Mechanisms of Regulation of Promyelocytic Leukemia (PML) Proteins by Epstein-Barr Nuclear Antigen 1 (EBNA1) and

Ubiquitin Specific Protease 7 (USP7)

by

Cameron Landry

A thesis submitted in conformity with the requirements

for the degree of Master’s of Science

Department of Molecular Genetics University of Toronto

© Copyright by Cameron Landry 2013

ii

Mechanisms of Regulation of Promyelocytic Leukemia (PML) Proteins by Epstein-Barr Nuclear Antigen 1 (EBNA1) and

Ubiquitin Specific Protease 7 (USP7)

Cameron Landry

Master of Science

Department of Molecular Genetics University of Toronto

2013

Abstract

Promyelocytic leukemia (PML) nuclear bodies (NBs) are important nuclear structures

that mediate tumour suppressive and antiviral functions. Epstein-Barr virus (EBV) has evolved

mechanisms to disrupt these structures and abrogate their functions, through the EBNA1

protein. EBNA1-mediated PML NB disruption requires its interaction with two cellular

proteins, CK2 and USP7, both of which negatively regulate PML in the absence of EBNA1. I

determined that USP7-null cells have greatly increased PML NBs. I have demonstrated the

importance of serine 393 of EBNA1, a phosphosite essential for the EBNA1-CK2 interaction.

An S393A mutation abrogated the ability of EBNA1 to disrupt PML NBs, while a S393T

mutation restored this ability. I have also determined that EBNA1 and USP7 can directly

interact with PML proteins, primarily with PML IV. In summary, my data supports a model in

which EBNA binds PML proteins directly and negatively regulates them through recruitment of

CK2 and USP7.

iii

Acknowledgements

First and foremost, I would like to sincerely thank my supervisor and mentor Dr. Lori

Frappier. Thank you for giving me the opportunity to pursue research in your lab. You have

always been extremely supportive and encouraged me to ask new questions and explore new

ideas. Your door was always open, whether it was to discuss results, provide feedback (and the

fastest feedback of any professor!), or to simply chat about our mutual love of travelling. Lori,

thank you so much for your understanding and everything you have done for me! I would not be

in a position to pursue my dreams without you.

Second, thank you to my supervisory committee, Dr. Jack Greenblatt and Dr. David

Bazett-Jones. Thank you for making my committee meetings quite enjoyable and for providing

valuable insight. I appreciate your interest in this project, as your feedback, suggestions, and

criticisms have all made me a better scientist! Thank you both for caring about and facilitating

my success as a scientist.

I would also like to extend my upmost gratitude to the members of the Frappier lab.

Much of my success as a graduate student is owed not only to their helpful suggestions or

Kathy’s killer baked goods, but also to our friendships. Kathy, you are seriously a pro-lab tech

and a pro-baker. Your birthday cakes were only one of the many things you did to made the

Frappier lab feel like home. Without your insight and suggestions and our many

discussions/problem-solving chats, I would not have had anywhere near the success I did. Thank

you also for your sense of humour! You are simply amazing and I have learned so much from

you. Tin, you are the protein master! Thank you so much for your invaluable suggestions and

encouraging feedback! Patrick and Sheila, thank you for coffee time. I looked forward to this

everyday, as not only was there fantastic coffee but there was also fantastic conversations. I

loved being able to talk science (and sometimes not science) so early in the morning! Also,

congrats on the soon-to-be-new-borns (I’m stoked I won baby-bets!). Madhav, future lab

meetings will never be the same without you. You were always willing to challenge everyone’s

ideas and ask critical questions (without any hesitation mind you!) that made us all better

scientists. Jen, thank you for our countless discussions about PML, EBNA1, and CK2 (yes there

have really been so many I can’t count them)! Your enthusiasm for science is simply unmatched

and I will miss this very much. Natasha, thank you so much for all your support (whether it was

in the lab or not) along the way. You have been a big sister to me and have always had my back.

You have also been the best “work-wife” anyone could ever ask for and I really appreciate our

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mutual love of sarcasm! Anna, thanks for joining the lab! You always bring happiness into the

lab and have already increased our fun factor by 10! Thank you also to everyone for the many

after-lab-beers we shared! You have all made my experience in the lab as awesome as my

experience outside the lab. Lori and everyone in the Frappier lab, I am forever grateful.

My fellow graduate students (you know who you are), thank you for taking this journey

with me, supporting me, and ensuring the time we did have away from the lab was well spent!

Thank you also to the Varsity Blues tennis team; you guys are awesome and you made our

Egypt adventure the trip of a lifetime! Special shout out to Cyrus, Cecelia, Farhaad, Jill, Julia,

Arren, Anna, Jocelyn, and our favourite coach Nabil! And thanks to my roommates, Kylie,

Richard, and Paulette. You guys are amazing!

I would finally like to thank my family. Thank you Mom, Dad, and Kevin for all your

support along the way. You guys have truly shaped who I am today and have always

encouraged me to pursue my goals and dreams, whether they have taken me across the country

or across the globe!

Thank you so much everyone. My time during graduate school in Molecular Genetics

has been a blast!

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Table of Contents

Abstract………………………………………………………………………………………… ii Acknowledgements……………………………………………………………………………. iii Table of Contents………………………………………………………………………………. v List of Tables…………..……………………………………………………………………… vii List of Figures………………………………………………………………………………... viii List of Abbreviations…………………………………………………………………..……… ix

1   Introduction ........................................................................................................................... 1  1.1   Promyelocytic leukemia (PML) nuclear bodies (NBs) .............................................................. 1  

1.1.1   Structure, composition, and dynamics of PML NBs ............................................................... 1  1.1.2   Regulation of PML proteins and PML NBs ............................................................................ 6  1.1.3   PML NB associated functions ................................................................................................. 6  

1.1.3.1   Tumour suppressive functions ........................................................................................................ 7  1.1.3.2   Antiviral functions .......................................................................................................................... 8  

1.1.3.2.1   PML and RNA viruses ............................................................................................................ 9  1.1.3.2.2   PML and DNA viruses ......................................................................................................... 10  

1.1.4   Viral proteins that disrupt PML NBs .................................................................................... 11  1.2   Epstein-Barr virus (EBV) .......................................................................................................... 14  

1.2.1   Overview of the EBV life cycle ............................................................................................ 14  1.2.2   Epstein-Barr nuclear antigen 1 (EBNA1) ............................................................................. 16  

1.2.2.1   Functions of EBNA1 on the EBV episome .................................................................................. 18  1.2.2.2   Cellular functions of EBNA1 ....................................................................................................... 20  

1.3   Ubiquitin specific protease 7 (USP7) ......................................................................................... 23  1.3.1   Structural organization and catalytic activity ........................................................................ 23  1.3.2   Cellular functions of USP7 .................................................................................................... 25  

1.4   Thesis rationale ........................................................................................................................... 26  2   Materials and Methods ....................................................................................................... 28  

2.1   Cell lines ....................................................................................................................................... 28  2.2   Plasmid construction .................................................................................................................. 28  2.3   Transfections ............................................................................................................................... 28  2.4   Cell proliferation assay ............................................................................................................... 29  2.5   Transient replication assay ........................................................................................................ 29  2.6   Plasmid labeling .......................................................................................................................... 30  2.7   Southern blot ............................................................................................................................... 30  2.8   Immunofluorescence microscopy .............................................................................................. 31  2.9   Western blots ............................................................................................................................... 31  2.10   EBNA1 and USP7 purification ................................................................................................ 31  2.11   In vitro binding assay ................................................................................................................ 32  

3   Results ................................................................................................................................... 33  3.1   Contributions of PML NBs to EBNA1-mediated DNA replication ....................................... 33  3.2   Analysis of PML NBs in USP7-null cells .................................................................................. 34  3.3   Contributions of Serine 393 in EBNA1 to PML NB disruption ............................................. 36  3.4   Characterization of the direct interactions of EBNA1 and USP7 with specific PML isoforms ................................................................................................................................................. 39  

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4   Discussion and Future Directions ...................................................................................... 46  4.1   Thesis summary .......................................................................................................................... 46  4.2   PML NBs do not contribute to oriP-plasmid replication ........................................................ 47  4.3   USP7-null cells have increased amounts of PML NBs and PML protein ............................. 48  4.4   Serine 393 of EBNA1 plays a critical role in EBNA1-mediated PML NB disruption .......... 49  4.5   EBNA1 and USP7 preferentially interact with PML isoform IV in vitro .............................. 50  4.6   Future directions ......................................................................................................................... 51  

4.6.1   Determining how EBNA1 and USP7 interact with PML ...................................................... 51  4.6.2   Determining the role of autophagy in PML NB degradation ................................................ 52  

4.7   Concluding remarks ................................................................................................................... 61  5   References ............................................................................................................................. 62  

vii

List of Tables

Table 1. Viral proteins that alter or disrupt PML NBs through PML proteins 11

Table 2. Epstein-Barr virus latency programs 15

viii

List of Figures

Figure 1-1: PML NBs and associated functions 2

Figure 1-2: Schematic of PML isoforms 4

Figure 1-3: Overview of the EBV lifecycle 17

Figure 1-4: EBNA1 structural organization 19

Figure 1-5: Schematic of USP7 structural organization 24

Figure 3-1: PML silencing does not reduce plasmid replication efficiency in CNE2 cells 35

Figure 3-2: HCT116 USP7 -/- Cells Have Increased PML NBs Compared to HCT116 WT 37

Figure 3-3: The ability of EBNA1 to disrupt PML nuclear bodies is abrogated by the 38 S393A mutation and restored by the S393T mutation

Figure 3-4: EBNA1 and USP7 Interact Directly with PML isoforms I, IV, & VI 40 Figure 3-5: EBNA1 titration demonstrates PML IV binding specificity 42 Figure 3-6: USP7 titration demonstrates PML IV binding specificity 44 Figure 4-1: EBNA1 can induce PML NB disruption in CNE2Z cells in the presence of a 53 proteasomal inhibitor (MG132) Figure 4-2: Schematic of autophagy 55 Figure 4-3: An autophagy inhibitor (3MA) and a proteasomal inhibitor (MG132) 58 additively block PML protein degradation in CNE2E cells

Figure 4-4: Inhibiting autophagy in CNE2 cells causes an increase in PML NBs 60

ix

List of Abbreviations

aa amino acid AIDS Acquired immune deficiency syndrome APL Acute promyelocytic leukemia Atgs Autophagy related genes ATM Ataxia telangiectasia mutated protein ATR Ataxia telangiectasia and Rad3-related protein BLM Bloom syndrome protein Bp Base pair BRCA1 Breast cancer type 1 susceptibility protein Brd4 Bromodomain 4 BRLF1 BamHI R leftward frame 1 BSA Bovine serum albumin BZLF1 BamHI Z leftward frame 1 CAT Chloramphenicol acetyl transferase CBP CREB binding protein CDK Cyclin-dependent kinase CHK2 Checkpoint kinase 2 CK2 Casein kinase 2 CMV Cytomegalovirus CREB cAMP response element-binding Daxx Death associated protein DAPI 4'-6-Diamidino-2-phenylindole dATP Deoxyadenosine triphosphate dCTP Deoxycytidine triphosphate dGTP Deoxyguanosine triphosphate DMEM Dulbecco’s modified Eagle’s medium DS Dyad symmetry DTT Dithiothreitol dTTP Deoxythymidine triphosphate DUB Deubiquitinating enzyme E3 ubiquitin ligase EBER Epstein-Barr expressed RNA EBNA Epstein-Barr nuclear antigen EBNA-LP Epstein-Barr nuclear antigen leader protein EBV Epstein-Barr virus ECL Enhanced chemiluminescence EDTA Ethylene diamine tetraacetic acid FBS Fetal bovine serum FLASH FLICE-associated huge protein FLICE FADD-like IL-1β-converting enzyme FOXO4 Forkhead box O 4 FR Family of repeats GA Glycine alanine GAM Goat-anti mouse GAR Goat-anti rabbit GC Gastric carcinoma GFP Green fluorescent protein

x

GAPDH Glyceraldehyde 3-phosphate dehydrogenase GMPS Guanosine monophosphate synthetase H2A Histone 2A H2B Histone 2B HAUSP Herpesvirus associated ubiquitin specific protease HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HCMV Human cytomegalovirus HCV Hepatitis C virus HDACs Histone deacetylases HDV Hepatitis D virus HFV Human foamy virus HIPK2 Homeodomain interacting protein kinase 2 HIV Human immunodeficiency virus HPV Human papilloma virus HSV Herpes simplex virus ICP0 Infected cell protein 0 ICP27 Infected cell protein 27 IE Immediate early IF Immunofluorescence IFN Interferon IM Infectious mononucleosis KSHV Kaposi’s sarcoma associated herpes virus LAMP1 Lysosomal-associated membrane protein 1 LANA Latency associated nuclear antigen LC3 Microtubule-associated protein 1 light chain 3 LCL Lymphoblastoid cell line LCMV Lymphocytic choriomeningitis virus LMP Latent membrane protein 3MA 3-methyladenine MCMs Minichromosome maintenance Mdmx Mouse double minute x Mdm2 Mouse double minute 2 MHC Major histocompatibility complex NAP Nucleosome assembly protein NBR1 Neighbor of BRCA1 gene ND10 Nuclear Domain 10 NEM N-ethylmaleimide NES Nuclear export signal NLS Nuclear localization signal NPC Nasopharyngeal carcinoma ORC Origin recognition complex PCNA Proliferating cell nuclear antigen PAR Poly-ADP-Ribosylation PARP Poly-ADP ribose polymerase PML Promyelocytic leukemia PML NBs Promyelocytic leukemia nuclear bodies PMSF Phenylmethylsulfonyl fluoride POD PML oncogenic domains pRB Retinoblastoma protein oriP Origin of plasmid replication

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RARα         Retinoic acid receptor alpha RBCC Ring finger, B-box, Coiled-coil RING Really interesting new gene RNF4 RING finger protein 4 RSV Respiratory synctial virus SAHFs Senescence-associated heterochromatin foci SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis SENP SUMO specific protease SIM SUMO interacting motif siRNA Small interfering RNA shRNA Small hairpin RNA Sp100 Speckle pattern 100 protein SQSTM1 Sequestosome 1 SUMO Small ubiquitin-like modifier SV40 Simian virus 40 TAF-1 Template activating factor 1 TNF-R Tumor necrosis factor receptor TRADD TNF-R1 associated death domain protein TRAF Tumor necrosis factor receptor associated factor TRIM Tripartite motif TUNEL Terminal deoxynucleotidyl transferase dUTP nick end labeling UBC9 Ubiquitin carrier protein 9 USP7 Ubiquitin specific protease 7 UV Ultraviolet VSV Vesicular stomatitis virus VZV Varicellar zoster virus

1

1 Introduction

1.1 Promyelocytic leukemia (PML) nuclear bodies (NBs)

The promyelocytic leukemia protein (PML) has been actively researched for over a

decade since it was identified to be important in acute promyelocytic leukemia (APL). The PML

gene was originally identified in APL patients as a result of a reciprocal chromosome

translocation that juxtaposes the PML gene on chromosome 15 and the retinoic acid receptor

(RAR)α gene on chromosome 17 (Kakizuka et al., 1991). Consequently, an oncogenic chimeric

PML-RARα protein is produced that disrupts the function of wild-type PML proteins (i.e. the

disruption of nuclear domains containing PML protein, termed PML nuclear bodies (NBs)) in a

dominant negative manner (Brown et al., 2009; Dyck et al., 1994; Wang and Chen, 2008).

