.8

SIK 229.3 191.1

type specific, we also examined the sensitivity of a colon cancer cell line, HT-29, and a

melanoma cell line, B16-F10 to UNC3810A. Both colon cancer and melanoma have previously

been shown to upregulate Tyro3 to promote cell proliferation and chemoresistance (160, 162,

168, 169) and we confirmed HT-29 and B16-F10 express Tyro3 (Figure 2.9E). The sensitivity of

both HT-29 and B16-F10 to UNC3810A was more similar to BCBL1 cells compared to BJAB

cells (Figure 2.9F). To examine if UNC3810A also attenuated Tyro3-mediated PI3K signaling,

we treated BCBL1 PEL cells with increasing amounts of UNC3810A. This resulted in a dose-

dependent decrease of Tyro3 phosphorylation (Figure 2.6D) and attenuated signaling through the

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PI3K/AKT/mTOR pathway as evidenced by decreased phosphorylation of p85 PI3K and AKT

(Figure 2.6E). Overall, these results suggest that UNC3810A is a selective inhibitor of Tyro3.

Inhibition of Tyro3 results in decreased cell survival

Tyro3 can activate the PI3K/AKT/mTOR pathway, whose inhibition in PEL cells results

in cell death (6, 73, 159). If Tyro3 contributed to the activation of the PI3K/AKT/mTOR

pathway, then reducing Tyro3 should affect PEL viability. We found that the PEL lines (BCBL1

and BC1) exhibited reduced cell viability in serum-starved conditions upon lentivirus-mediated

shRNA knockdown of Tyro3 compared to cells transduced with a non-targeting control (NTC)

shRNA (Figure 2.10A and Figure 2.7B). These results suggest Tyro3 activates PI3K signaling to

promote survival of PEL cells. Indicative of the lack of functionality of Tyro3 in non-PEL lines,

knockdown of Tyro3 in two non-PEL cell lines, BJAB and BL41, had little to no effect on cell

viability under the same conditions (Figure 2.10A and Figure 2.7D). BCBL1 and BC1 PEL cells

targeted with NTC shRNA or Tyro3 shRNA were also subjected to a colony formation assay,

and the number of colonies formed by the Tyro3-depleted cells was significantly less than

control cells (Figure 2.10B). Similarly, treatment of PEL cell lines (BCBL1, BC1) with

UNC3810A also resulted in a significant attenuation in the formation of colonies for the PEL

lines at 0.5 µM and 1 µM compared to the non-PEL NHL cell lines (BJAB, Ramos, BL41),

which exhibited none (Figure 2.10C, D).

Since PI3K and AKT phosphorylation levels decreased upon UNC3810A treatment, we

examined if the PEL lines were more sensitive to UNC3810A compared to the other NHL

subtypes. After treating cells for 72 hours with 2 µM of UNC3810A or DMSO control, we

measured cell growth using an MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl-

2-(4-sulfophenyl)-2H-tetrazolium) assay. PEL lines overall showed decreased cell growth

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Figure 2.10 Inhibition of Tyro3 results in a decrease of cell survival and colony formation in PEL. (A) Cell-Titer Glo analysis of non-PEL NHL or PEL cell lines stably knocked down with non-template control or Tyro3 shRNA over a time-course of 72 hours in serum-starved

56

conditions. Luminescence was normalized to the reading at 0 hours for each condition. (B) PEL cells were assessed for colony formation after being knocked down with non-template control or Tyro3 shRNA in a lentivirus-mediated manner. (C,D) Colony formation of (C) PEL (BCBL1 and BC1) or (D) non-PEL NHL (BJAB, Ramos, or BL41) cells was assessed over increasing concentrations of UNC3810A. Saline was used as the vehicle control. (B-D) Representative microscope pictures are shown capturing the full diameter of the tissue culture plate (length-wise) and also an area enlarged at the middle. The diameter of each plate is 35 mm in length. Significance is calculated using a one-way ANOVA test (*P<.05, **P<.01, ***P<.001, df = 8). Pictures were taken with a Leica MZ6 light microscope attached with an Leica MC170 HD camera. (A-D) All quantitative data is presented as mean ± SD (n=3 independent samples) and all experiments are representative of n=3 independent experiments.

compared to all other NHL subtypes (Figure 2.11A). We also examined if this inhibition of

cellular proliferation in PEL cells is cytostatic or cytotoxic. After treatment of PEL cells with

UNC3810A for 72 hours, we found that cell viability decreased with increasing concentrations of

UNC3810A (Figure 2.11B). Apoptotic cell death can be mediated by active caspase-3, which

cleaves many proteins upon activation such as PARP. To determine if cell death occurred in a

caspase-3 mediated manner, PEL cells were treated with UNC3810A for 72 hours. A dose-

dependent increase in caspase-3 activity was observed and corresponded with increased levels of

cleaved caspase-3 (the active form of caspase-3) and PARP (Figure 2.11C, D and Figure 2.9G).

We also examined if treating PEL cells with UNC3810A in combination with other inhibitors

resulted in a further decrease in cell viability. Several studies have shown that the TAM receptors

promote chemoresistance and that inhibition of the receptor(s) results in increased sensitivity to

chemotherapy (160, 162, 170, 171). Hence, we investigated the efficacy of combined treatment

of Tyro3 with two PI3K inhibitors, leniolisib and duvelisib, against PEL. We treated PEL lines

with either UNC3810A, leniolisib, or together in combination for 72 hours and measured cell

viability. Treatment of PEL cells with both resulted in lower cell viability at multiple

concentrations compared to treatment with either UNC3810A or leniolisib alone (Figure 2.12A).

We also examined if treatment of PEL cells with another PI3K inhibitor, duvelisib, and

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Figure 2.11 UNC3810A treatment of PEL cells results in cell death. (A) Growth was measured using CellTiter 96 AQueous One solution from different NHL cell lines that were treated with 2 µM UNC3810A for 72 hours. Absorbance readings in each cell line were normalized to the DMSO control from their respective cell line. Data is presented as mean ± SD (n=3 independent wells) and is representative of n=3 independent experiments (unpaired multiple t-test by Holm-Sidak method, *P<.05, **P<.01, ***P<.001, df =4). (B) PEL (BCBL1 and BC1) cells were treated with increasing concentrations of UNC3810A and harvested after 72 hours of treatment. Cells were counted for viability using trypan blue. Saline was used as the vehicle control. Data is presented as mean ± SD (n=3 independent samples, 4 counts from different 1 mm2 areas on the hemocytometer per sample) and is representative of n=3 independent experiments. (C) Western blot analysis of caspase-3 activation in BC1 cells treated for 72 hours with UNC3810A. Saline was used as the vehicle control. (D) Caspase-3 activity was measured in PEL (BCBL1 and BC1) cells treated with UNC3810A for 72 hours using the ApoAlertTM fluorescent assay kit. Saline was used as the vehicle control. Data is presented as mean ± SD (n=3 independent samples) and is representative of n=3 independent experiments.

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Figure 2.12 UNC3810A sensitizes PEL cells to treatment with PI3K inhibitors. (A) PEL cells were treated for 72 hours with UNC3810A alone, leniolisib alone, or together in combination at the indicated concentrations. (B) PEL cells were treated for 72 hours with UNC3810A alone, duvelisib alone, or together in combination at the indicated concentrations. (A,B) Cell viability was measured using Cell-Titer Glo. Luminescence readings were normalized to the vehicle control (UNC3810A – saline, leniolisib and duvelisib – DMSO, and for combinations – the corresponding mix of vehicle solutions used for the inhibitors in the combinatorial treatment). Data is presented as mean ± SD (n=3 independent wells) and is representative of n=3 independent experiments. (C) The combination index (CI) of the drug combinations from (A-B) calculated from CompuSyn at an effect level of 0.5 (Fa = 0.5). CI > 1 indicates antagonism, CI = 1 is an additive effect, and CI < 1 indicates synergism (148). Combo indicates combination.

UNC3810A in combination would have a similar effect. We saw there was a larger decrease in

cell viability when treating with both inhibitors compared to either alone (Figure 2.12B). We also

examined if either of these PI3K inhibitors were synergistic with UNC3810A. Using CompuSyn,

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we calculated the combination index (CI) at an effect level of 0.5 (Fa = 0.5) (148). In PEL cells,

both drugs were synergistic (CI < 1) with UNC3810A though leniolisib was less synergistic with

UNC3810A in comparison to duvelisib (Figure 2.12C). These results suggest that addition of

UNC3810A to this chemotherapeutic regimen may be beneficial.

Figure 2.13 Pharmacokinetic analysis of UNC3810A. (A,C) Mice were administered 3 mg/kg UNC3810A intravenously (i.v.), intraperitoneally (i.p.), or orally (p.o.), and plasma concentration levels of UNC3810A were determined over a course of 24 hours. Data presented is the mean ± SD (n=3 mice). (B,D) Mice were administered 3, 10, or 25 mg/kg of UNC3810A intraperitoneally (i.p.), and plasma concentration levels were determined over a course of 24 hours. Data presented is the mean ± SD (n=3 mice). (C,D) Parameters of analysis are presented: Tmax(h) is the time point which maximal concentration was achieved; T1/2(h) is the half-life; %F is oral bioavailability; Cmax is the maximum serum concentration achieved; AUClast is the area under the curve calculated up to the last measurable concentration; and AUCinf is the calculated area under the curve when extrapolating the time to infinity.

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Figure 2.14 UNC3810A retards PEL tumor growth in vivo. NOD-SCID mice in a BCBL1-PEL xenograft model were i.p. injected with 0.9% saline (control) or 25 mg/kg UNC3810A three times a week – Monday, Wednesday, and Friday (MWF) over a course of four weeks (n=8 mice per group). Luciferase expression in saline or UNC3810A treated mice was measured weekly by i.p. injecting mice with luciferin. (A) Luminescence images representing tumor burden taken at a one-minute exposure at 15 days after start of treatment. Luminescence images depict photon emission from tissue and are expressed as radiance (photons/second/centimeter squared/steradian). (B) Luminescence images were quantified using Living Image v4.4 to measure photon emission from tissue and are expressed as radiance (photons/second/centimeter squared/steradian). Data is presented as a box and whiskers plot with the whiskers marking the minimum and maximum values. The lowest, middle, and top lines of the box respectively represent the 25th percentile, median, and 75th percentile. (C) Effusions collected from mice treated with saline or UNC3810A were measured at the end of the study. Data represents mean ± SD (n=8 mice per group). (D) Western blot analysis of solid tumors from mice treated with saline or UNC3810A (n = 7 per group). (E) Representative pictures of immunohistochemistry (IHC) stains against p-AKT (Thr308) and p-S6 (Ser235/Ser236) of mice treated with saline or UNC3810A. Sections were developed with DAB (3,3′-diaminobenzidine) and counterstained with hematoxylin. Scale bar at lower left is 50 µm. Slides were imaged using a Leica microscope (DMLS) and camera (DMC2900), and acquired using the Leica Application Suite v4.8 software. For (B,C) a two-tailed unpaired Mann-Whitney test was used to determine significance (*P<.05, **P<.01, ***P<.001).