Interestingly, PML NBs are restored by two anti-APL therapies which trigger PML-RARα

degradation: retinoic acid and arsenic trioxide (Quignon et al., 1997; Zhu et al., 2001). This

identified the first parallel between the status of the nuclear bodies and that of the cell.

Accumulating evidence has confirmed that PML has many tumour suppressive functions

beyond APL in non-haematopoietic tumours. Regulation of appropriate levels of PML proteins

and their accumulation in NBs is crucial since loss of PML NBs accompanies the development

and/or progression of several tumours (Salomoni et al., 2008; Bernardi and Pandolfi, 2007;

Gurrieri et al., 2004). PML NBs can serve as scaffolds for a multitude of cellular processes,

including p53 activation, apoptosis, DNA damage responses, transcriptional regulation,

replicative senescence, and innate antiviral responses via the interferon pathway (Figure 1-1)

(Salomoni et al, 2008; Dellaire and Bazett-Jones, 2004; Bernardi et al, 2008; Lallemand-

Breitenback and de Thé, 2010; Everett & Chelbi-Alix, 2007; Geoffroy and Chelbi-Alix, 2011).

Hence, understanding the regulation of PML proteins and the NBs formed by them is very

biologically important.

1.1.1 Structure, composition, and dynamics of PML NBs

PML proteins form distinct nuclear foci called PML nuclear bodies (NBs), also known

as nuclear domain 10s (ND10s) or PML oncogenic domains (PODs) (Stuurman et al., 1992).

These subnuclear protein structures are spheres of 0.1-1.0 µm in diameter and typically range

from 5-30 NBs per cell, in a cell-cycle and cell-type dependent way (Maul et al., 2000; Dellaire

and Bazett-Jones, 2004). Electron microscopy studies have shown that PML NBs are composed

of a ring-like protein structure that does not contain detectable nucleic acids in the center;

2

Figure 1-1: PML NBs and associated functions. PML NBs, which contain the PML proteins, form distinct nuclear structures (the green foci) that are visualized in the centered immunofluorescence microscopy (IF) image. In this image, PML NBs are stained with an antibody against PML proteins in nasopharyngeal carcinoma (NPC) cells. The exact number of PML NBs varies from cell to cell and between cell lines as well. The variety of PML-associated functions are shown around the IF image; these functions are mediated by the numerous proteins that either stably or transiently associate with PML NBs.

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however, newly synthesized RNAs have been found to associate with the periphery of the ring

(Boisvert et al., 2000). Extensive protein-based threads also extend from the core facilitating

contacts with chromatin fibers important for maintaining NB integrity and positional stability

(Eskiw et al., 2004). Moreover, PML NBs are non-randomly associated with transcriptionally

active areas of the genome; specific associations with chromosome loci have been detected with

the major histocompatibility (MHC) gene cluster, the p53 gene locus, and most recently the

ABCA7, TFF1 and the PML loci (Wang et al., 2004; Shiels et al., 2001; Kumar et al., 2007; Sun

et al., 2003; Ching et al., 2013).

The human PML gene is expressed as 7 alternatively spliced mRNAs, from 9 different

exons, each encoding a distinct protein that varies in size from 48-97 kDa (Jensen et al., 2001)

(Figure 1-2). PML proteins contain domains important for protein-protein interactions as well as

self-multimerization. They are members of the tripartite motif (TRIM) family of proteins since

the N-terminal region contains an RBCC motif (composed of a Really Interesting New Gene

(RING), B-box, and coiled-coil regions) (Nisole et al., 2005). All seven PML isoforms (I-VII)

contain the TRIM domain in the N-terminal region (exons 1-4); however, they differ in their C-

terminal sequences (exons 5-9), resulting in unique PML protein profiles and the potential for

isoform specific functions (Jensen et al., 2001; Salomoni et al., 2008). PML isoforms I-VI

contain a nuclear localization signal (NLS) in exon 6, whereas isoform VII does not contain an

NLS and is therefore cytoplasmic (Melnick and Licht, 1999). Notably, PML isoform I also

contains a nuclear export signal (NES), consistent with its localization to the nucleus and the

cytoplasm (Salomoni and Bellodi, 2007). Moreover, multiple bands of PML can be observed,

when analyzed by SDS-PAGE followed by Western blotting, ranging from 50-160 kDa due to

post-translational modifications of the multiple PML isoforms (Burkham et al., 2001; Jensen et

al., 2001; Pandolfi et al., 1991).

PML is post-translationally modified by the small ubiquitin-like modifier (SUMO) at

three lysine residues (K65, K160, and K490) via the catalytic activity of the SUMO-conjugating

enzyme UBC9 (Boddy et al., 1996; Ishov et al., 1999; Sternsdorf et al., 1997; Kamitani et al.,

1998; Borden, 2002). These SUMO modifications are critical for NB formation as they mediate

interactions with the SUMO-interacting motifs (SIMs), defined by a VVVI consensus sequence,

present in the 7a exon of PML (Shen et al., 2006). PML can be modified by any of the three

SUMO isoforms 1, 2, and 3 and polymetric SUMO2/3 chains are important for nuclear

accumulation of PML and the structure of PML NBs (Duprez et al., 1999; Vertegaal et al., 2006;

Fu et al., 2005; Mukhopadhyay et al., 2006). Overexpression of SUMO-specific proteases such

4

Figure 1-2: Schematic of PML isoforms. PML protein comprises six nuclear isoforms I-VI and one cytoplasmic isoform VII, encoded from a single gene that is alternatively spliced. The domain organization is shown (top), indicating the positions of the Ring domain (R), Beta Box 1 (B1), Beta Box 2 (B2), Coiled-coil domain (C), and the Exonuclease domain (E). The Nuclear localization signal (N), not in PML VII, and the sites of sumoylation (black arrows) are also shown. The lower panel indicates exon usage; exons 1-6 (dark blue) are common to all nuclear isoforms, where the variability of the C-terminal regions is indicated by several alternatively spliced exons (light blue). (Adapted from Salomoni et al., 2008).

5

as SENP-1, SENP-2, or SENP-3, which remove SUMO modifications from target proteins,

results in the redistribution and reorganization of PML NBs and associated proteins (Best et al.,

2002; Nefkens et al., 2003). Interestingly, some viruses have evolved mechanisms to prevent

SUMOylation of PML, thereby counteracting the anti-viral functions of the NBs (Adamson and

Kenney, 2001; Geoffroy and Chelbi-Alix, 2011).

Although PML proteins form the structural basis of the NBs and are essential for NB

formation, many other proteins either stably or transiently associate with PML NBs, sometimes

through SUMO/SIM interactions (Bernardi and Pandolfi, 2007). To date, over one hundred

proteins have been documented to either stably or transiently interact with PML NBs. Core

resident proteins that localize to typical PML NBs include the Speckled protein of 100 kDa

(Sp100) and death-associated dead protein (Daxx) (van Damme et al., 2010).

PML NBs can significantly change in size, number, and position in both normal cellular

conditions and in response to stress conditions (Dellaire and Bazett-Jones, 2004). PML proteins

are dynamic components of the NBs that exchange from the nucleoplasm to NBs or between

NBs, with the majority of the PML protein pool (more than 90% in most cell lines) not being

NB-bound (Wiesmeijer et al., 2002; Lallemand-Breitenbach et al., 2001). During cell-cycle

progression, PML NBs change profoundly in number. There are few PML NBs in G0 and the

numbers increase as cells enter G1 and proceed to S and G2 phases (Dellaire et al., 2006; Koken

et al., 1995; Terris et al., 1995; Chang et al., 1995; Kurki et al., 2003). In early S phase, PML

NBs fragment into smaller bodies by fission mechanisms and become distorted. In G2,

consequently, there are twice as many NBs as there were in G1 phase (Dellaire et al., 2006). In

response to cellular stresses, such as heat shock and heavy metals, PML NBs redistribute into

microspeckles (numerous small nuclear dots) (Nefkins et al., 2003; Eskiw et al., 2003; Maul et

al., 1995). Similarly, transcriptional inhibition, exogenous nucleases, and some DNA damaging

agents (such as UV irradiation and alkylating agents) cause the dispersal of PML NBs into

micro-NBs. Conversely, other DNA damaging agents, such as γ-irradiation, result in an increase

in the number of PML NBs (Dellaire and Bazett-Jones, 2004; Salomoni et al., 2005; Seker et al.,

2003; Conlan et al., 2004; Carbone et al., 2002; Dellaire et al., 2006). The ability of PML NBs

to respond to many different cellular conditions and stresses highlights its importance as a

tumour suppressor.

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1.1.2 Regulation of PML proteins and PML NBs

PML NB regulation is important to determine appropriate levels of cell proliferation and

survival in response to stimuli (Salomoni et al., 2008). Furthermore, regulation of appropriate

levels of PML proteins and their accumulation in NBs is crucial, since loss of PML NBs

accompanies malignant transformation and/or tumor progression (Gambacorta et al., 1996;

Gurrieri et al., 2004). The ubiquitin/proteasome-mediated turnover of the PML proteins is an

important factor in regulating PML-associated functions, but little is known about how PML

proteins are recognized and what cellular factors regulate them. Scaglioni et al (2006) have

identified casein kinase 2 (CK2) as an important factor in regulating PML protein levels by

promoting its ubiquitin-mediated degradation, dependent on direct phosphorylation at serine

517. The E3 ligase responsible for PML polyubiquitination and subsequent proteasomal

degradation remains elusive. However, the two SUMO-targeted E3 ubiquitin ligases RNF4 and

Arkadia have been identified as regulators of PML upon arsenic trioxide treatment. Arsenic

trioxide triggers an initial polySUMOylation of PML, which allows RNF4 and Arkadia to bind

and polyubiquitinate PML proteins, leading to proteasomal degradation (Lallemand-Breitenbach

et al., 2008; Erker et al., 2013). Most of the factors known to regulate PML NBs in the absence

of arsenic trioxide are viral proteins (see section 1.1.4).

1.1.3 PML NB associated functions

PML NBs have a variety of cellular functions. For example, they can function as a depot

for nuclear proteins (e.g. SUMO and Daxx) waiting to be used or degraded. In support of this is

the observation that the suppression of Daxx properties correlates with its accumulation at PML

NBs (Lehembre et al., 2001). This may be a mechanism to fine tune protein levels that function

in the nucleoplasm. PML NBs can also act as a scaffolding site for post-translational

modifications of some proteins (e.g. p53 phosphorylation and acetylation by homeodomain

interacting protein kinase 2 (HIPK2) and creb binding protein (CBP), respectively). Moreover,

PML NBs are also known to non-randomly associate with a subset of genes and have been

implicated in regulating their expression (both activation and repression) (Negorev et al., 2001;

D’Orazi et al., 2002; Pearson et al., 2000; Ching et al., 2005; Dellaire and Bazett-Jones, 2004;

Ching et al., 2013).

Since the link between PML and the pathogenesis of APL was discovered, PML has

been implicated in having tumour suppressive functions in a variety of human cancers. Notably,

PML-null mice, which are viable and whose development appears normal, are significantly

7

more susceptible to tumorigenesis and to infections by at least some viruses (e.g. lymphocytic

choriomeningitis virus (LCMV) and vesicular stomatis virus (VSV) infections) (Wang et al.,

1998; Bonilla et al., 2002; Geoffory and Chelbi-Alix, 2011). In the following sections, I will

highlight the roles of PML NBs involved in tumor suppression and antiviral defense as these

functions are particularly relevant for my thesis.