UNC3810A retards PEL tumor growth in vivo

To determine if UNC3810A was efficacious in vivo, we examined if UNC3810A can

attenuate tumor growth in a PEL xenograft mouse model. Using Trex-RTA BCBL1 cells, we

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introduced a constitutively active red-shifted luciferase (luc) gene thereby allowing us to monitor

tumor growth upon addition of luciferin in the mice (137). To examine if UNC3810A would

reach sufficient sustained plasma level concentrations, we performed a pharmacokinetic analysis

of UNC3810A administered through an intravenous (I.V.), intraperitoneal (I.P.) or oral route

(Figure 2.13). Detectable plasma levels were sustained for 12 hours when UNC3810A was

administered through an I.P. route. At 25 mg/kg UNC3810A plasma concentration levels

reached a maximum concentration of 5.72 µM (Figure 2.13D). After four weeks of treatment at

25 mg/kg, tumor growth in mice treated with UNC3810A was significantly suppressed in

comparison to the saline vehicle control group (Figure 2.14A, B). Control mice also had a

significant amount of ascites (pleural effusion) collected from the peritoneal cavity compared to

those treated with UNC3810A (Figure 2.14C). Conversely in a BJAB xenograft mouse model on

the same dosing regimen, there was no attenuation in tumor volume between mice treated with

25 mg/kg UNC3810A and the saline vehicle control (Figure 2.15). This correlated with the data

Figure 2.15 UNC3810A treatment does not reduce tumor burden in a non-PEL lymphoma xenograft model. NOD-SCID mice in a BJAB-xenograft model were administered 0.9% saline (vehicle control) or 25 mg/kg UNC3810A intraperitoneally (i.p.) three times a week – Monday, Wednesday, and Friday (MWF) - over a course of three weeks (n = 6 mice per group). Tumor volume was measured MWF using calipers. Data is presented as a box and whiskers plot with the whiskers marking the minimum and maximum values. The lowest, middle, and top lines of the box respectively represent the 25th percentile, median, and 75th percentile. Significance was tested using a two-tailed unpaired Mann-Whitney test (*P<.05, **P<.01, ***P<.001). No significance was observed on any day.

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from cell culture studies. Immunoblotting of the PEL tumors showed decreased levels of

activating phosphorylation marks of S6 (Ser235/Ser236) in some of the mice treated with

UNC3810A compared to those treated with the saline control (Figure 2.14D). S6 is a ribosomal

protein activated by the mTOR complex 1 (mTORC1) and phosphorylation of serine residues

235/236 are an indicator of active protein translation. Confirming these results,

immunohistochemistry analysis of these solid tumors showed decreased levels of activated Akt

(Thr308) and ribosomal S6 (Ser236/236) in mice treated with UNC3810A compared to mice

treated with the saline control (Figure 2.14E and Figure 2.16). The activation of Akt can promote

cell survival and the activation of S6 can promote protein synthesis. Therefore, tumors in mice

treated with UNC3810A may exhibit delayed tumor progression due to inhibition of Tyro3 and

several survival pathways influenced by Akt.

Discussion

Kinases control the activity of many cellular pathways and are the target of the fastest

growing class of FDA approved oncology drugs (21). Performing MIB-based kinome profiling

allows us to interrogate the kinome using a proteomic-based approach to identify expressed and

activated kinases in the cell (53). Overexpression of a kinase does not necessarily correlate to its

activity (172, 173), therefore characterizing the activity of kinases can identify candidate kinases,

which may not be considered a hit from more traditional genomic and transcriptomic approaches.

In the case of PEL (and other virally associated cancers), expression of viral proteins or

microRNAs can mediate the activation or repression of host kinases in signaling pathways (174,

175).

To identify possible targets of interest, we kinome profiled key NHL subtypes and

identified kinases whose activity was specifically upregulated in one or a few subtypes. The most

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Figure 2.16 Immunohistochemistry analysis of solid tumors from BCBL1-xenograft mice. Solid tumors were harvested at the end of the experiment from BCBL1-xenograft mice treated with either saline control or 25 mg/kg UNC3810A. Paraffin-embedded sections from the tumors were placed on slides, and then probed for either (A) p-AKT threonine 308 or (B) p-S6 serine 235/serine 236. A control slide where the section was not incubated in primary antibody is demarcated by the red line. Sections were developed with DAB (3,3′-diaminobenzidine) and counterstained with hematoxylin. Scale bar at lower left is 50 µm. Slides were imaged using a Leica microscope (DMLS) and camera (DMC2900), and acquired using the Leica Application Suite v4.8 software.

significant hit in the kinome profiling screen for PEL was Tyro3, whose upregulation in other

cancers promotes survival and resistance to chemotherapy (146, 160-162). Tyro3 can activate

several pathways, including the PI3K/AKT/mTOR pathway, whose activity is important for PEL

survival (6, 73, 75, 159). Interestingly in Kaposi’s sarcoma (KS), another KSHV-mediated

malignancy, Axl was upregulated and shown to drive PI3K signaling (79). Knockdown of Axl

resulted in an attenuation of cell growth and invasion in KS cell lines. In normal B-cells,

expression of TAM receptors is very low or absent (170, 171, 176). Of the subtypes screened,

only PEL showed high expression and activity of Tyro3. We then developed a novel Tyro3-

specific inhibitor, UNC3810A. In multiple cell culture assays, signaling and viability was

inhibited by UNC3810A in PEL cell lines expressing Tyro3 while NHL cell lines with low or

absent levels of Tyro3 were not significantly altered. Further testing in xenograft models

demonstrated that treatment of PEL bearing mice with UNC3810A resulted in a lower tumor

burden compared to the saline control that was not seen in a non-PEL NHL model. These data

provide proof-of-principle that targeting Tyro3 may be effective in treating PEL.

Several studies have looked at the efficacy of using inhibitors to the PI3K/Akt/mTOR

pathway in the treatment of PEL (6, 73, 159, 177-179). Since several other NHL subtypes have

been shown to develop resistance to PI3K inhibition (180-182) inhibition of Tyro3 may be more

beneficial than just targeting PI3K as Tyro3 can signal into multiple pathways including the

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MAPK/extracellular-signal-regulated kinase (ERK) (183-186), p38 MAPK (187), Janus kinase

(JAK)/signal transducer of activation (STAT) (164), and phospholipase C (PLC) pathways (164,

186). Both the MAPK/ERK and p38 MAPK pathways are activated in KSHV-infected PEL (24,

48, 49). Furthermore, KSHV also encodes a viral homologue of interleukin-6 (vIL-6), which can

activate the JAK/STAT pathway similar to cellular IL-6 (174). Since Tyro3 signals through

several pathways activated in KSHV-associated malignancies, inhibiting Tyro3 may be more

efficacious than inhibiting PI3K alone. Furthermore, as expression of Tyro3 in other cancers has

been shown to promote resistance to chemotherapy, which is a major obstacle in treating PEL,

addition of a Tyro3 inhibitor may help sensitize cells to chemotherapy (160, 161). Though

preliminary, we see that treatment of PEL cells with UNC3810A in conjunction with the PI3K

inhibitors, leniolisib and duvelisib, inhibited cell viability at higher levels compared to either

treatment alone. Further studies with Tyro3 inhibitors in PEL either as single agent or in the

adjuvant setting are warranted.

In this study, we only focused on the role of Tyro3 in PEL, but our screen uncovered

additional differentially activated kinases. For example, DLBCL exhibited higher activity of

LIMK2 compared to other subtypes. LIMK2 has been shown to play a role in pancreatic cancer

metastasis (188). Whether LIMK2 activity is an indicator or predictor of aggressiveness in

DLBCL is of interest. Our combined study of kinome profiling coupled with Tyro3 inhibitor

development revealed that the kinome profiles between NHL subtypes contain significant

differences, and that the interrogation of the kinome may help identify specific pathways that are

important for survival or pathogenesis in tumors. This approach is useful in devising

personalized therapies for different subsets of NHL, as well as many other solid and

hematological cancers.

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CHAPTER 3: VIRAL PROTEIN KINASE (VPK) MODULATION OF HOST

SIGNALING PATHWAYS3,4

Introduction

Similar to all herpesviruses, the life cycle of KSHV consists of a latent and lytic phase

(174, 189, 190). Both the latent and lytic phases of the life cycle are thought to be important for

the development of KSHV-associated diseases (189, 191, 192). Though complete lytic

reactivation results in cell death, it is proposed that lytic cells can contribute to the maintenance

of KSHV-associated diseases by secreting inflammatory cytokines or angiogenic factors that

provide growth signals in a paracrine manner. Another theory is that due to the instability of the

KSHV genome, lytic replication is necessary to produce more viral progeny to reseed the

reservoir of latently-infected cells. Indeed, cells extracted from Kaposi’s sarcoma (KS) lesions

tend to lose the viral genome after a few passages in cell culture (193).

KSHV encodes a viral protein kinase (vPK) on open reading frame 36 (ORF36), which is

normally expressed during the lytic cycle, and is packaged in the virion (67, 194, 195). While

vPK is mainly expressed during the lytic cycle, it can also be expressed during latency in

3Mass spectrometry data was generated by Aadra P. Bhatt in collaboration with Feng Yan and Erica Cloer from the laboratory of Ben Major. Plasmids for V5-vPK, the Flag-tagged PP6 regulatory subunits, and HA-PPP6C were generated by Aadra P. Bhatt and Louise C. Giffin. Small interfering RNA against PPP6C was tested and validated by Zhe Ma and Guoxin Ni. GST-PPP6 constructs were kindly provided by Xingzhi Xu. All other data were generated by the dissertation writer, Jason P. Wong.

4A small portion of the introduction has previously appeared as an article in the journal Biological Chemistry. The original citation is as follows: Wong, J and Damania, B. Copyright © Biological Chemistry. 2017 Jul 26; 398(8):911-918. doi: 10.1515/hsz-2017-0101.

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hypoxic conditions, which can occur in KS and primary effusion lymphoma (PEL) due to their

location in the body (196). In accordance with this theory, vPK was detected in 59% of KS

lesions (197). Deficiency in vPK results in an attenuation of lytic gene expression and production

of viral progeny (67). Virus produced from ORF36-deficient KSHV is also less effective at

infecting cells during primary infection. Therefore, expression of vPK is important for lytic

replication in the KSHV life cycle. Indicative of the role of vPK in KSHV-associated

malignancies, 80% of KSHV-MCD patients treated with a regimen targeting vPK attained major

clinical responses (5, 196). Furthermore, mice expressing the vPK-transgene developed

lymphomas at an eight-fold higher rate than the wild-type (WT) control (68).