1.1.3.1 Tumour suppressive functions

Numerous observations have implicated PML NBs as being involved in the DNA

damage response, an important pathway in protecting cells against cancer. These studies have

shown that following DNA damage, PML NBs increase in size and number in an ATM- and

ATR-dependent manner, PML NBs are closely juxtaposed with persistent DNA repair foci, and

PML is phosphorylated after genotoxic stress (Dellaire and Bazett-Jones, 2004; Carbone et al.,

2002; Varadaraj et al., 2007; Dellaire et al., 2006a; Dellaire et al., 2009; Bischof et al., 2001;

Bischof et al., 2002). Moreover, DNA damage results in many DNA repair and checkpoint

proteins dynamically localizing to PML NBs (e.g. BLM helicase, and ATR and CHK2 kinases)

(Dellaire and Bazett-Jones, 2004). In fact, one of the mechanisms by which PML NBs affect

DNA damage signaling may be through post-translational modification and/or sequestration and

release of checkpoint proteins. Specifically, p53 accumulation and activation at PML NBs is

observed following DNA damage (Hofmann et al., 2002; D’Orazi et al., 2002). CHK2 also

phosphorylates PML to activate a p53-independent apoptosis pathway (Yang et al., 2002). In

turn, PML is required for ATM-mediated activation of CHK2 by promoting its

autophosphorylation (Yang et al., 2006). Finally, PML may also play a direct role in the DNA

repair and recombination process (Boe et al., 2006).

Cellular replicative senescence, a state that permanently excludes the cell from the cell

cycle, is another important barrier against tumorigenesis and can be caused by an induction of

the DNA damage response (Bartkova et al., 2006; Mallette et al., 2007; Campisi et al., 2007).

PML NBs were first implicated in senescence following the observation that overexpressing

PML results in a cell cycle arrest in cancer cell lines (He et al., 1997; Le et al., 1996; Mu et al.,

1997). Overexpression of the PML IV isoform alone has also been shown to induce replicative

senescence (Bischof et al., 2002; Mallette et al., 2004). The role of PML in senescence is

associated with an increase in pRb levels, potentiating the suppressive functions of pRb (Khan

et al., 2001). PML NBs have also been shown to regulate the formation of senescence-

associated heterochromatin foci (SAHFs), domains that repress the expression of growth-

8

promoting genes (Narita et al., 2003). Chromatin remodeling, involved in SAHFs formation, is

mediated by PML NBs, since the histone chaperone HIRA involved in this remodeling must

translocate to NBs to initiate this process (Zhang et al., 2005).

PML NBs also contribute to tumour suppression by regulating apoptosis. PML-null mice

are protected from multiple apoptotic stimuli due to PML’s role in regulating several pro- and

anti-apoptotic factors (Wang et al., 1998). PML is important in both p53-dependent and p53-

independent apoptotic pathways. The activation of p53 occurs when p53 associates with PML

NBs, enabling its acetylation and phosphorylation. PML also increases the level of p53 by

binding and inhibiting the E3 ligase Mdm2, the main negative regulator of p53 (Dellaire and

Bazett-Jones, 2004; Bernardi and Pandolfi, 2003; Takahashi et al., 2004; Bernardi et al., 2008).

A specific interaction between p53 and the PML isoform IV has also been detected (Fogal et al.,

2000).

UV-induced apoptosis is p53-independent but is dependent on the pro-apoptotic factor c-

Jun. UV irradiation causes the dispersal of PML NBs into microspeckles that colocalize with

active, phosphorylated c-Jun (Bernardi et al., 2008; Salomoni et al., 2005). PML can also

contribute to Fas-induced apoptosis. FLICE-associated huge protein (FLASH) is a positive

regulator of Fas-induced apoptosis and localizes to PML NBs under steady-state conditions. In

response to Fas activation, FLASH is released from PML NBs and accumulates in the

mitochondria where it activates caspase-8 (a vital cysteine protease involved in apoptosis)

(Milovic-Holm et al., 2007). PML-null cells are resistant to Fas-induced apoptosis, suggesting

that FLASH may require modification/activation at PML NBs before being able to activate

caspase-8 (Wang et al., 1998). Finally, TRADD (TNF-R1 associated death domain protein) is

another adaptor protein, similar to FLICE, implicated in apoptosis induced by death receptors

(Morgan et al., 2002). TRADD localizes to PML NBs, works as an adaptor molecule to recruit

caspase-8 and FADD to the TNF-R1 DISC (death-inducing signaling complex), and can induce

apoptosis from the cytoplasm and from within the nucleus (Kischkel et al., 2000). Importantly,

TRADD-induced apoptosis in the nucleus is both PML- and p53-dependent but does not require

caspases (Morgan et al., 2002). Therefore, PML NBs are strongly implicated in multiple facets

of the apoptosis pathway.

1.1.3.2 Antiviral functions

In addition to the role of PML NBs in suppressing tumour formation/progression, PML

NBs also have important antiviral functions via the interferon pathway. Interferons (IFNs)

9

establish an antiviral state in cells by conferring resistance to infection at multiple stages of viral

replication, including entry, transcription, RNA stability, initiation of translation, maturation,

assembly, and release (Geoffroy and Chelbi-Alix, 2011). IFNs mediate their effects through

IFN-upregulated cellular proteins, which include PML, p53, and Sp100 (a PML NB resident

protein). In fact, IFNs are the best known inducers of PML, resulting in a sharp increase in both

PML mRNA and protein levels, and a drastic increase in PML NB size and number (Everett and

Chelbi-Alix, 2007; Chelbi-Alix et al., 1995). PML NBs have been implicated in suppressing

lytic viral infections, since PML knock out mice are more susceptible to several virus infections,

and since there have been numerous studies showing that silencing of PML enhances the

replication of several viruses (Bonilla et al., 2002; Blondel et al., 2002; Iki et al., 2005; Li et al.,

2009). It has also been well established that many viruses express proteins that counteract

various stages in the IFN pathway and many viruses also encode proteins that disrupt PML NBs

(see section 1.1.4), perhaps as a strategy to evade IFN action (Regad et al., 2001; Everett and

Chelbi-Alix, 2007). The antiviral activities of PML NBs and examples of how some RNA and

DNA viruses have evolved to overcome these defenses are discussed below.

1.1.3.2.1 PML and RNA viruses

One of the mechanisms by which PML NBs inhibit RNA virus infection is to induce

apoptosis in a p53-dependent manner. Human immunodeficiency virus type 1 (HIV-1) leads to

PML aggregation in syncytia of the brain, lymph nodes, and a population of blood leukocytes of

infected individuals, as well as in syncytia induced by the HIV-1 envelope glycoprotein

complex in vitro (Perfettini et al., 2009). HIV-1-induced syncytia trigger an apoptotic pathway

that requires PML for the phosphorylation of ATM (which colocalizes to PML NBs) (Perfettini

et al., 2009). Therefore, PML activation is a crucial event that contributes to the apoptotic

demise of HIV-1-elicited syncytia.

Poliovirus is another RNA virus that is counteracted by PML-mediated apoptosis.

During poliovirus infection, p53 and PML cooperatively act to inhibit viral replication. In

response to poliovirus infection ERK is activated as part of the innate immune response, which

triggers PML phosphorylation. This modification, required for poliovirus-induced PML

SUMOylation, results in a transfer of PML from the nucleoplasm to the nuclear matrix and an

increase in PML NB size (Pampin et al., 2006). Consequently, these events recruit p53 to PML

NBs, where it is activated by phosphorylation on Ser15 in a PML-dependent manner leading to

apoptosis and inhibition of viral replication. These events are augmented upon the

10

overexpression of PML III and abrogated when PML or p53 are silenced (Pampin et al., 2006;

Geoffroy and Chelbi-Alix, 2011).

1.1.3.2.2 PML and DNA viruses

Many DNA viruses that replicate in the nucleus have an innately close relationship with

PML NBs. This is exemplified by the observations that viral genomes and/or their replication

compartments are closely associated with PML NBs, the large DNA viruses (e.g. adenovirus

and herpesviruses) encode proteins that localize to and/or disrupt PML NBs, herpesvirus

disruption of NBs directly correlates with an increase in viral gene expression, and herpesvirus

genomes that fail to disrupt PML NBs are sensitive to cell-mediated viral repression (reviewed

in Everett and Chelbi-Alix, 2007). Obvious connections of PML NBs and the IFN pathway with

viral genome repression are also evident (Geoffroy and Chelbi-Alix, 2011).

Live cell imaging of herpes simplex virus 1 (HSV-1) provided the first evidence that

PML NBs move toward viral genomes as they enter the nucleus, thereby actively targeting the

virus to prevent replication (Everett and Murray, 2005). PML’s repression of lytic viral

replication is immediately counteracted by the HSV-1 protein ICP0 (discussed in detail in

section 1.1.4). PML’s ability to suppress HSV-1 infection are further demonstrated in the

observations that ICP0-null HSV-1 mutants have a greatly increased probability of becoming

repressed. These repressed viral genomes are subsequently maintained in a quiescent state that

resembles the viral genome during latency (Everett et al., 2004; Preston, 2000; Preston and

Nicholl, 1997; Samaniego et al., 1998). Moreover, silencing PML proteins leads to increases in

both gene expression and plaque formation in ICP0-negative HSV-1 mutants (Everett et al.,

2006). In addition to actively targeting and suppressing HSV-1 genomes, PML NBs have also

been shown to sequester newly assembled varicella zoster virus (VZV) nucleocapsids in

neurons, satellite ganglion cells, and skin cells by forming spherical cages that enclose the

nucleocapsids (Reichelt et al., 2011).

Most recently, PML NBs have been demonstrated to be important in repressing the

reactivation of latent Epstein-Barr virus (EBV) to the lytic cycle in epithelial cells, as silencing

of PML was shown to be sufficient to induce the EBV lytic cycle in latently infected cells

(Sivachandran et al., 2012b). Moreover, treating EBV positive cells with arsenic trioxide,

known to disrupt PML NBs, leads to EBV reactivation/lytic protein expression and confers

susceptibility to the lytic-phase-specific antiviral drug gancyclovir (Sides et al., 2011). It is also

well established that several DNA and RNA viruses encode proteins that colocalize with PML

11

NBs and disorganize them, suggesting that viruses have evolved the ability to disrupt PML NBs

as a strategy to evade host antiviral mechanisms (Everett and Chelbi-Alix, 2007). Taken

together, these observations strongly implicate PML in the host antiviral response. Specific

examples of viral proteins that disrupt PML NBs are discussed below.

1.1.4 Viral proteins that disrupt PML NBs

Given the roles of PML NBs in promoting apoptosis and in suppressing lytic viral

infections, it is not surprising that some viruses have evolved mechanisms to induce the loss or

disrupt the functions of PML NBs (Table 1). Replication of the single-stranded RNA genome

of rabies virus occurs entirely in the cytoplasm, and yet this virus disorganizes PML NBs. The

rabies viral protein P delocalizes PML III from NBs into cytoplasmic dots where both proteins

colocalize (Blondel et al., 2002). Although protein P alters the localization of PML III, recent

studies suggest that only overexpression of PML IV confers resistance to rabies virus infection

(Blondel et al., 2010). Lymphocytic choriomeningitis virus (LCMV) is another RNA virus that

alters the localization of PML from the nucleus to the cytoplasm. This effect is mediated by the

nonstructural RING finger protein Z, which interacts with endogenous PML. This complex

interacts with elongation factor eIF4E in the cytoplasm, reducing its Cap-binding activity and

leading to the inhibition of translation (Borden et al., 1998; Kentsis et al., 2001). Three influenza

A viral proteins (M1, NS1, and NS2) have been shown to colocalize with PML proteins (Iki et

al., 2005). Although the functional significance of this localization has not yet been determined,

expression of different nuclear isoforms of PML (i.e. PML III, IV, or VI) significantly reduces

the rate of influenza A viral propagation (Chelbi-Alix et al., 1998; Iki et al., 2005; Li et al.,

2009).

Table 1. Viral proteins that alter or disrupt PML NBs through PML proteins

Virus Viral Protein Effect Adenovirus E4orf3 Reorganizes PML into tracts HCMV (Human cytomegalovirus)

IEI Disrupts NBs, é IE genes

HSV-1 (Herpes simplex virus)

ICP0 Degrades PML, é replication

ICP27 Alters PML transcription

EBV (Epstein-Barr virus)

BZLF1 Disrupts NBs

EBNA1 Facilitates NB degradation

HPV (Human E6 ê PML IV, senescence

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papilloma virus) E7 ê PML IV, senescence

KSHV (Kaposi’s sarcoma associated herpesvirus)

LANA2 Degrades NB

Murine γHV68 ORF 75c Disrupts NB, degrades PML, é replication

HFV (Human foamy virus)

Tas ê TAS activation

VSV (Vesicular stomatitis virus)

Not determined PML over-expression, ê viral replication

Influenza A M1, NS1, NS1 PML over-expression, ê viral replication

HDV (Hepatitis D virus)

L-HDAg Reorganizes PML NBs

RSV (Respiratory syncytial virus)

Not determined Cytoplasmic redistribution

HCV (Hepatitis C virus)

Core protein ê PML IV-induced apoptosis

Rabies virus Protein P Reorganizes PML III to cytoplasm

LCMV (Lymphocytic choriomeningitis virus)

Protein Z Reorganizes PML to the cytoplasm

*Note: many viruses also encode proteins that are able to alter/disrupt PML NBs through interactions with PML NB-associated proteins

DNA viruses, such as the herpesvirus family, have also evolved mechanisms to disrupt

PML NBs. HSV-1, an α-herpesvirus subfamily member, provided the first documented example

of a virus that disrupted PML NBs during infection via the immediate-early protein ICP0 (Maul

et al., 1993). ICP0, a tegument protein possessing E3 ubiquitin ligase activity, localizes to PML

NBs early during HSV-1 infection and induces the proteasome-mediated degradation of PML

NBs and Sp100 (Boutell et al., 2002; Everett et al., 1998; Chelbi-Alix and de The, 1999;

Geoffroy and Chelbi-Alix, 2011). ICP0-defective HSV-1 viruses are unable to degrade PML

NBs at early times in infection, and consequently this results in repression of viral gene

expression and a greatly reduced lytic infection (Everett et al., 2006). ICP0 was recently shown

to have two distinct mechanisms for targeting PML; the first is through SUMO-modifications

(and thus it can interact with/affect all PML isoforms), while the second is through a PML I-

specific interaction (SUMO-independent) (Cuchet-Lourenco et al., 2012).