KSHV-encoded vPK may contribute to the development of KSHV-associated

malignancies by altering signaling pathways that are frequently dysregulated in cancer. Some of

the cellular processes vPK has been shown to regulate include those involved in proliferative

signaling (61, 198-200), inhibiting the immune response (201), inflammation (56), DNA damage

(198, 200, 202), angiogenesis (61), and cell migration (203). Some of these functions can be

attributed to vPK’s mimicry of the cellular p70 S6 kinase (S6KB1) and cyclin-dependent kinases

(CDKs) (61, 198). Since kinases can alter a wide range of cellular processes, we were interested

in characterizing cellular binding partners of vPK to gain a better understanding of which

pathways vPK regulates in the host cell. From our previous work, we saw that even though vPK

and cellular S6KB1 shared many substrates, a substantial portion of unique substrates were

identified for vPK (61). Since a phosphopeptide screen does not recapitulate physiological

conditions, we performed a mass spectrometry screen by pulling down vPK expressed in 293T

cells to identify cellular binding partners. One of our top hits was a serine/threonine protein

phosphatase, protein phosphatase 6 (PPP6).

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PPP6 is a heterotrimer composed of one of three possible regulatory subunits (PPP6R1,

PPP6R2, and PPP6R3), one of three possible ankyrin subunits (ANKRD28, ANKRD48, and

ANKRD52), and a catalytic subunit (PPP6C). The regulatory subunits of PPP6 confer substrate

specificity, while the ankyrin subunit acts as the scaffold for the phosphatase (204-206).

Considered a tumor suppressor, PPP6 is known to regulate cell cycle progression, DNA damage

repair, autophagy, and inflammation (204, 207-216). PPP6 is important for mitotic progression

as PPP6-deficient cells have improper spindle formation and chromosome alignment due to

improper activation of Aurora kinase A (AURKA) (204, 211). Expression of a dominant-

negative mutant of PPP6 also resulted in a stall in the G1 phase of the cell cycle (217). PPP6 can

also regulate JNK and canonical NF-κB signaling by modulating the activity of TAK1 (210).

PPP6 dephosphorylates Thr187 of TAK1, thereby inactivating TAK1 and attenuating

downstream signaling. Furthermore in Drosophila, deficiency in the regulatory subunit of PPP6

resulted in low levels of basal activation of JNK (218). Highlighting the potential importance

PPP6 may have in tumorigenesis, two separate groups have reported PPP6C driver mutations in

approximately 10% of melanoma samples tested (219, 220). Therefore, KSHV may alter PPP6

signaling to promote disease progression in KSHV-related malignancies.

The temporal activation and expression of different proteins in the cell cycle is important

for proper division of the parent cell into daughter cells. Arguably the most well-known proteins

that regulate cell cycle progression are the cyclin-dependent kinases (CDKs) and the cyclins

(221-224). Cyclin D binds to CDK4 or CDK6 to regulate G1 phase progression, while cyclin E

binds CDK2 to regulate G1- to S-phase transition. Cyclin A binds to CDK2 to regulate S-phase

progression, and then later binds to CDK1 to regulate S- to late G2-phase progression. Finally,

cyclin B binds to CDK1 to regulate progression through mitosis. Since the binding of a cyclin to

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their respective CDK results in activation of other downstream targets, examining the temporal

expression of certain cyclins can inform which cellular processes are activated. KSHV encodes

several proteins known to regulate cell cycle progression during latency (225, 226). Interestingly,

overexpression of the vPK and a homolog of vPK in Epstein-Barr virus (EBV) resulted in a

mitotic-like state in cells, where proteins have activating phosphorylation marks normally

present in mitosis (200). As mentioned previously, vPK has been shown to have CDK-like

activity. Since PPP6 has been shown to have roles in mitotic progression, we were interested if

vPK may regulate PPP6 activity to alter progression through mitosis.

Another process we were interested in examining was autophagy as PPP6 has been

implicated with autophagy, while vPK has not (215, 216). The knockdown of PPP6C resulted in

the attenuation of mammalian target of rapamycin complex 1 (mTORC1)-dependent starvation-

induced autophagy (215). PPP6C was shown to activate the stress-kinase GCN2 during amino

acid starvation to induce phosphorylation of the alpha subunit of the eukaryotic translation

initiation factor 2 (eIF2α) to promote autophagy. Furthermore in NK T-cells (NKT), PPP6 is

thought to regulate the mTORC1-dependent inhibition of ULK1 in a TBK1 binding protein 1

(Tbkbp1)-dependent manner (216). Knockdown of PPP6C resulted in an increase in the

phosphorylation of the serine 757 residue of ULK1, which is normally phosphorylated by

mTORC1 to inhibit ULK1 activity (84). This corresponded to a decrease in phosphorylation of

serine residue 555 of ULK1, which is normally phosphorylated by 5’ AMP-activated protein

kinase (AMPK) to induce autophagy (227). Interestingly, eIF2α was determined to be a substrate

of vPK from our phosphopeptide screen (61). Since, the regulation of S6 by vPK was

independent of mTORC1, we were wondering if vPK can promote autophagy and protein

synthesis, which are two processes normally regulated by mTORC1 in opposing manners (61).

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In this chapter, we elucidate the interaction between vPK and different subunits of PPP6. We

also examine if vPK expression alters processes normally regulated by PPP6, and vice versa.

Then, we examine more in-depth if vPK has a role in autophagy and mitophagy, which is the

selective degradation of mitochondria by autophagy.

Materials and methods

Cell culture.

293T, 293GP, and iSLK.219 cells were grown in DMEM supplemented with 10% FBS,

1% penicillin-streptomycin, and 1% L-glutamine. 500 µg/mL of G418 was added to the 293GP

cells for selection. 250 µg/mL of G418, 400 µg/mL of hygromycin, and 10 µg/mL of puromycin

were also added to the iSLK.219 cells for selection as described previously (228). To reactivate

iSLK.219 cells from latency, 0.1 µg/mL of doxycycline as added to cells (229). A FLUOstar

OPTIMA plate reader was used to quantify RFP and GFP fluorescence as a measure for

reactivation. Human umbilical vein endothelial cells (HUVEC) were grown in endothelial cell

basal medium 2 (PromoCell, C-22211) supplemented with 1% L-glutamine, 10% FBS, 1%

penicillin-streptomycin, and the growth medium 2 supplement pack (PromoCell, C-39211)

excluding the addition of heparin and ascorbic acid from the pack (230).

Retrovirus infection of cell lines.

To generate cell lines stably expressing empty vector (EV) or viral protein kinase (vPK),

plasmids containing the TK502 backbone containing an EV or vPK transgene were used as

described previously (61). 293GP cells in a 10 cm dish were transfected with plasmids

containing VSV-G (5 µg) and either the EV or vPK transgene (5 µg) using Lipofectamine 2000

(Invitrogen, 11668019). The transfection was performed as detailed in the manufacturer’s

protocol. Three days after transfection, viral supernatants were collected and clarified of cellular

debris. Cells were centrifuged with either the retroviruses at 2500 rpm for 90 minutes at 30oC in

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serum-free media containing 10 µg/mL polybrene (Sigma, 107689). The next day, media was

refreshed and 800 µg/mL of G418 was added for selection.

Knockdown of genes.

To knockdown PPP6C, cells were transfected with 50 nM of a pool of two PPP6C siRNA

(Dharmacon, J-009935-07 and J-009935-08) or the non-targeting siRNA control (D-001810-01)

using Lipofectamine RNAiMAX (Invitrogen, 13778150) according to the manufacturer’s

instructions. Cells were harvested or analyzed after 48 or 72 hours of transfection.

Transfection of cell lines.

Plasmids containing either pcDNA3.1 Flag-tagged PPP6R1, Flag-tagged PPP6R2, Flag-

tagged PPP6R3, HA-tagged PPP6C, V5-tagged vPK, or an empty vector (EV) control were used

for transfection. 293T cells were transfected with a total of 2 µg (for 6-well plates) or 10 µg (for

10 cm plates) of plasmid using X-tremeGENE HP (Roche, 6366244001) or X-tremeGENE 9

(Roche, 6365779001) according to the manufacturer’s protocol. Cells were harvested three days

after transfection.

Nocodazole block and release.

Empty-vector (EV) or vPK HUVECs were knocked down with PPP6C or non-targeting

control siRNA 48 hours prior to performing nocodazole block and release. Cells were treated

with nocodazole (400 ng/mL) for 18 hours to arrest cells in mitosis. The next day, cells were

gently washed 2X with PBS, and fresh complete media was added. Cells were harvested at the

indicated time points, lysed in 0.1% NP-40 lysis buffer, and analyzed by immunoblotting.

Immunoblotting.

Immunoblotting was performed in a similar manner as described previously (230). Cells

were washed once in cold PBS before being lysed in 0.1% NP-40 lysis buffer [50 mM Tris

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(tris(hydroxymethyl)aminomethane)-HCl (pH 8), 150 mM NaCl, and 0.1% NP-40] containing

phosphatase inhibitors [30 mM β-glycerophosphate, 50 mM sodium fluoride, and 1 mM sodium

orthovanadate] and 1x Roche protease inhibitor tablet (Sigma-Aldrich, 11836145001). After 15

minutes of lysis on ice, lysate was clarified by centrifugation at 12,000 xg for 10 minutes at 4oC.

Protein quantification was performed afterwards using a Bradford assay (Bio-Rad, 500-0006).

Lysates were loaded with 5x urea buffer [10% w/v SDS, 10 mM β-mercaptoethanol, 20% v/v

glycerol, 0.2 M Tris-HCl (pH 6.8), 0.05% w/v bromophenol blue, and 8M urea] or 2X Laemmli

buffer (Millipore Sigma, S3401), and proteins were resolved by SDS-PAGE and then transferred

onto a nitrocellulose membrane. Primary antibodies used from Cell Signaling included HA-HRP

(1:5000, 14031), α-tubulin-HRP (1:5000, 9099), p-S6 (1:10000, 4858), total S6 (1:5000, 2217),

p-eIF2α (1:1000, 9721), total eIF2α (1:1000, 9722), LC3B (1:1000, 2775), ATF4 (1:1000,

11815), CHOP (1:1000, 2895), p-ULK1 S555 (1:1000, 5869), P-ULK1 S757 (1:1000, 14202),

total ULK1 (1:1000, 8054), p-Beclin-1 S90 (1:1000, 86455), p-Beclin-1 S93 (1:1000, 14717),

total Beclin-1 (1:1000, 3495), cyclin D1 (1:1000, 2978), cyclin A2 (1:1000, 4656), p-Aurora A

(T288)/B (T232)/C (T198) (1:1000, 2914), total Aurora A (1:1000, 4718), p-Optineurin S177

(1:1000, 57548), total Optineurin (1:1000, 58981), NDP52 (1:1000, 60732), p62 (1:1000, 5114),

and total PERK (1:1000, 3192). Primary antibodies from Santa Cruz included cyclin B1 (1:1000,

sc-245), vinculin (1:2500, sc-25336), GST (1:5000, sc-138), and cyclin E (1:1000, sc-481).

Other primary antibodies used were V5-HRP (Thermo Scientific, R961-25), DYKDDDDK-HRP

(Thermo Scientific, MA1-91878-HRP), PPP6C (Bethyl, A300-844A), PPP6C (EMD Millipore,

07-1224), p-PERK (Abcam, ab192591), MTCO2 (Abcam, ab79393), MTCO2 (Thermo

Scientific, A6404), and vPK (obtained as previously described (61)). Blots that were probed for

phosphorylated proteins were stripped with RestoreTM Stripping Buffer (Thermo Scientific,

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46430) and reprobed with the total antibody. Blots were developed with Pico West (Thermo

Scientific, 1863096), Clarity Western ECL substrate (Bio-Rad, 1705060), or ECL Prime (GE

Healthcare, RPN2232). Blots were imaged using a ChemiDoc imaging system and viewed with

Image Lab v5.2.1 software or with film.

Co-immunoprecipitation (Co-IP).

50 µg of protein was used for the input, and 2 mg of cell lysate was used for co-

immunoprecipitation (co-IP). The lysate was pre-cleared with mouse IgG (Santa Cruz, sc-2025)

and protein A/G agarose beads (sc-2003) for 45 minutes. The co-IP was performed according to

the manufacturer’s instructions with slight deviations. For pulling down V5-tagged vPK, 40 µL

of V5 beads (Sigma, A7345) were incubated with lysate overnight at 4oC. Afterwards, the beads

were washed 2X with 0.1% NP-40 lysis buffer, and then 2X with PBS. Protein was eluted using

V5 peptide (Sigma, V7754), and then a second elution was performed with 2X Laemmli buffer

(Millipore Sigma, S3401). The samples were then analyzed by Western blot. For pulling down

Flag-tagged PPP6 regulatory subunits, 40 µL of EZview Red Anti-Flag M2 affinity gel beads

(Sigma, F2426) were incubated with lysate overnight at 4oC. The number of washes and elutions

were performed similar to the V5-beads, except TBS was used for washing and the first elution

was performed using 3X Flag peptide (Sigma, F4799).

Purification of GST-tagged proteins.

Purification of Flag-tagged vPK protein was done as previously described before (61).

Constructs used for purifying the individual regulatory and catalytic subunits of PPP6 was

generously provided by Xingzhi Xu (213). The constructs used for recombinant protein

expression are pGEX-6P-1 GST, pGEX-4T1 GST-PPP6R1, pGEX-4T1 GST-PPP6R2, pGEX-

4T1 PPP6R3, and pcDNA3 3X-Flag-GST PPP6C. The protocol for expressing and purifying the

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GST-tagged PPP6 regulatory and catalytic subunits was performed using a mix of protocols as

described before (61, 67, 213, 231). BL21 Escherichia coli (E. coli) bacteria were transformed

with the plasmids encoding the GST constructs. A bacterial culture prepared from a starter

culture with an OD of 0.4-0.6 was induced with 200 µM IPTG overnight for 10 hours at 18oC.

Afterwards, the bacteria was pelleted at 6500 rpm for 10 minutes, and resuspended in NETN [20

mM Tris pH 7.5, 100 mM NaCl, 1 mM EDTA, 0.5% NP-40, 1 mM DTT (Sigma, 43815), 1 mM

PMSF, and 1 Roche Protease Inhibitor tablet (Sigma-Aldrich, 11836145001)]. Bacteria were

then lysed using a Branson Digital Sonifier at max amplitude for 2x 30 seconds on ice. The

lysate was rocked for 30 minutes at 4oC, and then clarified by centrifugation at 10,000 xg for 10

minutes at 4oC. For each milliliter of bacterial lysate, 50 µg of prepared Glutathione Sepharose

beads (GE Healthcare, 17-0756-01) were used to pulldown the expressed GST-fusion proteins.

The bacterial lysate was incubated on beads for one hour (or overnight) at 4oC. After washing

beads 3X with NETN buffer, and then 3X with PBS, GST-fusion proteins were eluted with 10

mM reduced glutathione, pH 8 (Sigma, G4251). A second elution was performed and combined

with the first elution. Glycerol (final concentration 10%) was added to the combined elutions,

before dialysis was performed with Slide-A-Lyzer dialysis G2 cassettes (Thermo Scientific,

87718) in PBS buffer containing 10% glycerol, 1 µg/ml leupeptin, 1 µg/ml pepstatin A, 0.15 mM

PMSF, and Roche Protease Inhibitor tablets (Sigma-Aldrich, 11836145001) at 4oC for 4 hours.

The eluent was removed from the cassette and concentrated using Amicon centrifugal filters (10

kDA – UFC801024 or 50 kDA – UFC805024) by spinning at 4000 rpm for 40 minutes at 4oC.

The concentrated eluent was removed and quantified using a Bradford assay (Bio-Rad, 500-

0006). The purified protein was aliquoted and snap-frozen in a dry ice-ethanol bath and stored in

the -80oC for future use.

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In vitro binding assay.

Glutathione Sepharose beads (GE Healthcare, 17-0756-01) were prepared as indicated in

the manufacturer’s protocol. Recombinant GST-tagged regulatory or catalytic subunits were

added to the beads in NETN buffer, and rotated on a Nutator for 1 hour at 4oC to allow binding.

Afterwards, beads were washed 3x with NETN buffer, and then incubated with 3% BSA in

NETN buffer for 30 minutes – 1 hour on a Nutator at 4oC to block non-specific binding. Next,

recombinant Flag-vPK was added to the beads complexed with the recombinant GST-fusion

proteins, and was incubated at 4oC for one hour on a Nutator. The beads were washed 5X with

NETN buffer, and bound proteins were eluted with 2X Laemmli buffer (Millipore Sigma,

S3401). The elution was separated into two fractions and ran on separate SDS-PAGE gels: one

fraction was used for Coomassie staining and another was used for immunoblotting.

Coomassie staining.

Gel staining was performed as described from a protocol from the lab of Julianne Grose

(https://mmbio.byu.edu/grose-lab/lab-protocols) with minor deviations (232). Briefly, the gel

was rocked in gel-fixing solution [50% (v/v) ethanol and 10% (v/v) acetic acid in water] for 30

minutes. Afterwards, the gel was incubated in gel-washing solution [50% (v/v) methanol and

10% (v/v) acetic acid in water] overnight. The gel was then stained with Coomassie blue [0.1%

(w/v) Coomassie blue R350, 20% (v/v) methanol, and 10% (v/v) acetic acid in water] for 2

hours, and washed with gel-washing solution until the background was sufficiently low.

Afterwards, the gel was incubated in storage solution [5% (v/v) acetic acid in water] for 1 hour.

The gel was imaged on the ChemiDoc imaging system and viewed with Image Lab v5.2.1

software.

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In vitro kinase assay.

The in vitro kinase assay was performed in a similar manner as described before (61).

Recombinant Flag-tagged vPK was diluted in kinase buffer [50 mM Tris-HCl (pH 7.5), 0.1 mM

EGTA, 1 mg/mL BSA, and 0.1% (v/v) 2-betamercaptoethanol (BME)] before performing the in

vitro kinase assay. 100 ng of recombinant vPK was incubated with 2 µg of either a recombinant

PPP6 regulatory/catalytic subunit or the GST-control in reaction buffer [20 mM HEPES (pH

7.5), 5 mM MnCl2, and 10 mM BME] with 10 µCi γ-P32 labeled ATP (Perkin Elmer,

NEG002A250UC) at 37oC for 30 minutes. The reaction was quenched with 2x Laemmli buffer

(Millipore Sigma, S3401) and ran on a SDS-PAGE gel. Afterwards, the gel was fixed and

stained, and imaged using a radiation cassette. The cassette was developed using a Typhoon FLA

9500 (GE Healthcare) and the gel was imaged using the ChemiDoc imaging system.

Tubule formation assay.

The tubule formation assay was performed in a similar manner as described previously

(233). Empty-vector (EV) or vPK HUVECs were knocked down of PPP6C (or the non-targeting

control) using siRNA for 48 hours prior to starting the tubule formation assay. 500 µL of

Matrigel (Corning, 354230) was plated in 24-well plates and allowed to solidify at 37oC for 30

minutes. Afterwards, EV or vPK HUVECs were detached using trypsin, and then plated in 1 mL

of complete media on top of the Matrigel at a concentration of 1x105 cells per well. Tubule

formation was assessed every hour between two to four hours and images were taken using a

Nikon Eclipse Ti microscope with a 4x objective. Tubules were counted manually as nodes

containing three or more branches.

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Mitochondrial fractionation.

Mitochondria were collected from cells using a mitochondrial fractionation kit (Thermo

Scientific, 89874) according to the manufacturer’s instructions. Briefly, cells from 10 cm dishes

were collected, and resuspended in 400 µl of Reagent A. Two minutes after resuspension, 5 µl of

Reagent B was added. Cells were vortexed every minute for 5 minutes, and then 400 µl of

Reagent C was added. Cells were spun down at 700 xg for 10 minutes at 4oC. The supernatant

was transferred to a new tube, and then further spun down at 3000 xg for 15 minutes at 4oC. The

supernatant containing the cytosolic fraction was collected, and the pellet containing the

mitochondria was washed with 500 µl of Reagent C. After centrifugation at 12,000 xg for 5

minutes at 4oC, the wash was removed, and the pellet was resuspended in 2X Laemmli buffer

(Millipore Sigma, S3401). Samples were later ran on a SDS-PAGE gel for immunoblotting.

Mitophagy dye stain.

Mitochondria were stained and mitophagy was assessed using a mitophagy dye kit

(Dojindo, MD01-10) according to manufacturer’s instructions. Poly-L lysine coated coverslips

(Corning, 354086) were placed in 24-well plates and coated with a gelatin solution (Sigma, ES-

006-B) for 30 minutes at 37oC. Afterwards, the coverslips were air-dried for one hour before

2x105 HUVEC EV or vPK cells were added to the well. The next day, cells were washed 2x with

PBS, and then incubated with 500 µL of Mtphagy Dye working solution at 37oC for 30 minutes

in serum-free media. Afterwards, the dye solution was removed, and cells were washed 2X with

PBS. After cells were grown for an additional 24 hours, the cells were fixed by incubating them

in a 3.7% formaldehyde solution in PBS for 15 minutes. Cells were washed 2X with PBS and

mounted on a slide using an anti-fade mounting stain for DAPI (Thermo Scientific, P10144).