Human cytomegalovirus (HCMV), a member of the β-herpesvirus subfamily, causes

PML NB reorganization by preventing or removing SUMO modifications on PML via the

immediate-early protein IE1 (Lee et al., 2004). An IE1-mutant HCMV displays a similar

13

phenotype to the ICP0-defective HSV-1, in that failure to disrupt PML NBs correlates with

increased repression of viral genome replication (Gawn and Greaves, 2002). Two members of

the γ-herpesvirus subfamily, EBV and Kaposi’s sarcoma herpesvirus (KSHV), also disrupt PML

NBs through their BZLF1 and LANA2 proteins respectively (Adamson and Kenney, 2001;

Marcos-Villar et al., 2009). BZLF1 acts to disperse PML from PML NBs by reducing the

amount of SUMO1-modified PML. It is likely that BZLF1 outcompetes PML for a limiting

amount of SUMO, as BZLF1 itself was shown to be SUMO modified (Adamson and Kenney,

2001). LANA2 increases the levels of SUMO2-ubiquitin-modified PML, which induces the

disruption of NBs via a proteasome-dependent mechanism (Marcos-Villar et al., 2009). LANA2

is SUMOylated and contains a SIM motif, both which have been shown to be important for

PML NB disruption. Mutating either the SIM motif or the SUMOylated lysine of LANA2

abrogates PML disruption abilities and it is therefore likely that SUMO modifications bridge

interactions between LANA2 and other SUMO-modified proteins required for PML disruption

(Marcos-Villar et al., 2009). It is evident that the herpesvirus family have evolved as experts of

PML NB disruption in order to facilitate effective viral infection.

Another DNA virus known to affect PML NBs is adenovirus, whose infection results in

the reorganization of PML NBs from punctate foci into elongated nuclear tracts (Puvion-

Dutilleul et al., 1995). The E4orf3 protein mediates this effect by specifically targeting PML II.

Mutants of E4orf3 unable to interact with PML II could no longer cause PML NB

rearrangement (Hoppe et al., 2006). Furthermore, an adenovirus mutant lacking E4orf3 was

compromised for efficient viral replication due to its inability to alter PML, and silencing of

PML proteins restored the replication abilities of the E4orf3 mutant (Ullman et al., 2007).

Of particular interest for my thesis is the effect of the EBV protein Epstein-Barr nuclear

antigen 1 (EBNA1) on PML NBs. EBNA1 has been shown to facilitate the disruption of PML

NBs through the degradation of PML proteins in epithelial cells (i.e. nasopharyngeal carcinoma

(NPC) cells and gastric carcinoma (GC) cells) (Sivachandran et al., 2008; Sivachandran et al.,

2012a). The degradation of PML proteins correlates with a loss in PML NB-associated

functions. For example, lower steady-state p53 levels and less acetylated p53 following DNA

damage are observed in gastric carcinoma cells that have lower PML NB levels due to either

EBV or EBNA1 expression (Cheng et al., 2010; Sivachandran et al., 2012a). For a detailed

discussion, see section 1.2.2.2 Cellular functions of EBNA1.

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1.2 Epstein-Barr virus (EBV)

EBV was discovered over 45 years ago from examining electron micrographs of cells

cultured from an endemic pediatric cancer common in sub-Saharan Africa, later to be known as

Burkitt’s lymphoma. Anthony Epstein and Yvonne Barr identified and characterized the viral

particles found in the cancer specimens (Epstein et al., 1964a; Epstein et al., 1964b). Due to this

analysis EBV was classified as a γ-herpesvirus, a herpesvirus subfamily that also includes

KSHV (Epstein et al., 1964a; Young and Rickinson, 2004). Far from having a restricted

distribution, EBV was later found to be widespread in all human populations and persists in the

vast majority of individuals as a lifelong, asymptomatic infection in B-lymphocytes (Young and

Rickinson, 2004). In fact, EBV is so successful that it infects over 90% of the adult human

population. Most individuals are infected as children (before age 3), but newly infected

adolescents or adults commonly present with infectious mononucleosis, also known as the

kissing disease (Rickinson and Kieff, 2001; Gerber et al., 1972). Importantly, latent infection of

the virus can induce cell proliferation and immortalization. Therefore, latency has been

recognized as a causative factor in the development of various cancers. Cancers that are the

most highly recognized as having EBV as a causative factor are nasopharyngeal carcinoma

(NPC), gastric carcinoma (GC; approximately 10% of total GC), Burkitt’s lymphoma,

Hodgkin’s lymphoma, and a variety of lymphomas in AIDS and organ transplant patients as a

result of immune suppression (reviewed in Young & Rickinson, 2004; Frappier, 2013). For

these reasons, EBV latent infection has become an active area of research.

1.2.1 Overview of the EBV life cycle

EBV spreads through saliva and initially infects epithelial cells of the oropharynx

producing virions that subsequently infect underlying B-lymphocytes (Rickinson and Kieff,

2001). These B cells act as the main site of EBV persistence, and through various stages of

latency, a stable reservoir of resting viral-genome positive memory B-cells are established

(Thorley-Lawson and Gross, 2004; Young and Rickinson, 2004). Latent infection is defined by

viral genome maintenance in the absence of virion production. Four main types of latency have

been described in B cells (Thorley-Lawson and Gross, 2004). Latency stages I, II, and III (also

known as EBNA1-only, default, and growth latency programs) occur in proliferating cells and

involve the expression of specific viral proteins, whereas latency 0 occurs in resting cells in

which no viral proteins are expressed (Table 2) (Thorley-Lawson, 2001; Thorley-Lawson and

Gross, 2004).

15

Table 2. Epstein-Barr virus latency programs

Latency Program EBNAs LMPs Detected in

0 --- --- Healthy individuals

I (EBNA1 only) EBNA1 --- BL

II (Default) EBNA1 LMP1, 2A NPC Ψ, GCΨ*, Hodgkin’s disease

III (Growth) EBNA1, 2, 3, A, B, C, and LP

LMP1, 2A, 2B EBV-associated diseases in immunocompromised individuals

ΨBARF1 also expressed in NPC and GC *EBV-positive gastric carcinoma display a latency-II-like profile, except LMP1 is not expressed

Latency program III is involved in the initial infection of naïve B-cells in the resting

state; growth transformation occurs to activate these cells to become proliferating blasts. A

combination of viral protein expression (i.e. six EBV nuclear antigens and three latent

membrane proteins) immortalizes B-cells to become lymphoblastoid cell lines (LCLs)

(Rickinson and Kieff, 2001). The EBNA’s, other than EBNA1, are highly immunogenic and

thus cells expressing this latency program are commonly found in immunocompromised

patients, as well as in vitro. It is believed that cells can transition between latency stages, and

thus, following latency III, EBV infection transits into latency I or 0 in order to avoid detection

by the immune system (Thorley-Lawson and Gross, 2004; Thorley-Lawson and Allday, 2008;

Young and Rickinson, 2004). In latency I, proliferating memory cells only express EBNA1,

which is the only viral protein necessary for viral genome maintenance (i.e. replication and

segregation of the viral episomes) in the dividing cells (Yates, 1996). Latency I is commonly

found in healthy individuals as well as in Burkitt’s lymphoma (Rickinson and Kieff, 2001). The

default program, latency II, expresses some latent membrane proteins, in addition to EBNA1.

Latency II has been associated with NPCs, GCs, and Hodgkin’s lymphoma (Rickinson and

Kieff, 2001). It is evident that EBV-associated tumours have distinct viral gene expression

profiles correlating with the different latency programs I, II, and III. In addition to the various

viral gene expression patterns observed, all latency programs express EBERs (double-stranded,

nuclear, non-coding viral RNA molecules) and viral micro RNAs. The functional role of these

molecules remains poorly defined and is an active area of study (Young and Rickinson, 2004).

The lytic form of EBV infection is required for the production of viral progeny and thus

responsible for cell-to-cell spread and transmissions from host to host. Lytic reactivation from

latency can be caused by multiple stimuli. For example, in vivo, antigen stimulation of infected

16

memory B-cells, causing them to differentiate into antibody producing plasma cells, results in

lytic replication that can be detected in peripheral B-lymphocytes (Laichalk and Thorley-

Lawson, 2005). Differentiation in epithelial cells can also activate the lytic form of EBV

infection (Tovey et al., 1978; Young et al., 1991). EBV reactivation in cell culture systems,

however, can be caused by a variety of chemical agents, including phorbol ester, 12-0-

tetradecanoyl phorbol-13-acetate (TPA), sodium butyrate, calcium ionophoes, and B-cell

receptor stimulation (Faggioni et al., 1986; zur Hausen et al., 1978). In instances of reactivation,

two EBV immediate early genes, BZLF1 and BRLF1, are actively transcribed from the latent

viral genome (Biggin et al., 1987; Flemington et al., 1991). These regulators of lytic gene

expression regulate their own and each other’s promoters, and subsequently activate a cascade

of approximately 80 viral genes that includes early and late proteins (Liu and Speck, 2003;

Adamson et al., 2000; Flemington et al., 1991; Speck et al., 1997). Early viral genes encode

proteins important for viral genome amplification, in order to produce linear molecules for

packaging into virions. Late viral genes encode structural proteins important for forming the

viral particle (Amon and Farrell, 2005). This infectious viral particle may infect new epithelial

cells or B-cells or is shed in saliva and is able to infect new hosts (Murray and Young, 2001).

See Figure 1-3 for a schematic of the EBV life cycle.

Important for my thesis is the role of EBNA1 in manipulating the host cellular

environment. EBNA1 has an essential role in EBV latency, reflected in the fact that EBNA1 is

expressed in all forms of latency in proliferating cells. Moreover, it is the only viral protein

detected in all EBV-associated tumors, pointing to its importance in cancers caused by EBV

(Frappier, 2012).

1.2.2 Epstein-Barr nuclear antigen 1 (EBNA1)

EBNA1 was the first viral protein detected in EBV infected cells and is the only viral

nuclear protein expressed in both lytic and latent modes of infection (Reedman and Klein,

1973). EBNA1 has been shown to be required for the persistence of EBV genomes in latency

through contributions to episome replication and segregation (Yates et al., 1984; Yates et al.,

1985). In addition to maintaining a constant EBV genome copy number, EBNA1 regulates the

expression of itself and other latency genes (Rickinson and Kieff, 2001). This function requires

the ability of EBNA1 to bind specific DNA elements in the EBV genome (reviewed in Frappier,

2013). Increasing evidence also supports the notion that EBNA1 can manipulate the host

cellular environment to facilitate viral infection and cell immortalization.

17

Figure 1-3: Overview of the EBV life cycle. A) Primary infection. Incoming viruses establish lytic replication in the oropharynx, after which the virus spreads throughout the lymphoid tissues as a latent (latency III) growth-transforming infection of B cells. Many of these proliferating cells are removed by a T-cell response, but some cells escape and subsequently establish a stable reservoir of resting viral-genome-positive memory B cells. At this point, viral antigen expression is mostly suppressed (latency 0). B) Persistent infection. The memory B-cell reservoir (harbouring latent EBV) is subject to physiological controls governing memory B-cell migration and differentiation as a whole. Therefore, some of these EBV-positive cells are recruited to germinal centres, resulting in the activation of different latency programs, after which they might re-enter the reservoir as memory cells, or commit to plasma differentiation. Cells committed to differentiation may move to mucosal sites in the oropharynmx, and in the process, activate the lytic viral cycle. Virions produced at these sites might initiate lytic replication in epithelial cells, allowing shedding of infectious virions into the oropharynx. These new infectious viral particles can replenish the B-cell reservoir or be passed through the saliva to a new host; most of the virions, however, will be removed by the memory T-cell response. (Adapted from Young and Rickinson, 2004).

18

The exact size of EBNA1 varies in different viral isolates due to the varying length of

the Gly-Ala repeat; however, the most commonly studied version of the protein is 641 amino

acids in length (Baer et al., 1984; Rickinson and Kieff, 2001). EBNA1 has a variety of

functional domains outlined in Figure 1-4. The N-terminal and central regions contains

sequences critical for DNA replication, segregation, and transcriptional regulation (Wu et al.,

2002; Yates and Camiolo, 1988). The C-terminal domain is important for DNA binding and

dimerization and is essential for interactions with elements in the EBV genome (e.g. the dyad

symmetry (DS) and the family of repeats (FR)) (Ambinder et al., 1990; Ceccarelli and Frappier

2000; Chen et al., 1993; Goldsmith et al., 1993; Rawlins et al., 1985; Wu et al., 2000; Wu et al.,

2002). Important for EBNA1’s manipulation of the host cellular environment is its ability to

interact with two cellular proteins, CK2 and USP7, mediated through regions 387-394 and 436-

450, respectively (Sivachandran et al., 2010; Saridakis et al., 2005).

1.2.2.1 Functions of EBNA1 on the EBV episome

Similar to replication in a typical eukaryotic cell, the EBV episome undergoes

replication once every cell cycle (Adams 1987). Analysis of different EBV DNA fragments that

would allow the replication of plasmids in cells latently infected with EBV identified the latent

origin of DNA replication, termed oriP (Yates et al., 1984). Replication of oriP-plasmids was

shown to require only EBNA1 and no other viral proteins (Yates et al., 1985). OriP contains

two functional elements, the dyad symmetry (DS) and the family of repeats (FR). These

elements each contain multiple copies of an 18 bp palindromic sequence that are bound by

EBNA1 (Rawlins et al., 1985; Reisman et al., 1985). The DS element contains four EBNA1

binding sites and is sufficient for replication of oriP-plasmids in the presence of EBNA1

(Harrison et al., 1994; Wysokenski and Yates 1989; Yates et al., 2000).

EBNA1 lacks any enzymatic activities but recruits cellular replication proteins to oriP

(Frappier and O’Donnell, 1991). Both the cellular origin replication complex (ORC) and the

minichromosome maintenance (MCM) complex were found to associate with the DS element

(Chardhuri et al., 2001; Dhar et al., 2001; Schepers et al., 2001). EBNA1 likely recruits ORC to

the DS, which in turn recruits the MCM complex, as EBNA1 and EBNA1 binding sites in the

DS have been shown to be important for ORC association (Dhar et al., 2001; Julien et al., 2004;

Schepers et al., 2001). Another factor that EBNA1 recruits to the DS is the nucleosome

assembly protein template activating factor (TAF-1). TAF-1 negatively regulates replication

19

Figure 1-4: EBNA1 structural organization. A schematic of the functional regions of EBNA1 is shown and amino acid numbers are indicated. The DNA binding domain, shown in purple, is important for EBNA1 binding to the EBV genome. Regions 61-83 (green) and 325-376 (red) are important for transactivation, and region 325-376 is important for genome segregation. The nuclear localization signal is depicted in black, although EBNA1 mutants with specific deletions in this area are still able to enter the nucleus. EBNA1 regions known to mediate interactions with CK2 and USP7 are also indicated.

20

from oriP, most likely through effects on chromatin structure (Wang and Frappier, 2009).

Notably, the EBNA1 DNA binding domain is not sufficient to activate oriP replication; regions

8-67 and 325-376 are also required and make redundant contributions to replication function

(Ceccarelli and Frappier, 2000; Wu et al., 2002).

Mitotic segregation of EBV episomes is crucial to maintain a stable copy number of the

genome in dividing cells. This function requires two viral components, EBNA1 and the FR

element in oriP (Lupton and Levine, 1985; Krysan et al., 1989; Lee et al., 1999). EBNA1 is

tightly associated with the condensed cellular chromatin in mitosis, and evidence indicates that

EBNA1 functions in segregation by tethering the EBV episome to the mitotic chromosomes.

Both EBV episomes and oriP-containing plasmids have been shown to associate with mitotic

chromosomes, and this association is dependent on EBNA1 and on the ability of EBNA1 to

attach to the chromosomes (Simpson et al., 1996; Harris et al., 1985; Delecluse et al., 1993;

Grogan et al., 1983; Petti et al., 1990; Kanda et al., 2001; Kapoor et al., 2005). The central Gly-

Arg repeat region of EBNA1 (amino acids 325-376) is critical for both mitotic chromosome

attachment and segregation function (Shire et al., 1999; Wu et al., 2000; Wu et al., 2002).

EBNA1 binding to the FR element is also important for transcriptional activation of

some EBV latency genes (Rickinson and Kieff, 2001). EBNA1 binding to FR specifically

enhances the transcription of the Cp promoter (from which all EBNA genes are transcribed) and

the promoter controlling LMP1 expression (Sugden and Warren, 1989; Wysokenski and Yates,

1989). Two regions of EBNA1 (65-83 and 325-376) are critical for this activity and mediate

host protein interactions that may impact transcriptional activities. Region 65-83 binds

bromodomain protein 4 (Brd4), a protein that interacts with chromatin to regulate transcription

(Lin et al., 2008; Wu and Chiang, 2007). Region 325-376 mediates interactions with P32/TAP

and two nucleosome assembly proteins implicated in transcriptional regulation, NAP1 and TAF-

1 (Wang et al., 1997; Holowaty et al., 2003; Park and Luger, 2006; Wang and Frappier, 2009).

The importance of EBNA1 transactivation is exemplified in the observation that an EBV mutant

defective in EBNA1 transactivation activities, but not replication or segregation functions, was

severely impaired in its ability to transform cells (Altmann et al., 2006).

1.2.2.2 Cellular functions of EBNA1

The effects of EBNA1 on replication, segregation, and transcriptional activation were

originally thought to be its only functions. Increasing evidence over the past decade strongly

supports the notion that EBNA1 is also an effective manipulator of the host cellular

21

environment. One example of this is EBNA1’s ability to interfere with the function of p53, a

tumour suppressor that is commonly down-regulated by DNA tumour viruses in order to

promote cell survival and proliferation. The link between EBNA1 and p53 was first discovered

from proteomic experiments revealing that EBNA1 stably binds the ubiquitin specific protease 7

(USP7 or HAUSP) (Holowaty et al., 2003). USP7 binds and stabilizes p53 and Mdm2 (the

major ubiquitin E3 ligase of p53) by removing polyubiquitin chains that signal degradation via

its deubiquitinating (DUB) activity (Cummins et al., 2004; Li et al., 2002; Li et al., 2004).

EBNA1 has a higher binding affinity for the N-terminal binding pocket on USP7 bound by p53

and Mdm2, and consequently EBNA1 outcompetes p53 and Mdm2, preventing their interaction

with USP7 (Holowaty et al., 2003; Hu et al., 2006; Saridakis et al., 2005; Sheng et al., 2006). In

keeping with these observations, EBNA1 expression in U2OS cells decreased the amount of p53

that accumulated in response to DNA damage, while a USP7 binding mutant of EBNA1 did not

have this effect (Saridakis et al., 2005). In addition, EBNA1 expression in CNE2 NPC cells

resulted in lower p53 levels following DNA damage (Sivachandran et al., 2008). Finally, the

expression of EBNA1 in all of the above cell lines resulted in decreased apoptosis following

DNA damage. This observation is likely in part due to EBNA1 binding USP7 and destabilizing

p53, but also in part due to EBNA1’s ability to disrupt PML NBs, a known site of p53 activation

(discussed below).

EBV latent infection or expression of EBNA1 on its own induces the degradation of

PML proteins in NPC and GC cells, leading to a loss of PML NBs and their functions

(Sivachandran et al., 2008; Sivachandran et al., 2012a). EBNA1 was shown to associate with

PML NBs and a specific interaction with PML isoform IV was also detected, as this isoform

was recovered by an EBNA1 immunoprecipitation more than any other isoforms (Sivachandran

et al., 2008; Sivachandran et al., 2012b). When analyzing cells expressing individual PML

isoforms (each of which forms NBs), EBNA1 had the greatest effect on PML IV NBs, again

suggesting that EBNA1 has an important interaction with this isoform (Sivachandran et al.,

2012b).

EBNA1’s ability to induce the degradation of PML NBs was subsequently shown to

require its interaction with two host proteins, protein kinase CK2 (casein kinase 2) and the

ubiquitin specific protease USP7 (or HAUSP), which also associates with PML NBs in

uninfected cells (Sivachandran et al., 2010; Sarkari et al., 2011; Everett et al., 1997). Silencing

of the catalytic subunit of CK2 (CK2α) or USP7 abrogates EBNA1’s ability to disrupt PML

NBs. Similarly, EBNA1 mutants that do not bind USP7 or CK2 fail to disrupt PML NBs

22

(Sivachandran et al., 2010 and Sarkari et al., 2011). Moreover, EBNA1 was shown to increase

the association of CK2 and USP7 with PML proteins and/or NBs. This increased association of

CK2 with PML proteins increased phosphorylation of PML proteins at serine 517 by CK2 (a

known trigger for polyubiquitination and proteasomal degradation) (Sivachandran et al., 2010;

Scaglioni et al., 2006). EBNA1 was shown to bind directly to the CK2β regulatory subunit of

CK2 and does so without disrupting the CK2 holoenzyme. EBNA1 interacts with CK2β and

USP7 through distinct sequences and can bind both proteins simultaneously (Sivachandran et

al., 2010). Taking the above observations together, it is likely that during EBV infection,

EBNA1 shifts the equilibrium by increasing the amount of both CK2 and USP7 that is localized

to PML NBs, which in turn increases PML phosphorylation, polyubiquitination, and

degradation.

The effects of EBNA1 on apoptosis, DNA damage, and p53 acetylation have also been

investigated, since PML NBs are known to be important for these processes. EBNA1 was found

to decrease apoptosis, DNA repair, and p53 acetylation in response to etoposide-induced DNA

damage, consistent with a loss of PML NBs and their functions (Sivachandran et al., 2008;

Sivachandran et al., 2012a). These results indicate that cells expressing EBNA1 may be more

susceptible to DNA damage, which could contribute to cell transformation in epithelial

carcinomas. Importantly, PML staining in EBV-positive and EBV-negative gastric carcinoma

biopsy specimens revealed greatly reduced PML levels in the presence of EBV, verifying the

effect of EBV latent infection on PML NBs in gastric carcinomas (Sivachandran et al., 2012a).

As discussed earlier, PML NBs retain antiviral properties and are suppressive for lytic

viral replication of some herpesviruses (HSV-1 being the most studied example). EBNA1 is

known to play various important roles in the context of latent EBV infection; however, the

reason for its continued expression during the lytic cycle was unknown. The possibility that

EBNA1’s function in the lytic cycle is related to its modulation of PML NBs was recently

investigated by Sivachandran et al (2012b) in EBV-positive GC cells. Two roles for EBNA1 in

EBV reactivation from latent to the lytic cycle were determined. First, EBNA1 silencing

positively contributed to spontaneous EBV reactivation, suggesting EBNA1 may repress

reactivation in latency. Second, upon induction of the lytic cycle, EBNA1 silencing decreased

viral genome amplification and lytic gene expression, suggesting EBNA1 positively contributes

to lytic infection. This effect was shown to be mediated by EBNA1’s disruption of PML NBs,

as EBNA1 did not promote the lytic cycle when PML proteins were silenced. In keeping with

this conclusion, silencing of PML NBs or arsenic trioxide disruption of PML NBs is sufficient

23

to induce EBV reactivation to the lytic cycle (Sivachandran et al., 2012b; Sides et al., 2011).

Notably, the ability of EBNA1 to induce PML NB degradation has only been observed in

epithelial cells, and thus, promoting the lytic cycle through PML NB disruption may be a

primary function of EBNA1 in epithelial cells that is responsible for virion production for host-

to-host spread.

It is evident that EBNA1 plays essential roles in EBV infection and EBV-associated

tumours. On one hand, the ability of EBNA1 to directly maintain the viral episome and allow

for continued expression of EBV genes most likely makes a major contribution to cell

transformation. On the other hand, increasing evidence suggests EBNA1 itself may be playing a

direct role in modulating the host cellular environment (i.e. promoting the survival of

transformed cells via interactions with USP7 and PML), thereby contributing to tumorigenesis.

1.3 Ubiquitin specific protease 7 (USP7)

Of particular interest to my thesis is the role of USP7 in regulating PML NBs, and I will

therefore provide a brief introduction to USP7. USP7 was discovered as a binding partner of the

HSV-1 protein ICP0 (Meredith et al., 1994). Additional studies revealed that this ICP0-

associated protein had conserved motifs belonging to the ubiquitin specific protease (USP)

family, and thus, USP7 was originally named herpesvirus associated ubiquitin specific protease,

or HAUSP (Everett et al., 1997). ICP0’s interaction with USP7 was shown to be important in

protecting itself from autoubiquitination and degradation (Boutell et al., 2005; Canning et al.,

2004). USPs remove ubiquitin moieties from target substrates, reversing the actions of ubiquitin

ligases, and consequently act to stabilize proteins (Nijman et al., 2005).

1.3.1 Structural organization and catalytic activity

USP7 is usually considered to have 3 domains (two protein interactions domains

separated by a catalytic domain), as depicted in Figure 1-5 (Holowaty et al., 2003). The catalytic

domain of USP7 is responsible for its deubiquitinating (DUB) activity and contains a conserved

triad of Cys, His, and Asp residues common to all USPs (Hu et al., 2002). An amino acid point

mutation in the catalytic cleft of USP7 at cysteine 223 (to serine, i.e. C223S) has been shown to

abrogate its DUB activity (Li et al., 2002). Regions upstream and downstream from the central

catalytic domain mediate protein-protein interactions and provide substrate recognition. For

example, the N-terminal TRAF domain mediates interactions with EBNA1, p53, Mdm2, and

Mdmx (Saridakis et al., 2005; Zapata et al., 2001; Li et al., 2002; Hu et al., 2002; Holowaty et

24

Figure 1-5: Schematic of USP7 structural organization. USP7 is thought to have 3 domains, the N-terminal TRAF domain (NTD) shown in blue, the catalytic domain (CAT) shown in red, and a C-terminal domain (CTD) containing 5 ubiquitin-like (UBL) motifs arranged in 2-1-2 units. Known protein interactors of the NTD and UBL domains are indicated, as well as amino acid numbers. The C223S mutation site that abrogates USP7’s deubiquitinating (DUB) activity is also indicated.