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Cells were visualized three days later on a Nikon Eclipse Ti microscope with a 60x objective

using oil.

Mass spectrometry analysis.

Pulldown of Flag-tagged vPK for mass spectrometry analysis was performed as

previously described (234). Flag-tagged vPK was expressed in 293T cells. Cells were lysed and

lysates were clarified. The clarified lysate was added to Flag-M2 beads (Sigma) to pull down

Flag-vPK. After washing the beads, samples were eluted using 3x FLAG peptide (Sigma).

Samples were digested using trypsin with a FASP protein digestion kit (Protein Discovery).

After tryptic peptide cleanup, samples were analyzed by mass spectrometry as previously

described (234). Peptide counts generated from a GFP negative control was used to filter out

non-specific interactions.

Ingenuity Pathway Analysis (IPA).

Analysis was performed using the IPA program from Qiagen. Data was uploaded as

differential peptide counts between the pull-down of a GFP control and Flag-tagged vPK. Core

analysis was performed with no changes to the default parameters except only human

interactions were examined. The canonical pathways analysis bar graph was generated based on

pathways containing –log(p-values) > 1.3 and a z-score > ±1.5.

Results

vPK interacts and phosphorylates the regulatory subunits of PPP6

To identify new host cellular binding partners to vPK, we pulled down Flag-tagged vPK

from 293T cells and performed mass spectrometry (Table 3.1). We found several proteins which

had previously been identified to interact with vPK, such as tripartite motif-containing 28

(TRIM28) and DExH-box helicase 9 (DHX9). Multiple subunits of protein phosphatase (PPP6),

including all three regulatory subunits (PPP6R1, PPP6R2, and PPP6R3), two of three ankyrin

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Table 3.1 Peptide counts of proteins identified to interact with vPK using mass spectrometry.

Protein Peptide Count PPP6R3 14 TRIM28 12 PPP6C 8 DHX9 5

PPP6R1 5 ANKRD28 3

PPP6R2 1 ANKRD52 1

subunits (ANKRD28 and ANKRD52), and the catalytic subunit, PPP6C were pulled down in our

mass spectrometry screen. In concordance with this data, a group which pulled down all the open

reading frames (ORFs) in KSHV (to determine binding partners for all KSHV-encoding ORFs)

also pulled down PPP6R3, PPP6C, and ANKRD28 with vPK (ORF36) (116). To confirm our

mass spectrometry data, we performed a co-immunoprecipitation (co-IP) in 293T cells

overexpressing Flag-tagged regulatory subunits of PPP6 and V5-tagged vPK. We determined all

three regulatory subunits of PPP6 immunoprecipitated with vPK (Figure 3.1A). Similarly,

PPP6C, which was HA-tagged, pulled down with vPK as well (Figure 3.1B). To confirm the co-

IP data, we performed a reverse co-IP by pulling down each of the regulatory subunits, and

likewise saw that vPK interacted with each of the regulatory subunits (Figure 3.1C). Even though

vPK interacts with the catalytic and regulatory subunits of PPP6, vPK may not directly bind to

the PPP6 subunits, but rather bind another protein, which acts as an intermediary between them.

To determine if vPK binds the PPP6 subunits directly, we purified recombinant Flag-tagged

vPK, and GST-tagged PPP6 regulatory and catalytic subunits to perform an in vitro binding

assay (Figure 3.1D, E). We found that the catalytic and all three regulatory subunits of PPP6 can

individually bind vPK directly. Since phosphorylation of substrates is the main mechanism by

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Figure 3.1 vPK binds and phosphorylates several subunits of the PPP6 complex. (A) 293T cells were transfected either with empty vector (EV) or V5-tagged vPK, and Flag-tagged PPP6R1, PPP6R2, or PPP6R3. V5-tagged vPK was immunoprecipitated, and elutions were analyzed by Western. (B) 293T cells were transfected with HA-tagged PPP6C, and in different combinations with or without V5-tagged vPK and all of the regulatory subunits, which were Flag-tagged. V5-tagged vPK was immunoprecipitated, and elutions were analyzed by Western.

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R1, R2, R3 (+) samples indicate cells that were transfected with all three Flag-tagged regulatory subunits in equal amounts. (C) 293T cells were transfected either with empty vector (EV) or V5-tagged vPK, and Flag-tagged PPP6R1, PPP6R2, or PPP6R3. Individual Flag-tagged regulatory subunits were immunoprecipitated, and elutions were analyzed by Western. (D) Recombinant purified GST, GST-tagged regulatory subunits (PPP6R1, PPP6R2, or PPP6R3) or GST-tagged PPP6C were incubated with or without recombinant Flag-tagged vPK in an in vitro binding assay. GST proteins were eluted from GST-beads, and presence of vPK was analyzed by Western. The blue asterisks indicate the location of the vPK band. (E) A portion of the elution harvested in (D) was run on an SDS-PAGE gel and Coomassie stained. A sample containing recombinant vPK alone was also run as a marker. The red asterisks indicate the respective location of each of the purified GST proteins in each sample. (F) An in vitro kinase was performed with samples containing recombinant GST, GST-tagged regulatory subunits, or GST-tagged PPP6C with or without recombinant vPK. The red arrow indicates the location of phosphorylated regulatory subunits, while the blue arrow indicates the location of autophosphorylated vPK. The Coomassie stain of the gel is below. The blue asterisks indicate the location of the vPK band. The red asterisks indicate the respective location of each of the purified GST proteins in each sample.

which kinases alter their substrates’ activity or function, we examined if vPK could

phosphorylate any of the PPP6 subunits by performing an in vitro kinase assay (Figure 3.1F). We

saw that vPK phosphorylates all three regulatory subunits, but not PPP6C. Interestingly, the

autophosphorylation band, which is indicative of the kinase activity of vPK, is severely reduced

in the PPP6C sample (195, 235). This could possibly be due to PPP6C dephosphorylating vPK;

however, PPP6C produced from bacteria was previously reported to be inactive (236, 237), and

follow-up p-nitrophenyl phosphate (pNPP) experiments demonstrated that PPP6C had no activity

(data not shown). A possible explanation is EDTA levels may have been too high in the PPP6C

reaction. Since the Roche protease inhibitor tablet contains EDTA, which is used during the

dialysis step, EDTA is potentially present in the solution upon storing the recombinant proteins.

Due to the difficulty of purifying PPP6C, the concentration of PPP6C was approximately 8-fold

lower compared to the regulatory subunits. This necessitates the presence of more storage

solution in the in vitro kinase reaction; therefore, more EDTA is present in the reaction. EDTA

acts as a metal chelator and may be sequestering the manganese ions in the reaction buffer. Metal

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ions are important for the enzymatic activity of kinases as they frequently coordinate the transfer

of the phosphate group within the catalytic pocket (238). Therefore, we cannot fully rule out the

possibility that vPK can phosphorylate PPP6C. Optimization of conditions and a repeat of the in

vitro kinase assay are necessary to confirm that vPK does not phosphorylate PPP6C. While we

examined if vPK can phosphorylate the PPP6 subunits, it is possible that PPP6 may have

enzymatic activity towards vPK. An in vitro phosphatase assay to examine if the PPP6 complex

can dephosphorylate vPK can be done in the future.

Examining the role of PPP6 on pathways modulated by KSHV

PPP6 is a family member of the protein phosphatase 2A (PP2A) group, which include

PP2A, protein phosphatase 4 (PP4), and PPP6 (236). Similar to PPP6, PP2A can form a trimeric

holoenzyme consisting of a regulatory, scaffolding, and catalytic subunit to dephosphorylate

substrates (239). Several reports have demonstrated that the phosphorylation of the regulatory

subunit of PP2A can regulate PP2A activity (240, 241). Since vPK phosphorylated the regulatory

subunits of PPP6 (Figure 3.1F), we were interested in examining if expression of vPK cells

altered some of the signaling pathways regulated by PPP6. One of the more well-studied

pathways that PPP6 regulates is the dephosphorylation of Aurora kinase A (AURKA) to control

progression through mitosis (204, 209, 211, 242, 243). AURKA is responsible for proper spindle

pole formation, and depletion of AURKA results in mitotic arrest (244). However,

overexpression of AURKA in cancers commonly leads to chromosome instability (245).

Therefore, the activation and inactivation of AURKA is important for proper progression

through mitosis. During mitosis, AURKA is bound to TPX2, which results in the

autophosphorylation and activation of AURKA (246, 247). PPP6 recognizes the AURKA-TPX2

complex and dephosphorylates AURKA, resulting in the inactivation of the complex (211).

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Figure 3.2 Examining the role of PPP6C in multiple pathways and cellular processes relevant in the KSHV life cycle. (A) Empty-vector (EV) or vPK-expressing HUVECs were knocked down with either a non-template control (NTC) or PPP6C-targeting siRNA. After 48 hours of transfection, cells were blocked with 2 µM nocodazole for 18 hours. Cells were released, and collected at 0, 15, 30, 60, 90, and 180 minutes after release, and are indicated by the numbers 1, 2, 3, 4, 5, and 6, respectively. The red arrow indicates the location of the p-AURKA band, and the black dashed lines are to help with clarity. (B) iSLK.219 cells were transfected with siRNA against NTC or PPP6C for 48 hours, and then reactivated with doxycycline. The ratio of RFP:GFP fluorescence intensity (indicative of reactivation) was measured using a plate reader 0 to 72 hours after doxycycline treatment. (C) Western blot examining knockdown of EV or vPK-HUVECs transfected with NTC or siPPP6C siRNA used in the tubule formation assay in (D, E). (D) EV and vPK-HUVECs were transfected with NTC or siPPP6C siRNA for 48 hours. Cells were collected and 1x105 cells/well were plated on Matrigel

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for a tubule formation assay. Images were taken 3 hours after plating, and the number of branching points was counted in each field. (E) Representative pictures of (D).