25

al., 2003; Hu et al., 2006; Sheng et al., 2006; Sarkari et al., 2010). The C-terminal domain of

USP7 contains five ubiquitin-like domains (UBLs) organized into 2-1-2 UBL units. The last di-

UBL unit is sufficient to activate USP7 through binding a switching loop component of the

catalytic domain, promoting ubiquitin release (Faesen et al., 2011). The C-terminal domain has

also been shown to mediate protein interactions with ICP0, FOXO4, and GMP synthetase (van

der Horst et al., 2006; Holowaty et al., 2003b; Faesen et al., 2011), the latter of which stimulates

the ability of USP7 to cleave monoubiquitin from histone H2B (Sarkari et al., 2009; van der

Knaap et al., 2005).

1.3.2 Cellular functions of USP7

The p53 tumour suppressor acts as a transcription factor to activate and express genes

involved in apoptosis and cell-cycle arrest and, therefore, provides an essential network

protecting cells against cancer. USP7 has been found to regulate the p53 pathway in multiple

ways. Initially, USP7 was found to stabilize p53 through a specific interaction that promoted its

deubiquitination, resulting in p53-dependent cell-cycle arrest and apoptosis in response to DNA

damage (Li et al., 2002). Additional studies proved to complicate USP7’s role in p53 regulation.

Decreased USP7 expression had the expected effect and was found to destabilize p53, whereas a

complete knockdown of USP7 was found to have the opposite effect, as p53 stabilization was

observed (Li et al., 2004). This unexpected effect can be attributed to the ability of USP7 to

specifically bind and stabilize Mdm2, the main E3 ligase responsible for ubiquitinating p53

(Cummins et al., 2004; Cummins and Vogelstein, 2004; Li et al., 2004). Furthermore, USP7 can

bind, ubiquitinate, and stabilize Mdmx, an Mdm2 homologue that inhibits transcription

activation by p53 (Meulmeester et al., 2005). ATM/ATR-mediated phosphorylation of Mdm2

and Mdmx, following DNA damage, disrupts their interaction with USP7 and leaves USP7

available to interact with and stabilize p53 (Meulmeester et al, 2005). Having the ability to

stabilize p53 and its negative regulators, USP7 offers a graceful way to adjust the levels and

therefore the activity of p53.

USP7 function extends beyond its involvement in the p53 pathway. For example, USP7

regulates the trafficking of the transcription factor Forkhead box O (FOXO4). FOXO4 is

monoubiquitinated in response to oxidative stress, which allows its nuclear translocation and

subsequent transcriptional function (van der Horst et al., 2006). As a potential feedback

mechanism, USP7 can remove ubiquitin from FOXO4 to counteract its nuclear translocation;

26

this promotes FOXO4 nuclear exclusion and inhibition of transcriptional activities (van der

Horst et al., 2006).

USP7 is also known to partially associate with PML NBs, originating from observations

by Everett et al (1997). Additional analyses demonstrated that, at least in some cell

backgrounds, ICP0 required its interaction with USP7 to disrupt PML NBs (Parkinson and

Everett, 2000). These were the first findings implying USP7 and PML may be functionally

linked. More recently, the finding that USP7 is used by EBNA1 to disrupt PML NBs also

suggested that USP7 itself regulates PML NBs. Indeed, additional studies in the absence of

EBV or EBNA1 showed that USP7 overexpression induced the loss of PML NBs, while USP7

silencing increased the level of PML NBs and PML proteins (Sarkari et al., 2011). The

observation that USP7 promotes PML degradation is counterintuitive, since USPs generally

stabilize proteins (Frappier and Verrijzer, 2011). However, it was shown that this effect of USP7

is independent of its DUB activity, since overexpression of a catalytically inactive USP7, or the

N- or C-terminal protein interaction domains of USP7, also induced the loss of PML NBs

(Sarkari et al., 2011). It is possible that USP7 recruits a negative regulator, perhaps an E3 ligase

of PML, to PML NBs. Similar to the observation that EBNA1 preferentially interacts with PML

IV, immunoprecipitation with USP7 also recovered more PML IV than any other PML

isoforms. Moreover, overexpression of USP7 in cells expressing individual PML isoforms was

found to decrease PML IV and PML I NBs, with the greatest effect on PML IV NBs (Sarkari et

al., 2011). Although the precise mechanism by which USP7 negatively regulates PML NBs

remains elusive, it is evident that its regulation is important and requires further investigation.

1.4 Thesis rationale

PML NBs are subnuclear structures that are important in a variety of tumour suppressive

and antiviral functions. Previous observations indicated that the replication of EBV plasmids by

EBNA1 was less efficient if PML proteins were silenced. Therefore, my first objective was to

determine the role of PML NBs in EBV episome replication. I present data demonstrating that

PML NBs do not have a suppressive role in episome replication. EBNA1 and USP7 have both

been determined to be negative regulators of PML NBs, in that they induce the loss of PML

NBs and degradation of PML proteins. Moreover, EBNA1 and USP7 preferentially interacted

with and affected PML isoform IV in cells. My second objective was to further investigate the

mechanisms by which EBNA1 and USP7 recognized PML proteins. I first present data

confirming the ability of USP7 to negatively regulate PML NBs by examining PML NBs in

27

USP7-null cells. Next, I examined the importance of serine 393 of EBNA1 (a site important for

CK2 interaction) in EBNA1-mediated PML NB disruption and determined that the ability of

this serine to be phosphorylated is crucial for EBNA1 function. Finally, I tested the hypothesis

that EBNA1 and USP7 directly interact with PML proteins in vitro. I provide data supporting

this hypothesis and show that EBNA1 and USP7 exhibit binding specificities to PML IV.

28

2 Materials and Methods

2.1 Cell lines

The NPC CNE2Z cell line (referred to here as CNE2) was derived from an EBV-positive NPC

but lost the EBV genome after growth in culture (Sun et al., 1992). CNE2 was maintained in

alpha minimal essential media (αMEM, Gibco) supplemented with 10% fetal bovine serum

(FBS). The generation of the CNE2-shPML cell line is described next (originally detailed in

Sarkari et al., 2011). pLKO.shPML1 expressing anti-PML shRNA (Everett et al., 2006) was

provided by Dr. Roger Everett and is described elsewhere (Cuchet et al., 2011). A lentivirus was

generated from this plasmid as previously described (Everett et al., 2008). One ml of filtered

culture medium containing the shRNA-lentivirus was added to 1x105 CNE2 cells with

polybrene (Sigma) at a final concentration of 8µg/µl, which was replaced after 24 hours with

medium containing 2µg/ml puromycin. 72 hours later, puromycin was removed and cells were

cloned by serial dilution and checked for PML expression by immunofluorescence microscopy

(described below) using an anti-PML antibody able to recognize all PML isoforms (PG-M3;

Santa Cruz). Western blotting (described below) was also used to confirm PML silencing. The

colon carcinoma cell lines HTC116 and HTC116-USP7-/- (provided by Bert Vogelstein;

Cummins et al., 2004) were cultured in Dulbecco’s modified Eagle’s medium (DMEM, Gibco),

supplemented with 10% FBS. The human embryo kidney HEK293T cell line was also

maintained in DMEM, supplemented with 10% FBS.

2.2 Plasmid construction

EBNA1 was expressed from the pc3oriP plasmid (referred to as pc3oriPE), which contains the

EBV oriP element, and the construction of these plasmids was previously described (Shire et

al., 1999). EBNA1 point mutants (i.e. EBNA1-S393A and EBNA1-S393T) were generated by

QuickChange site-directed mutagenesis (Stratagene) of pMZS3F-EBNA1 and were sequenced

to verify the mutation.

2.3 Transfections

To generate CNE2 and CNE2-shPML cells transiently expressing EBNA1 proteins (i.e.

EBNA1, EBNA1-S393A, or EBNA1-S393T), 1.5x105 cells were transfected with 2 µg

pc3oriPE plasmids in 6-well dishes. Polyjet transfection reagent (SignaGen) was used at a ratio

of 2:1 (polyjet:DNA). 293T cells were transfected with 8 µg DNA of individual PML isoform

29

constructs (pCIFlagPML) in 10 cm plates. Lipofectamine LTX (Invitrogen) transfection reagent

was used at a ratio of 2:1 (LTX:DNA).

2.4 Cell proliferation assay

The proliferation rates of CNE2 and CNE2-shPML cells following transfection were monitored.

Approximately 1x105 cells were seeded in 6 cm plates, followed by transfection with an oriP

plasmid the next day. Plasmid were transfected with Lipofectamine LTX (Invitrogen) in a 1:2

ratio (DNA:Lipofectamine). Cell cultures were split reseeded into 10 cm dishes when they

reached 90% confluency. The 6 cm plates were subsequently harvested at 1, 2, 4, 6, and 8 days

post-transfection and counted using a haemocytometer.

2.5 Transient replication assay

CNE2 and CNE2-shPML cells were transfected with either an oriP plasmid control (pc3oriP) or

an oriP plasmid that expresses EBNA1 (pc3oriPE), and then propagated for 72 hrs. Plasmids

were recovered from cells by Hirt’s method (Hirt, 1967). Cells were harvested by centrifugation

at 114x g for 10 mins and the cell pellet was resuspended in 350 µl of PBS. Cells were washed

with PBS, spun down to remove the supernatant, and resuspended in 350 µl of PBS. 350 µl of

2X Hirt’s solution (0.6% SDS, 10 mM EDTA, 10 mM Tris-HCl, pH 7.5 final concentration)

was added to samples. Samples were inverted 10 times to mix and incubated at 25oC for 10

mins. Following the incubation, 140 µl of 5 M NaCl was added and samples were incubated

overnight at 4oC. The next day, samples were subjected to centrifugation for 1 hour at 21,130x g

at 4oC. Supernatants were then transferred to microfuge tubes containing 6 µl of RNase A (10

mg/ml, Fermentas) and incubated at 37oC for 1 hour. 8 µl of Proteinase K (20 mg/ml,

Fermentas) was added and samples were incubated at 50oC for two hours. Samples were then

extracted with phenol:chloroform followed by chloroform and DNA was precipitated with

isopropanol. To linearize the DNA, samples were resuspended in 30 µl of Xho I digestion buffer

containing10 units of Xho I (New England Biolabs) and incubated for 3 hours at 37ºC. 9/10ths

of the linearized DNA was then incubated with 10 units Dpn I (New England Biolabs) for 3

hours to digest the input plasmid that had not undergone replication. The remaining 1/10th of the

DNA was used as input control for the efficiency of plasmid recovery. Finally, plasmid samples

were separated on 0.8% agarose gel electrophoresis, transferred to Hybond-XL membranes

(Amersham) as described below, and probed with 32P-labelled pc3oriP. Bands were visualized

30

by autoradiography and signals were quantified by PhosphorImager analysis using the

ImageQuant software.

2.6 Plasmid labeling

300 ng of linearized pc3oriP and 8 µg of random hexamer primer (Roche) were diluted to 10 µl

with dH20 and incubated at 95oC for 5 mins, followed by a 5 min incubation on ice. The mixture

was brought to a final volume of 25 µl by adding 10 mM Tris-HCl, 50 mM NaCl, 10 mM

MgCl2, 1 mM DTT pH 7.9, 50 µM dNTP mix (dATP, dTTP, and dGTP), 3 units of Klenow

DNA polymerase, and 5 µl of 3000 Ci/mmol α32P-dCTP. This final mixture was incubated at

37oC for 1.5 hours. The labeled plasmid was passed through an IllustraTMMicrospinTM G-25

column. Plasmids were then denatured by incubating at 95oC for 5 mins, followed by 5 min on

ice.

2.7 Southern blot

Plasmid samples from the transient replication assays were separated on a 0.8% agarose gel

overnight at 30V. The gel was soaked in 0.25 M HCl for 10 mins (to allow for DNA

depurination), washed in dH20, and then DNA fragments were denatured by soaking the gel in

0.4 M NaOH/0.6 M NaCl for 30 mins. DNA fragments were then transferred to a Hybond-XL

membrane in the same solution for 1 hour using a vacuum pump. Next, a 20X SSC solution (3M

NaCl, 0.3 M sodium citrate, pH 7) was added for 1 hour, and then membranes were washed in

2X SSC solution and air dried. Dry membranes were then incubated in blocking buffer (50%

formamide, 5X Denhardts, 0.5% sodium dodecyl sulphate, 6X SSC, and 0.2 mg/ml sheared

salmon sperm DNA) in a hybridizing oven for 1.5 hours at 42oC. Blocking buffer was removed

and replaced with new blocking buffer containing the 32P-labelled pc3oriP probe (5x106 cpm in

15 ml of buffer), for an overnight incubation at 42oC. The next day, membranes were washed

twice for 10 mins at room temperature in 1% SDS and 2% SSC, then twice more in the same

solution for 30 mins at 65oC. The final wash was in 0.1% SDS and 0.2% SSC for 10 mins at

room temperature. Membranes were subsequently dried and subjected to autoradiography

overnight. Radiolabelled DNA bands were quantified by PhosphorImager analysis using

ImageQuant software.

31

2.8 Immunofluorescence microscopy

Cells grown on coverslips were fixed with 3.7% formaldehyde in phosphate buffered saline

(PBS) for 20 mins, washed twice in PBS and then permeabilized with 1% Triton X-100 in PBS

for 5 mins at room temperature. Samples were blocked with 4% bovine serum albumin (BSA) in

PBS, followed by an hour incubation with primary antibodies against EBNA1 (R4 rabbit serum

at 1:1000 dilution (Holowaty et al., 2003)) and PML (Santa Cruz PG-M3 at 1:100 dilution) in

4% BSA. Samples were washed three times with PBS and then incubated for an hour with

secondary antibodies goat anti-rabbit Alexafluor 555 (Molecular Probes) and goat anti-mouse

Alexfluor 488 (Molecular Probes) in 4% BSA. Coverslips were subsequently mounted onto

slides using ProLong Gold antifade medium containing DAPI (Invitrogen). Images were

obtained using the 40x oil immersion objective on a Leica inverted fluorescent microscope and

processed using OpenLab (ver.X.0) software. PML NBs were quantified by counting all visible

PML foci in 100 cells.

2.9 Western blots

Cells were lysed in 9 M urea, 5 mM Tris-HCl (pH 6.8), sonicated briefly and clarified by

centrifugation for 5 minutes at 15 000 rpm in a microcentrifuge. 50 µg of total protein was

subjected to 10% SDS-PAGE and transferred to a nitrocellulose membrane. Membranes were

blocked in 5% non-fat dry milk in PBS, then incubated with primary antibodies against PML

(Bethyl, A301-167A; 1:2000 dilution), USP7 (Bethyl, A300-033A; 1:10 000 dilution), EBNA1

(R4; 1:5000 dilution), or actin (Santa Cruz, I-19; 1:20 000 dilution). Membranes were washed in

PBS with 0.1% Tween 20 three times. After washing, blots were probed with secondary

antibodies goat anti-mouse peroxidase (Santa Cruz, SC-2055; 1:5000) or goat anti-rabbit

peroxidase (Santa Cruz, SC-2004; 1:5000). Blots were washed again in PBS with 0.1% Tween

and then developed using chemiluminescence reagents (ECL, Perkin Elmer).

2.10 EBNA1 and USP7 purification

EBNA1 (lacking most of the Gly-Ala repeat region) was expressed as a hexahistidine fusion

from a baculovirus and purified from insect cell nuclei as previously described (Frappier and

O’Donnell, 1991). Thirty 15 cm plates of High Five cells at 80% confluency were infected with

His-tagged EBNA1 baculovirus for 72 hours. Cells were harvested, washed with PBS (at 4oC),

and resuspended on ice in 50 ml of hypotonic buffer (20 mM HEPES pH 7.5, 1 mM MgCl, 1

mM PMSF) using a Dounce homogenizer and pestle B. Nuclei were collected by centrifugation

32

at 1000x g for 10 mins at 4oC, washed twice in 50 ml of cold hypotonic buffer, and resuspended

in 25 ml lysis buffer (20 mM HEPES pH 7.5, 1 M NaCl, 1% Nonidet P-40, 10% glycerol, 1 mM

MgCl2, and protease inhibitor p8849 (Sigma)) with the Dounce homogenizer and pestle B,

followed by a 30 min incubation on ice. The nuclear extract was clarified by centrifugation at 34

000x g for 1.5 hours at 4oC. This lysate was then incubated with 500 µl of nickel resin

(QIAGEN) in 500 µl of lysis buffer for 1 hour at 4oC on a rotator, and then transferred to a

column. The resin was washed 5 times with 5 column volumes of the nuclear lysis buffer

containing 250 mM imidazole. EDTA and DTT were immediately added to the elutions, up to a

final concentration of 10 mM. The eluted protein was dialyzed overnight against buffer A (50

mM HEPES pH 7.5, 1 M NaCl, 10% glycerol, 0.5 mM EDTA, and 0.1 mM DTT) and the

purified EBNA1 was stored in aliquots at -80oC. Full length USP7 was previously purified by

Teresa Sanchez-Alcaraz according to Holowaty et al., 2003.

2.11 In vitro binding assay

10 cm plates of 293T cells at 80% confluency were transfected with 8 µg of pCIFlagPML

plasmid expressing FLAG-tagged full-length PML isoforms I, IV, or VI (from Beech et al.,

2005). The next day, cells in each 10cm plate were moved to two 15cm plates, and cells were

harvested two days later. Harvested cells were washed in PBS and then resuspended in 1 ml of

lysis buffer (150 mM NaCl, 30 mM ZnCl2, 50 mM Tris, 1 mM EDTA, 1 % Triton, pH 7.4;

protease inhibitor P8340 (Sigma) and SUMO protease inhibitor NEM (Sigma) were added fresh

to the buffer for each use) per plate of cells. Cells were incubated in the lysis buffer for 30 mins

on ice, followed by centrifugation at 12, 000 x g for 1 hour at 4oC. The supernatant was

transferred to a new tube and stored at -70oC. 1 ml of the prepared lysate was incubated with 30

µl of anti-FLAG M2 resin (A2220, Sigma) for 2-3 hours at 4oC. Samples were washed five

times in lysis buffer containing 500 mM NaCl to disrupt any interactions from the cell lysate.

Purified EBNA1 (0.11-4.5 µg titration) or USP7 (0.26-2.1 µg titration) were then incubated with

the resin in lysis buffer containing 250 mM NaCl for 2-3 hours. Samples were washed five

times with the same 250 mM NaCl buffer and then PML and associated proteins were eluted

with two 15 min washes of 4x bead volume of 0.85 M ammonium hydroxide. Eluted proteins

were lyophilized and resuspended in 2X SDS loading dye. 1/10th of the input and the flow

through, along with the entire elution were analyzed by SDS-PAGE and Western blotting for

EBNA1, USP7, and PML.

33

3 Results

3.1 Contributions of PML NBs to EBNA1-mediated DNA replication

Boichuk et al (2011) recently demonstrated that the simian virus 40 (SV40) initiates

replication near PML NBs and that PML silencing inhibits viral replication. This indicated that

PML NBs might have a positive role in some viral replications. Moreover, an initial observation

by a previous student in our laboratory suggested that the replication of EBV-based plasmids by

EBNA1 is less efficient if the cells are silenced for PML. In the context of EBV, this suggests

that, while PML bodies are suppressive for lytic infection, they may positively contribute to

latent infection. Using the NPC cell lines CNE2 and CNE2-shPML (PML silenced cells), we

sought to further investigate if the absence of PML would decrease the amount of EBV genome

replication.

In order to assess the amount of replication from oriP in conditions with and without

PML NBs, CNE2 and CNE2-shPML cells were transfected with two different oriP plasmids,

oriP, which should not replicate, and oriP-EBNA1, which should replicate due to the presence

of EBNA1. OriP-EBNA1 plasmids will replicate once every cell doubling and, therefore, to

accurately compare relative amounts of plasmid replication, the cell doubling times of CNE2

and CNE2-shPML cells were determined first. The number of cells was counted with a

haemocytometer on days 1-8 (cell proliferation assay) following transfection (Figure 3-1, A). I

determined that CNE2-shPML cells were growing more slowly than CNE2 cells. Therefore, in

order for cells to undergo the same number of cell doublings, CNE2 cells were harvested on day

3, whereas CNE2-shPML cells were harvested on day 4. Plasmids from the cell lysates were

isolated by Hirt’s method (Hirt, 1967) and linearized. 90% of each sample was treated with the

Dpn I restriction enzyme to determine plasmid replication, while the remaining 10% was used

as an indication of plasmid yield (i.e. input). Plasmids that have replicated at least once are

resistant to Dpn I digestion, whereas the parental transfected plasmid will be digested by Dpn I.

Plasmids produced in E. coli are methylated on both DNA strands due to the presence of Dam

methylase and, consequently, Dpn I will cleave the DNA at its methylated GATC recognition

sequence. Mammalian cells do not contain Dam methylase and, thus, following one plasmid

replication in CNE2 or CNE2-shPML cells, the plasmids are hemi-methylated and resistant to

Dpn I cleavage. The amount of oriP-plasmid replication was analyzed by Southern blotting

(Figure 3-1, B). Lanes 1-4 and 7-10 indicate 10% of the plasmid input. As expected and due to

the absence of EBNA1, oriP plasmids were not able to replicate and were completely digested

34

by Dpn I treatment (lanes 5,7 and 13,15). Plasmid replication in the presence of EBNA1 (lanes

6,8 and 12,14) was similar with and without PML, suggesting that PML NBs have no obvious

effect on latent replication. The DNA bands in experiment 2 were normalized to the input

samples and quantified by Image Quant. This quantification revealed that CNE2-shPML cells

exhibited only a 24% decrease in the amount of plasmid replication (Figure 3-1, C) and

confirmed that PML NBs do not positively or negatively contribute to oriP-plasmid replication.

Note that while the amount of plasmid replication in lane 8 (without PML) does appear less than

lane 6 (with PML), the Dpn I portion of lane 8 is also less compared to lane 6 and most likely

accounts for the difference observed.

3.2 Analysis of PML NBs in USP7-null cells

We have previously demonstrated that, in addition to EBNA1’s regulation of PML NBs,

USP7 alone can negatively regulate PML (Sarkari et al., 2011). Importantly, both

overexpressing and silencing USP7 demonstrated this negative regulation. USP7’s regulation of

p53 has been shown to be different when USP7 is down regulated by siRNA as compared to

when it is completely absent in USP7-knock out cells, in that p53 is destabilized upon USP7

down regulation but stabilized in USP7-null cells (Li et al., 2002; Li et al., 2004). Since the

effect of a complete knockdown of USP7 on PML NBs has never been investigated, I examined

how USP7 affected PML NBs in WT HCT116 cells and HCT116 USP7-/- cells. First I

investigated the number of PML NBs by immunofluorescence microscopy with PML-specific

antibody (Figure 3-2, A). The cell nuclei are DAPI stained and shown in blue, and PML NBs are

shown in green. The number of PML NBs in WT cells was counted by determining all visible

NBs in fifty cells; on average there were 4.6 NBs per cell. The number of PML NBs in HCT116

USP7-/- cells could not be accurately quantified by IF, as most of the cells had greater than 30

NBs per cell. There is an obvious increase in the number of PML NBs in HCT116 USP7-/- cells

(i.e. the amount of green staining) as compared to control WT HCT116 cells (Figure 3-2, A).

Moreover, many of the PML NBs in the USP7-null cells appear much larger than NBs in the

WT control.

Next, I determined the differences in PML protein levels by western blotting. I compared

equal amounts of whole cell lysate from each cell line using antibody that recognizes all PML

isoforms and antibody against actin as a loading control. I found that the HCT116 USP7-/- cells

have a striking increase in the levels of PML proteins compared to control WT HCT116 cells

(Figure 3-2, B). Taken together, these results support our previous findings that USP7 is a

35

A

Figure 3-1: PML silencing does not reduce plasmid replication efficiency in CNE2 cells. (A) Proliferation assay comparing the doubling times of CNE2 and CNE2-shPML cells post-transfection with an oriP plasmid for 8 days. (B) Replication assay in CNE2 and CNE2-shPML (sh) cells comparing the bands from oriP (negative control without EBNA1) and oriP-EBNA1 plasmids after 3 cell doublings (3-4 days). (C) DpnI resistant bands were normalized to input lanes, and then the values from experiment 2 were plotted on a bar graph. The values are shown relative to the positive control (oriPE in CNE2), which was set to 100%.

0  

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shPML

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Input

EBNA1 - + - + - + - + - + - + - + - +

0"

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OriP CNE2Z

OriPE CNE2Z

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Experiment 1 Experiment 2

Lane: 1 2 3 4 5 6 7 8 7 8 9 10 11 12 13 14

36

negative regulator of PML. Moreover, USP7’s regulation of PML NBs is dissimilar to its p53

regulation, as downregulating USP7 with siRNA yields the same effect on PML NBs as

complete USP7 knockout.

3.3 Contribution of Serine 393 in EBNA1 to PML NB disruption

The only known functional role of the EBNA1-CK2 interaction is in EBNA1-mediated

PML disruption (Sivachandran et al., 2010). EBNA1 recruits CK2 to PML NBs, resulting in an

increase in CK2-mediated phosphorylation of PML proteins, signaling their polyubiquitination

and degradation (Sivachandran et al., 2010, Scaglioni et al., 2006). CK2’s ability to bind to

EBNA1 was recently shown to be dependent on the phosphorylation of EBNA1 S393 (a CDK

site) (Cao et al., submitted). An S393A mutation disrupts CK2 binding, while S393T (which can

also be phosphorylated by CDK) partially restores it. Importantly, an S393D mutation that

reconstitutes the negative charge is not sufficient to restore binding to CK2, indicating that

phosphorylation is critical for the interaction (Cao et al., submitted).

In order to demonstrate a functional role of this EBNA1 phosphosite, I transiently

expressed EBNA1, EBNA1-S393A, and EBNA1-S393T in CNE2 cells (the cell line that has

previously been used to characterize EBNA1’s regulation of PML) and examined the effects on

PML NBs by immunofluorescence microscopy (Figure 3-3, A). The cells were stained with

antibodies specific to PML (shown in green) and EBNA1 (shown in red), and nuclei were

visualized by DAPI staining (blue). PML NBs were quantified by counting all visible NBs in

100 cells expressing the various EBNA1 constructs for two independent experiments. This

quantification was graphed as the average number of PML NBs per cell for all cells counted

(Figure 3-3, B). The number of PML NBs in cells expressing the EBNA1-S393A mutant was

very similar to cells not expressing EBNA1 (negative control). On the other hand, EBNA1-

S393T was able to reduce the number of PML NBs to levels almost as low as WT EBNA1

(positive control). Therefore, the S393A mutation abrogated the ability of EBNA1 to disrupt

PML NBs, whereas the S393T mutation largely restored this ability. These results accentuate

the functional importance of the EBNA1-CK2 interaction for EBNA1-mediated PML

disruption, as the effects of the EBNA1 mutants on PML NBs correlates with their ability to

interact with CK2.