BGLF4, which is the homolog to vPK in Epstein-Barr virus (EBV), has been shown to induce a

mitotic-like state in cells (200). Overexpression of BGLF4 resulted in increased expression of

proteins highly expressed during mitosis including AURKA, Aurora kinase B, cyclin B1, and

nucleolar and spindle associated protein 1 (NUSAP1). For instance, AURKA expression

normally increases during the G2-phase, and is degraded during mitosis and the G1-phase of the

cell cycle (245). These two observations would suggest vPK (and potentially BGLF4) is

mediating a mitotic-like state by either altering the stability of proteins expressed during mitosis

or by preventing the cell from properly progressing through mitosis. To elucidate if vPK alters

cell cycle progression through mitosis, we treated EV- and vPK-expressing human umbilical

vein endothelial cells (HUVECs) with nocodazole, which arrests cells in mitosis by preventing

the proper assembly of the mitotic spindle (Figure 3.2A) (248). We then removed media

containing nocodazole and added fresh complete media to the cells, thereby allowing progression

through mitosis. Since mitosis lasts around two hours in most cells, we harvested cells at

multiple time points ranging from zero hours to three hours after nocodazole release. As

expected, cells had the highest expression of cyclin B1 (a mitotic cyclin) at the zero hour time

point of harvest. Consistent with literature, knocking down PPP6C resulted in prolonged

expression of cyclin B1 in both EV- and vPK-expressing cells, indicating a slower progression

through mitosis (211). The expression of vPK alone also resulted in prolonged expression of

cyclin B1, which was further exacerbated when PPP6C was knocked down. Furthermore, the

onset of cyclin D1 expression (a G1-phase cyclin) was almost completely attenuated in vPK-

expressing cells with PPP6C knocked down. These results suggest vPK expressing cells take

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longer to progress through mitosis, which is further exacerbated by PPP6C knock down.

Consistent with this conjecture, phosphorylation of AURKA was sustained longer in vPK-

expressing cells. These results suggest vPK may be inhibiting PPP6 activity to promote a

mitotic-like state in cells. Therefore, to see if inhibiting PPP6 results in a pro-viral state within

the cell, we knocked down PPP6C in iSLK.219 cells, and reactivated them using doxycycline.

Interestingly, cells depleted for PPP6C had lower lytic viral reactivation compared to those

knocked down with the non-template control (NTC) (Figure 3.2B). To further investigate this

interaction, we examined tubule formation in EV- and vPK-expressing HUVECs with or without

PPP6C knocked down (Figure 3.2C). We had previously shown that vPK augmented tubule

formation, which is a hallmark for angiogenic potential. Similar in pattern to our reactivation

data, PPP6C knock down resulted in decreased formation of tubules in both EV- and vPK-

expressing cells (Figure 3.2D, E). This suggests that vPK inhibition of PPP6C may not be

beneficial for lytic viral replication and KSHV pathogenesis. Further work would need to be

done to determine if PPP6 is antagonistic or beneficial for the KSHV life cycle. PPP6 may also

play different roles during lytic replication, latency, and KSHV-associated pathogenesis.

Autophagy markers are upregulated by vPK expression

PPP6 has been implicated in the induction of autophagy (215, 216). Since vPK has not

been shown to regulate autophagy, we were interested in examining if vPK may regulate PPP6

activity to modulate autophagy during the KSHV life-cycle. Previous reports have shown that

autophagy is induced during lytic replication, though the maturation of the autophagosome is

blocked (97, 99). We also noticed that eukaryotic translation initiation factor 2 subunit alpha

(eIF2α) may potentially be a substrate for vPK from our phosphopeptide screen (61). The

phosphorylation of eIF2α results in the attenuation of 5’ cap-dependent translation, and promotes

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Figure 3.3 Ingenuity pathway analysis (IPA) of host interacting partners of vPK identified using mass spectrometry. Core analysis was performed on the host interacting partners that were identified from the mass spectrometry screen using IPA (Qiagen) to identify potential pathways regulated by KSHV. Graph was generated with cutoff values for pathways having a –log(p-value) > 1.3 and z-score > ±1.5.

the translation of short upstream open reading frames (85). Some of these encode for

transcription factors such as ATF4 and CHOP that upregulate autophagy (85, 89). Interestingly,

IPA analysis of host proteins identified in the vPK mass spectrometry analysis indicated that

eIF2 signaling may be regulated by vPK (Figure 3.3). The expression of vPK in HUVEC cells

resulted in an upregulation of the phosphorylation of eIF2α, and the autophagy marker LC3B-II

(Figure 3.4A). The conversion of LC3B-I to LC3B-II allows LC3B to become membrane-

associated which allows it to be recruited to the phagophore assembly site (PAS) (81, 82).

Previous literature had shown that PPP6C upregulates the phosphorylation of eIF2α in nutrient-

starved conditions. However, we saw that knock down of PPP6C resulted in variable results, and

frequently led to an increase in the phosphorylation of eIF2α rather than a decrease (Figure

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3.4A). A caveat to this observation is that we performed this experiment under full nutrient

conditions. It is possible that different results may be obtained when performing this experiment

in nutrient-starved conditions. We also examined if autophagic flux was impaired by treating

cells with chloroquine for three hours (Figure 3.4B). An increase in LC3B-II expression may be

due to the inhibition of the fusion of the lysosome with the autophagosome (249). This would

result in an accumulation of autophagosomes containing LC3B-II even though autophagy is

inhibited. Chloroquine inhibits autophagy by raising the pH within lysosomes, which prevents

the degradation of cargo within the autolysosome. If treatment of cells with chloroquine does not

increase the levels of LC3B-II, this would indicate the increase in LC3B-II formation in vPK-

expressing cells is due to a block in autophagy. An increase in LC3B-II was observed in the

vPK-expressing HUVECs upon chloroquine treatment suggesting that autophagic flux was not

impaired in vPK expressing cells (Figure 3.4B).

Various kinds of cellular stress can induce different kinases to phosphorylate eIF2α.

Under endoplasmic reticulum (ER)-stress, the PKR-like ER kinase (PERK) becomes activated,

and phosphorylates eIF2α. Under amino acid depletion, detection of double-stranded RNA

(which occurs under viral infection), and heme deficiency, the kinases GCN2, PKR, and HRI

become activated and phosphorylate eIF2α, respectively. We examined if any of these kinases

are activated by vPK to upregulate eIF2α phosphorylation. Since, PKR had previously been

shown to inhibit KSHV-infection, and there is no good antibody to detect HRI phosphorylation,

we did not examine these two kinases (250). We did not see any difference in PERK activation

between EV- and vPK-expressing HUVECs (Figure 3.4C). We also tried examining

phosphorylation of GCN2, but results were variable and detection by immunoblotting was

difficult (data not shown). Therefore, to determine if phosphorylation of eIF2α led to differential

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Figure 3.4 Autophagic markers are induced but not the integrated stress response (IRS) in vPK-HUVECs. (A-C) Empty-vector (EV) or vPK-HUVECs were transfected with siRNA against a non-template control (NTC) or PPP6C for 48 hours. Lysates were collected and assessed by Western. (B) 48 hours after transfection, EV or vPK-HUVECs were treated with H2O control or 25 µM chloroquine for 2 hours before lysates were collected. (D) Lysates from EV or vPK-HUVECs were assessed for ATF4 and CHOP expression by Western. EV-HUVECs treated with tunicamycin were used as a positive control. (E) EV or vPK-HUVECs were treated with H2O control or 100 µM CoCl2 for 24 hours before lysates were harvested and analyzed by

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Western. Expression of BNIP3 was assessed as its expression is upregulated by CoCl2. Red asterisk indicates the band that is p-ULK1 S555.

translation of mRNAs, we examined ATF4 and CHOP protein expression. There was no

detectable protein expression of ATF4 or CHOP by Western in the EV- or vPK-expressing

HUVECs (Figure 3.4D). In contrast, treatment of EV-HUVEC cells with tunicamycin, a drug

that induces ER stress resulted in robust expression of ATF4 and CHOP. Therefore, while vPK-

expressing HUVECs have upregulated phosphorylation of eIF2α, this induction may not be

sufficient enough to cause physiological changes. We also examined if the rest of the autophagy

signaling pathway was activated by examining the activation state of the ULK1-Beclin1 axis. We

used cobalt (II) chloride as a positive control as it induces autophagy by mimicking a hypoxic

state within the cell. Interestingly, both serine 90 and serine 93 in Beclin1 had increased

phosphorylation in untreated vPK-HUVECs suggesting Beclin1 was more activated in vPK-

expressing cells (Figure 3.4E). Concordantly, serine 555 of ULK1, an indicator of ULK1

activation, showed higher phosphorylation levels in vPK-expressing cells. Overall, the

phosphorylation status of ULK1 and Beclin1 suggests the ULK1-Beclin1 axis of autophagy is

activated in vPK-expressing cells.

vPK expression promotes mitophagy

Since vPK expression did not induce differential translation of mRNAs, but still led to

activation of the ULK1-Beclin1 complex, we wondered if instead of macroautophagy, a type of

selective autophagy was induced. A recent study showed that the viral interferon factor 1

(vIRF1) encoded by KSHV was able to interact with one of the mitophagy receptors, NIX, to

induce mitophagy (115). Furthermore, reactivation in PEL cells resulted in an induction of

mitophagy and a decrease of mitochondrial protein levels. Another study had also identified that

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Figure 3.5 vPK expression promotes mitophagy. (A, B) EV or vPK-expressing HUVECs were treated with DMSO control or 10 µM CCCP, which is a drug that induces mitophagy. (A) Cells were harvested 3 hours after treatment and the mitochondria were extracted. The mitochondrial fraction was analyzed by Western. (B) Cells were harvested 24 hours after treatment, and whole cell lysates was analyzed by Western. (C) EV and vPK-HUVECs were seeded on coverslips and stained with a mitophagy dye. 24 hours later, cells were fixed, and mounted with a DAPI anti-fade mounting solution. Staining was analyzed by fluorescence microscopy using oil with a 60x objective. The dim red color is stained mitochondria, while the bright red puncta are stained mitochondria localized to autolysosomes.

vPK interacted with many proteins related to ribosome biogenesis and mitochondria translation

(116). Therefore, we were interested to examine if vPK may induce mitophagy. We fractionated

mitochondria from cells either treated for three hours with DMSO control or 10 µM of carbonyl

cyanide 3-chlorophenylhydrazone (CCCP), which induces mitophagy by depolarizing the

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mitochondrial membrane (Figure 3.5A) (112). The three hour time point is an acute time point to

examine the recruitment of mitophagy receptors. In EV-HUVECs, we see an increase of LC3B-

II in the mitochondria upon CCCP treatment, indicative of formation of the phagophore.

Likewise, an increase in the recruitment of PINK1-Parkin-related mitophagy receptors,

optineurin (Optn), p62, and NDP52 occurs. However in vPK-expressing HUVECs, there is a

reduction protein level of all three mitophagy receptors and LC3B-I, and reduced recruitment of

LC3B-II compared to EV cells with CCCP treatment. Likewise, there is sharp reduction in

protein levels of the mitochondrial proteins leucine rich pentatricopeptide repeat containing

(LRPPRC) and mitochondrially encoded cytochrome c oxidase II (MTCO2). Furthermore, we

see a reduction in expression levels of vPK in the mitochondria upon CCCP treatment (possibly

due to vPK being degraded with the mitochondria). These results suggest that vPK induces or

otherwise makes cells amenable for mitophagy. The phosphorylation of optineurin by tank

binding kinase 1 (TBK1) has been shown to promote autophagy (251). We see increased

phosphorylation of Optn in vPK-expressing HUVECs under basal conditions (Figure 3.5B).

Furthermore, when staining cells in normal conditions with a mitophagy dye, we see increased

red puncta in vPK-HUVECs, suggesting more mitochondria are undergoing mitophagy

compared to EV-HUVECs (Figure 3.5C). The mitophagy dye stains all mitochondria red, but has

increased fluorescence in acidic conditions (i.e. when mitochondria are within autolysosomes).

Therefore, the brighter red puncta indicate mitochondria within lysosomes. Overall, these results

suggest that vPK expression promotes mitophagy.

Discussion

As previously reported, inhibition of mitophagy results in an attenuation of lytic protein

expression and viral replication (115). There are several reasons why KSHV may induce

mitophagy during the lytic cycle. Autophagy and apoptosis crosstalk heavily with each other and

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in some cases induction of autophagy delays the onset of apoptosis (252). Damaged

mitochondria also induce apoptosis if not eventually cleared (253). The high metabolic and

energy stress the virus induces during the lytic cycle to produce progeny is likely to result in

mitochondrial stress. Several viruses have been reported to induce mitophagy to inhibit apoptosis

(254). Furthermore, the mitochondrion is a hub for innate immune signaling such as through

mitochondrial antiviral signaling protein (MAVS)-retinoic acid-inducible gene 1 (RIG-I)

signaling. Likewise, several viruses have induced mitophagy to inhibit innate immune signaling

(254). We have previously shown that MAVS and RIG-I inhibit KSHV lytic replication and

virion production (255). Therefore, KSHV may induce mitophagy to delay the onset of apoptosis

and inhibit the interferon response.

Interestingly, both vPK and vIRF1 have been shown to inhibit type 1 interferon responses

mediated by TBK1 (201, 256). Our lab has shown that vIRF1 prevents stimulator of interferon

genes (STING) from activating TBK1, thereby inhibiting interferon-beta (IFN-β) production.

Nevertheless, reactivation of iSLK.219 cells did induce some phosphorylation and activation of

TBK1. On the other hand, vPK binds activated interferon regulatory factor 3 (IRF3), but not the

inactive form of IRF3, and prevents it from properly acting as a transcription factor to inhibit the

type 1 interferon response (201). It is possible that the virus is able to subvert the activation of

TBK1 during viral infection to upregulate mitophagy while sufficiently ablating the activation of

the type 1 interferon response. Currently, one protein encoded by KSHV, vIRF1, is known to

induce mitophagy by in a NIX-dependent manner (115). However, the depletion of NIX only

resulted in a marginal reduction in lytic replication and KSHV ORF45 expression. This suggests

there are other mitophagy receptors and viral proteins which may mediate mitophagy during the

KSHV lytic cycle. Future work includes examining if the optineurin receptor is phosphorylated

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and activated in a TBK1-dependent manner during viral reactivation. Extra time points after

CCCP induction are also necessary to clarify the kinetics of vPK regulation of mitophagy.

Currently, we are able to determine that vPK interacts with the PPP6 holoenzyme and is

able to phosphorylate the regulatory subunits of PPP6. However, the physiological relevance of

the phosphorylation of the regulatory subunits by vPK remains to be elucidated. Our attempts to

determine the role of PPP6 in pathways regulated by vPK and KSHV has resulted in variable and

conflicting results. One possible explanation is that the pathways regulated by PPP6 are

regulated differently during different stages of the KSHV life cycle. For instance, AURKA is

upregulated and activated during latency by the latency-associated nuclear antigen (LANA) and

in KSHV-associated malignancies (230, 257). Furthermore, PPP6 inhibits prolonged activation

of NF-κB (210, 258), which is upregulated during latency and important for PEL survival (31,

33, 34, 36, 259). Since the inhibition of PPP6 results in prolonged activation of AURKA and NF-

κB, this would be beneficial for the latently-infected cell. However, the activation of NF-κB can

also lead to activation of the type 1 interferon response (260, 261), which is detrimental for lytic

replication (46, 201, 255, 256). There are some reports that NF-κB activity is downregulated in

PEL cells upon reactivation, and the rescue of NF-κB activity results in an inhibition of viral

replication (34, 262). PPP6C has also been shown to enhance MAVS-RIG-I signaling (263).

Therefore, activation of PPP6 during lytic replication would possibly be beneficial. Further work

would need to be done to examine if PPP6 activity is regulated differently during latency and the

lytic cycle, and whether or not this is dependent on vPK.

In this chapter we have shown vPK influences a wide variety of cellular pathways.

While we are unable to elucidate the manner by which vPK regulates PPP6 activity, we are able

to confirm vPK does bind to multiple subunits of PPP6. More work would need to be done to

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determine if the phosphorylation of the regulatory subunits by vPK alters PPP6 functionality.

Interestingly, DNA-protein kinase (DNA-PK) is a common substrate between these two proteins;

therefore, examining DNA-PK activity may be helpful in characterizing the interaction between

vPK and PPP6 (208, 264, 265). Furthermore, we also show that vPK is able to promote

mitophagy, which has several implications for the KSHV life cycle ranging from innate immune

signaling to energy metabolism. Further work is necessary to elucidate the mechanism by which

vPK promotes mitophagy. Targeting the mitophagic process during the KSHV life-cycle may

hold promise as a therapeutic target.

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CHAPTER 4: CONCLUSIONS AND FUTURE DIRECTIONS

Summary

My dissertation research, performed under the guidance of Dr. Blossom Damania, has

examined how both host (Chapter 2) and viral kinases (Chapter 3) play a role in the Kaposi’s

sarcoma-associated herpesvirus (KSHV) life cycle. Kinases frequently act as signaling

intermediaries within signaling pathways, and therefore are important in turning on or off certain

cellular processes. Since kinases are enzymes, they can amplify a signal quickly. My work

provides evidence that Tyro3, a host receptor tyrosine kinase, promotes cell survival and that

targeting Tyro3 may be beneficial in the treatment of primary effusion lymphoma. Furthermore,

my studies have examined host cellular binding partners for KSHV-encoded viral protein kinase

(vPK), and that it may have a role in promoting mitophagy. Due to kinases being a readily

“druggable” target (266), elucidating the role of kinases in the KSHV life cycle can open the

door to the initiation of drug discovery programs to treat KSHV-associated diseases. MIB/MS

has previously been used to identify potential host kinases that could be therapeutic targets for

human cytomegalovirus (HCMV) infection (267).

In Chapter 1, a broad overview of the KSHV life cycle is described with a focus on

pathways that are modulated by KSHV. This chapter also highlights kinases within these

pathways that are dysregulated by KSHV. For chapter 2, we examined which host kinases are

uniquely activated in primary effusion lymphoma (PEL) in comparison to other B-cell non-

Hodgkin lymphomas (NHL) to determine the kinases that PEL may be dependent on for

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pathogenesis. Since the prognosis for patients with PEL is poor and no standard treatment

regimen exists, we were interested in identifying new therapeutic targets in PEL (268). Kinases

that are uniquely activated in PEL may be an Achilles’ heel as PEL cells may be more dependent

on the activity of that particular kinase in comparison to healthy B-cells. We identified Tyro3 as

our most significant hit, and characterized its role in upregulating the PI3K/Akt/mTOR pathway

to promote cell survival. Furthermore, we developed an inhibitor against Tyro3 called

UNC3810A, which showed efficacy in both cell culture and in a PEL-xenograft mouse model.

For chapter 3, we examined host interacting partners of KSHV-encoded vPK. Since vPK is

important for late gene lytic expression and the production of viral progeny (67), identifying host

interacting partners of vPK can help determine pathways that are important in the lytic phase of

the KSHV life cycle. Furthermore, several points of evidence indicate that vPK may play a role

in several KSHV-associated malignancies (5, 14, 61, 68, 269). We identified that several

subunits of the protein phosphatase 6 (PPP6) interacted with vPK. PPP6 is known to be

dysregulated in melanoma and plays roles in multiple pathways associated with cancer including

cell cycle progression, DNA damage repair, and inflammation (204, 208, 210, 213, 218, 264).

Further work is necessary to elucidate the importance of the interaction between PPP6 and vPK.

Our analysis of vPK-interacting partners also led to a hypothesis that vPK may play a role in

eukaryotic initiation transcription factor 2 (eIF2) signaling and mitophagy. While we found that

the alpha subunit of eIF2 (eIF2α) was phosphorylated, we did not see any induction of

differential translation of mRNAs upregulated by the integrated stress response (IRS) that

normally occurs when eIF2α is phosphorylated (85, 89). However, we saw the induction of

several autophagy markers, and determined that mitophagy was upregulated by vPK.

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As summarized above, we took two different approaches with a focus on kinases to study

which pathways are important in the KSHV life cycle. The first project identified a host cellular

kinase that regulates a pathway that is known to be important in KSHV-associated pathogenesis

(10). The second project, which examined host interacting partners of a KSHV-encoded kinase,

identified a pathway that has only been recently characterized in the KSHV life cycle (115). The

subsequent part of this chapter further expands on the implications of these projects, and

proposes future studies to better characterize the role of these pathways and kinases in KSHV-

mediated pathogenesis.

Upregulation of several pathways in KSHV-associated PEL by Tyro3

As highlighted in chapter 1, several pathways are well-known to be dysregulated by

KSHV, which include the PI3K/Akt/mTOR, JAK/STAT, NF-κB, and the MAPK pathways.

Kinases tend to regulate more than one pathway as they can phosphorylate a variety of substrates

(50). Since most of the infected cells in KSHV-associated malignancies are primarily latent, both

host and a subset of viral factors are necessary to promote KSHV-associated tumorigenesis.

Interestingly, while many proteins have been identified to regulate the PI3K pathway (vGPCR,

vPK, vIL-6, K1, and K15), many of them are primarily lytic proteins (with some having low

expression during latency) and are thought to contribute to PI3K signaling in KSHV-associated

malignancies in a paracrine manner. Indeed infection by KSHV is necessary, but not sufficient to

result in KSHV-associated malignancies (1). Therefore, identifying host cellular kinases that are

upregulated in KSHV-associated malignancies is important.

Receptor tyrosine kinases can modulate several pathways in part due to their position in

the signaling cascade (270). Since they act as both the receptor and kinase, they initiate the

signaling cascade rather than acting as bridging molecules. This allows them to become activated

under a variety of stimuli to induce multiple signaling cascades (270). Highlighting this

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versatility, Tyro3 can modulate other pathways besides the PI3K pathway such as the

MAPK/ERK (184, 185), p38 MAPK (187), JAK/STAT (164, 184), and PLC-γ pathways (186).

Figure 4.1 Unresolved questions regarding the role of Tyro3 in PEL. A basic outline depicting the activation of the PI3K pathway by Tyro3, which UNC3810A, a Tyro3 inhibitor can inhibit. Four questions of interest still remain in our study of Tyro3 in PEL. (1) Does Tyro3 upregulate other pathways in PEL besides PI3K? (2) What induces Tyro3 in PEL? (3) Will treatment of PEL with UNC3810A sensitize PEL cells to standard chemotherapeutic regimens? (4) Does Tyro3 regulate glycolysis in PEL cells in a PI3K-dependent manner?

We only characterized the role of Tyro3 in PI3K signaling in PEL; this beseeches the question

does Tyro3 play a role in these other pathways, which are known to be upregulated in PEL?

From an efficiency standpoint, it would be worthwhile for KSHV to alter the activity of Tyro3 to

upregulate all of these pathways. We would hypothesize that Tyro3 would activate other

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pathways besides the PI3K pathway in PEL (Figure 4.1). While we identified Tyro3 was

activated in PEL cells, another group had previously identified Axl - a related family member to

Tyro3 - as being upregulated in KS tissues (79). Interestingly from our work, we did not find Axl

to be expressed at high levels in PEL cells. This suggests that the cellular origin, B-cells or

endothelial cells, may play a role in which TAM receptor is upregulated in certain KSHV-

associated malignancies (Figure 4.1).

From a therapeutic standpoint targeting Tyro3 is an attractive option. As discussed above,

Tyro3 may affect multiple pathways important in PEL pathogenesis. Furthermore, we have

shown that Tyro3 can be targeted with a kinase inhibitor with some specificity against the other

TAM family members. A prominent characteristic of the TAM receptors is that they promote

chemoresistance in many different cancers (76, 78). Similarly, PEL is generally resistant to

standard chemotherapy (271). While preliminary, we have shown that treatment of PEL cells

with UNC3810A sensitizes them to several PI3K inhibitors in a synergistic manner. Therefore,

combination regimens utilizing a Tyro3 inhibitor and standard chemotherapy drugs may have

some efficacy (Figure 4.1). Furthermore, many different cancers such as melanoma and colon

cancer have upregulated Tyro3 expression and activity (146, 160, 162, 168). Therefore,

inhibitors developed against Tyro3 would be beneficial in treating a wide range of cancers

besides PEL.

Another outstanding question is whether Tyro3-mediated activation of PI3K signaling

plays more of a role besides promoting cell survival. The PI3K/Akt/mTOR signaling pathway is

a central hub in the cell as it is the main regulator of energy- and nutrient-dependent cellular

processes. Previous work in our lab had identified that glycolysis is highly upregulated in PEL

cells, which was dependent on PI3K signaling (272). Therefore, Tyro3 inhibition may lead to

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attenuation of glycolysis in a PI3K-dependent manner (Figure 4.1). Considering inhibition of

Tyro3 resulted in an attenuation of PI3K signaling, we would expect glycolysis to be negatively

affected as well when inhibiting Tyro3. Many different cancers heavily utilize and upregulate

glycolysis (272). Healthy cells usually rely on oxidative phosphorylation, which occurs within

the mitochondria, to generate energy as it is a highly efficient way to generate ATP (273). A

possible reason on why cancer cells upregulate glycolysis is that it generates ATP at a much

faster rate compared to oxidative phosphorylation, which is important when considering the

limited amount of nutrients and resources in the tumor microenvironment (273). Indeed

inhibition of glycolysis in PEL cells resulted in cell death (272). Therefore, targeting PI3K to

attenuate PI3K signaling may serve two purposes: (1) inhibiting protein biosynthesis and (2)

inhibiting glycolysis-dependent energy production. However, cancer cells still require and rely

on mitochondrial metabolism not only for energy production, but also to regulate many different

cellular processes (274). It would be interesting to examine the interplay between glycolysis and

mitochondrial-dependent metabolism, and their respective contributions in PEL pathogenesis.

Targeting of mitochondria during the KSHV lytic cycle

If the PI3K/Akt/mTOR pathway is the main nutrient and energy regulator in the cell,

then the mitochondria would be the primary organelle related to energy production in eukaryotic

cells. Healthy mitochondria are necessary for proper energy homeostasis within the cell (109,

111). Highlighting the importance of the mitochondria, many apoptotic proteins (and thus

apoptosis) are inherently tied to mitochondrial-related signaling pathways (253). The degradation

and recycling of old or defective mitochondria can be completed by a selective autophagic

process known as mitophagy. While general autophagy (i.e. macroautophagy) has been

examined in multiple studies (97-99, 101, 105), only recently has mitophagy been appreciated to

play a role in the KSHV life cycle (115). A group reported that mitophagy was important for

101

KSHV lytic replication, and that induction of mitophagy was partially due to viral interferon

regulatory factor 1 (vIRF1)-dependent activation of NIP3-like protein X (NIX) (115). However,

depletion of NIX in reactivated PEL cells only modestly affected lytic replication. This suggests

other mitophagy receptors other than NIX and/or other viral proteins regulate mitophagy during

lytic replication.

Figure 4.2 Potential questions regarding the role of vPK-induced mitophagy in the KSHV life cycle. A basic outline is shown of two different cellular processes: (1) NIX-mediated and ubiquitin-dependent mitophagy and (2) MAVS/RIG-I and cGAS/STING dependent type I interferon induction. Both vIRF1 and vPK can block type I interferon in different manners. Furthermore, vIRF1 can induce mitophagy in a NIX-dependent manner. Several questions still remain regarding vPK induction of mitophagy, and the role of mitophagy during the lytic cycle of KSHV: (1) How does vPK induce the phosphorylation of optineurin (and mitophagy in general)? (2) Does inhibiting mitophagy result in increased type I interferon signaling due to upregulation of MAVS? (3) Does inhibiting mitophagy result in increased apoptotic signaling during the KSHV lytic cycle? OMM is an abbreviation for outer mitochondrial membrane.

102

In chapter 3, we show that vPK promotes mitophagy as degradation of mitochondria was

accelerated upon carbonyl cyanide 3-chlorophenylhydrazone (CCCP) treatment in comparison to

empty-vector (EV)-treated cells. Furthermore, a mitophagy receptor, optineurin (Optn), had

higher levels of a mitophagy-activating phosphorylation mark in vPK expressing cells. These

results would suggest that vPK can induce mitophagy by activating the mitophagy receptors.

However, several questions still remain. The most pertinent question is determining the

mechanism by which vPK induces the phosphorylation of Optn (Figure 4.2). Examining the

commonalities between vPK and vIRF1 may provide a clue. Both proteins are known to inhibit

type I interferon responses. A host protein known to regulate both type I interferon responses and

mitophagy is the tank binding kinase 1 (TBK1). Furthermore, TBK1 is known to phosphorylate

optineurin on the residue that is upregulated in vPK-expressing cells. It is possible TBK1 activity

is regulated by vPK or that vPK can phosphorylate the mitophagy receptors directly. Considering

reactivation of iSLK.219 cells is shown to result in an increase in TBK1 activity (256), KSHV

may downregulate the induction of type I interferon responses, while subverting the activation of

TBK1 to promote mitophagy.

Several possible reasons exist on why KSHV may promote mitophagy. Mitochondria can

act as a signaling hub for innate immune signaling through mitochondrial antiviral signaling

protein (MAVS). We have previously shown that MAVS inhibits KSHV lytic replication (255),

and other viruses have utilized mitophagy to evade immune responses (254). It would be

interesting to examine if inhibition of mitophagy resulted in an increase in antiviral responses

during lytic reactivation (Figure 4.2). As noted above, many apoptotic proteins are regulated by

mitochondrial-related pathways (253). The persistence of defective or excessive amounts of

mitochondria can lead to the activation of apoptosis. While lytic cells eventually undergo

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apoptosis, an inhibition of mitophagy may lead to an earlier onset of apoptosis in reactivated

cells (Figure 4.2). Therefore, the induction of mitophagy may be beneficial for KSHV lytic

replication on several levels.

Elucidating the vPK and PPP6 interaction

Our initial foray into mitophagy was due to our observation that vPK and PPP6 interacted

with each other and that PPP6 plays a role in the induction of autophagy (215). Although vPK

could play a role in the integrated stress response (ISR), there was no discernable induction in

downstream targets of eIF2α signaling in vPK-expressing cells. Furthermore, our attempt at

examining the role of PPP6 in the induction of autophagy had variable and sometimes different

results as reported in the literature. Therefore, the importance of the interaction between vPK and

PPP6 in the KSHV life cycle remains to be determined. A few possible leads for future

investigation exist. One pathway which we did not examine that PPP6 regulates is DNA repair.

PPP6 dephosphorylates γ-H2AX after the repair of double-stranded DNA breaks (204, 213, 264).

While expression of vPK does not induce γ-H2AX, its homologs murine γ-herpesvirus 68

(γHV68)-encoded orf36 and EBV-encoded BGLF4 did in mice fibroblast cells (265). However,

KSHV infection of endothelial cells is known to induce γ-H2AX. From these observations

several questions exist. (1) Is the lack of γ-H2AX induction by vPK in the previous report due to

its expression in mice fibroblasts rather than its physiological cell type? (2) After a double-

stranded DNA break is the dephosphorylation of γ-H2AX by PPP6 altered when vPK is

expressed? (3) Is PPP6 activity inhibited (or even relevant) during KSHV infection as seen by

the persistence of γ-H2AX upon KSHV infection? Elucidating these questions may help identify

if PPP6 has a role during the KSHV life cycle and if vPK mediates PPP6 activity.

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Conclusions

In my dissertation, we have shown that elucidating the role of kinases is a useful method

to better understand pathways that KSHV regulates during the different stages of its life cycle,

and in the context of KSHV-associated diseases. Studying these pathways has also provided

insight on possible targets for therapeutic intervention, which is important for PEL patients, who

generally have a poor prognosis. While my work focused on KSHV-related pathogenesis, the

work in my dissertation has wider implications in other diseases. A common theme throughout

my dissertation is how KSHV modulates pathways that tend to be dysregulated in cancers. While

we only examined Tyro3 signaling in PEL, many other cancers such as melanoma and colon

cancer are known to upregulate Tyro3 to promote proliferation and/or chemoresistance (160,

162, 168). The tool compound that we developed, UNC3810A, can be used to study other

cancers that have high Tyro3 activity. Finally, we also see that KSHV-encoded vPK can promote

mitophagy. There is an increasing appreciation in the oncology field that mitophagy may

promote chemoresistance in a variety of cancers (275-278). Considering vPK can mimic both

cellular cyclin-dependent kinases (CDKs) and the p70 S6 kinase, it would be interesting to

determine if any of these cellular kinases can also promote mitophagy. Many discoveries of host

cellular processes have been made by studying viruses, ranging from the identification of the Src

oncogene to the inner workings of the innate immune system. In summation, by studying how

KSHV alters its host cell, we can gain a better understanding of host cell biology.

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