37

Figure 3-2: HCT116 USP7 -/- Cells Have Increased PML NBs Compared to HCT116 WT. HCT116 WT and HCT116 USP7-/- cells were analyzed by immunofluorescence microscopy for the number of PML NBs (A). Cell nuclei are DAPI stained (blue) and PML NBs are depicted as green foci within the nuclei. The amount of PML protein was also compared in HCT116 WT and HCT116 USP7-/- cells by western blotting (B). Actin was used as a protein loading control.

HCT116 WT

HCT116 USP7-/-

A

HCT116 W

T

HCT116 U

SP7-/-

Actin

PML

B PML Overlay with DAPI

38

Figure 3-3: The ability of EBNA1 to disrupt PML nuclear bodies is abrogated by the S393A mutation and restored by the S393T mutation. (A) CNE2Z cells were transfected with plasmids expressing EBNA1 or the EBNA1 S393A or S393T mutants. Cells were then stained with antibodies against PML (green) and EBNA1 (red) and counter stained with DAPI (blue). The number of PML nuclear bodies per cell was counted for 100 cells in two independent experiments. (B) The average number of PML nuclear bodies per transfected cell is shown in the bar graph along with standard deviations.

DAPI

WT S393A S393T

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39

3.4 Characterization of the direct interactions of EBNA1 and USP7 with specific PML isoforms

The specific PML isoforms important for the association of EBNA1 and USP7 with

PML NBs have been recently investigated. Cell lines in which total PML was silenced and

single silencing-resistant PML isoforms were introduced (and confirmed to form NBs) were

used to investigate the effect of EBNA1 on different PML isoforms. Expression of EBNA1 in

these cells was found to primarily cause loss of PML IV bodies, with a lesser effect on the other

PML isoforms (Sivachandran et al., 2012b). Silencing of USP7 was found to increase PML IV

and PML I NBs, with the greatest effect on PML IV NBs (Sarkari et al., 2011). This data is

consistent with the observations that immunoprecipitation of EBNA1 or USP7 recovers the

PML IV isoform more than any other PML isoform (Sivachandran et al., 2008 and Sarkari et al.,

2011). Although EBNA1 and USP7 preferentially disrupt and co-IP PML IV, similar effects but

to a lesser degree, are seen on PML I (Sivachandran et al., 2008 and Sarkari et al., 2011). The 8a

exon is a common feature of PML IV and PML I and may account for this similarity (Figure 1-

2). Exon 8, a unique region of PML IV that flanks exon 8a, may also be a contributing factor

since a binding preference is exhibited with this isoform. I hypothesize that EBNA1 and USP7

specifically interact with sequences in PML IV (i.e. exon 8a and 8) that are absent in other PML

isoforms.

I tested this hypothesis by determining, in vitro, if EBNA1 and USP7 interact directly

with PML isoforms I, IV, or VI. PML VI does not contain any of the C-terminal sequences of

PML I and IV and was used as a negative control for binding. My approach was to express

FLAG-tagged full-length PML proteins (from Beech et al., 2005) in human cells (by

transfection of 293T cells), then isolate them via the FLAG-tag to use in in vitro binding assays

with purified EBNA1 or USP7. Importantly, each transfected PML isoform is expressed at

extremely high levels, and thus, only a small proportion of the isolated PML isoform could be

complexed with other endogenous PML isoforms from the 293T cells. Once the PML protein

was bound to the anti-FLAG resin, the samples were washed several times in high salt buffer to

disrupt any interactions from the cell lysate (i.e. other PML isoforms and known PML

interactors). Following this, purified EBNA1 or USP7 was incubated with the resin in a lower

salt physiological buffer. The samples were then washed several times with this buffer and PML

and associated proteins were eluted with base.

In my initial experiments using high levels of EBNA1 and USP7, I detected direct

interactions of EBNA1 and USP7 with all PML isoforms tested (PML I, IV, VI) (Figure 3-4).

40

Figure 3-4: EBNA1 and USP7 Interact Directly with PML isoforms I, IV, & VI. Individual FLAG-tagged PML isoforms were expressed in 293T cells and isolated on anti-FLAG resin. PML I, IV, and VI were incubated with purified EBNA1 (8.1 µg) and/or USP7 (18 µg) and samples were compared by Western blotting for EBNA1 (A) and USP7 (B) binding. 10% of the input and flow through of purified EBNA1 or USP7 was loaded, and is compared to the entire protein elution. The amount of EBNA1 and USP7 binding to resin alone was analyzed (C) and the samples were also analyzed for the amount of PML bound to the resin (D).

41

Concurrently with this experiment, an EBNA1 plus USP7 mixing experiment was done whereby

both purified proteins were incubated with PML on the resin. As a starting point, equal molar

ratios of EBNA1 and USP7 were used in these experiments (e.g. 8.1 µg EBNA1 and 18 µg

USP7), although the actual ratio of these proteins at PML NBs in vivo is unknown. In the

presence of USP7, EBNA1 exhibited less binding to PML and, in contrast, USP7 binding to

PML in the presence of EBNA1 was not decreased (Figure 3-4). Importantly, any potential

background binding of EBNA1 and USP7 with the FLAG resin did not account for the

interactions observed, as EBNA1 and USP7 did not bind to the resin without PML present

(negative control) (Figure 3-4, C). In the experiments where EBNA1 and USP7 binding was

observed, PML proteins I, IV, and VI were all eluted from the FLAG resin (positive control),

indicating the interactions seen were indeed mediated by PML proteins (Figure 3-4, D).

Moreover, the amount of PML I protein eluted from the resin appears to be slightly increased

compared to the amounts of PML IV and VI, which are very similar; overall, the FLAG-resin

did retain similar amounts of PML proteins, allowing for an adequate comparison for isoform

specific binding.

In order to determine if EBNA1 and USP7 had preferential binding with a specific PML

isoform, I titrated back the amounts of protein incubated with PML on the resin. EBNA1 was

titrated from 4.3 µg to 0.11 µg, and I demonstrate that EBNA1 first loses binding to PML VI

and then to PML I, while still exhibiting binding specificity to PML IV even at the lowest

concentrations tested (Figure 3-5, A and D). Again, EBNA1 did not exhibit any background

binding to the FLAG resin that could account for these observations (Figure 3-5, B and E) and

PML proteins were eluted off the resin (Figure 3-5, C and F). Importantly, the relative amounts

of all PML isoforms that were eluted from the resin is very similar and thus serves as an

accurate comparison for binding affinities. Since both EBNA1 and PML can bind to DNA, I

tested the possibility that the interaction I observed was mediated through DNA. PML lysates

were incubated with benzonase for 30 mins to digest the RNA and DNA, before the lysates were

incubated with FLAG resin. In this condition, EBNA1 was still able to bind to PML IV (no

other isoforms were analyzed for binding) (Figure 3-5, G).

Similarly to EBNA1, when titrating back the amount of USP7 protein (from 2.1 µg to

0.26 µg), I have demonstrated a binding specificity for PML IV at the lowest concentrations

tested (Figure 3-6, A). Interestingly, USP7 appears to have a higher binding affinity to PML VI

when compared to PML I at 0.525 µg of USP7. These results are unexpected, since in vivo data

suggests that USP7 preferentially interacts with PML IV and PML I over all other isoforms.

42

43

Figure 3-5: EBNA1 titration demonstrates PML IV binding specificity. Individual PML protein isoforms were expressed in 293T cells and isolated back out via their FLAG tags. Decreasing concentrations (4.35µg, 1.74µg, and 0.87µg of EBNA1 for A; 0.28µg and 0.11µg of EBNA1 for C) of purified EBNA1 were incubated with various PML isoforms bound to FLAG resin, washed, and subsequently eluted. Samples were analyzed by Western blotting with antibodies against EBNA1 (A, B, D, E, G) and PML (C, F) proteins. (G) PML IV-transfected cell lysate was incubated with benzonase before binding PML to FLAG resin, followed by incubation with EBNA1. 10% of the input and flow through of purified EBNA1 was loaded, and is compared to the entire protein elution.

44

Figure 3-6: USP7 titration demonstrates PML IV binding specificity. Individual PML protein isoforms were expressed in 293T cells and isolated via their FLAG tags. Decreasing concentrations (2.1µg to 0.26µg in 2-fold dilution steps) of purified USP7 were incubated with the indicated PML isoforms bound to FLAG resin, washed, and subsequently eluted. Samples were analyzed by Western blotting with antibodies against USP7 (A and B) and PML (C) proteins. 10% of the input and flow through of purified USP7 was loaded, and is compared to the entire protein elution.

45

This observation may be accounted for by the fact that much more PML VI was eluted from the

FLAG resin than PML I, providing more PML VI for USP7 to bind to (Figure 3-6, B). USP7

does not bind to the FLAG resin without PML protein, again indicating that the interactions

observed are mediated through PML protein (Figure 3-6, C). This in vitro analysis demonstrates

that EBNA1 and USP7 can both interact with PML proteins and, furthermore, these results

support our in vivo data from which I conclude that EBNA1 and USP7 have binding

specificities for PML IV.

46

4 Discussion and Future Directions

4.1 Thesis summary

Studying the mechanisms by which pathogens manipulate the host cellular environment

can be useful in understanding host cell biology. In this instance, I have investigated the

mechanisms by which the EBV protein EBNA1 regulates the host tumour suppressor protein

PML, which is required for the formation of subnuclear structures known as PML NBs.

Previously, EBNA1 has been shown to disrupt PML NBs in epithelial cells, leading to a

degradation of the protein and a loss of the NBs’ functions. This has important consequences for

the host cell, as a loss of PML NBs facilitates viral infection and may contribute to

tumorigenesis. A previous observation in our laboratory suggested that, although PML NBs are

suppressive for lytic viral replication, they might positively contribute to latent episome

replication. I therefore sought to investigate the role of PML NBs in EBV genome replication by

analyzing oriP-plasmid replication in cells with and without PML NBs. I have shown that PML

NBs neither positively nor negatively contribute to oriP-plasmid replication.

The ability of EBNA1 to disrupt PML NBs was demonstrated to be dependent on its

binding to two host proteins, USP7 and CK2. USP7 itself was subsequently shown to be a

negative regulator of PML NBs, in that silencing or overexpressing USP7 led to an increase or

loss in PML protein and NBs, respectively. Moreover, USP7 is a known regulator of the p53

tumour suppressor. This regulation, however, is quite complex, since USP7 silencing versus a

complete USP7 knock-down yields different effects on p53 levels. Due to this complex

regulation of p53, I investigated USP7’s regulation of PML NBs in USP7-null cells. Consistent

with USP7 silencing experiments, I found that USP7-null cells have a greatly increased number

of PML NBs, as well as an increase in PML protein levels.

Jennifer Cao in our laboratory recently determined that phosphorylation of serine 393 of

EBNA1 (part of a CDK site) was particularly important for its interaction with CK2. Mutating

this site to alanine abrogated the CK2-interaction, whereas a mutation to threonine (which

restores the CDK site) partially restored binding. These studies led me to investigate the

functional implications of these mutants on EBNA1-mediated PML NB disruption. I provide

data demonstrating that the EBNA1-S393A mutant cannot disrupt PML NBs, whereas the

EBNA1-S393T mutant is able to disrupt PML NBs almost as well as WT EBNA1. This

functional analysis directly correlates with the ability of the EBNA1 mutants to bind CK2,

47

highlighting the importance of the EBNA1-CK2 interaction in EBNA1-mediated PML NB

disruption.

Multiple types of evidence also indicated that both EBNA1 and USP7 likely regulate

PML NBs through interactions with PML isoform IV. I used an in vitro approach with purified

proteins to determine if EBNA1 or USP7 could directly interact with any PML proteins, and if

so, what isoforms are important. I first provide data indicating that both EBNA1 and USP7 can

bind PML isoforms I, IV, and VI; however, upon titrating back the amounts of EBNA1 and

USP7 used in the binding assay, I demonstrate that these proteins have a specific binding

preference for PML IV. This data directly recapitulates the in vivo findings that first indicated

EBNA1 and USP7 preferentially interact with PML IV.

4.2 PML NBs do not contribute to oriP-plasmid replication

The interactions between PML NBs and viral genomes first sparked interest upon

observations that the parental genomes and/or replication complexes of HSV-1 and adenovirus

were closely associated with PML NBs (Ishov and Maul, 1996; Maul et al., 1996). Soon after

these initial discoveries, SV40, polyomavirus, and all herpesvirus subfamily members were also

shown to be associated with PML NBs (reviewed in Maul, 1998; Everett, 2001). PML NBs were

demonstrated to have suppressive functions on lytic viral replication directly or mediated

through the IFN response (Everett and Chelbi-Alix, 2007). In fact, many of the viruses listed

above encode proteins to disrupt PML NBs in order to facilitate efficient viral infection

(Geoffory and Chelbi-Alix, 2011).

Surprisingly, a recent observation indicated that PML NBs actually facilitate SV40

replication; viral replication was shown to be near PML NBs and PML silencing inhibited viral

replication (Boichuk et al., 2011). Although PML NBs suppress most viral infections, this

suggested that PML NBs could positively contribute to the replication of some viruses. An

initial observation by a previous student in our laboratory suggested that the replication of EBV-

based plasmids by EBNA1 is less efficient if the cells are silenced for PML. This warranted

further investigation since this suggested that, like SV40 replication, PML NBs could positively

contribute to EBV latent replication. Using oriP-plasmid replication as a model of EBV episome

replication, I examined plasmid replication in cells with and without PML, and with and without

EBNA1. OriP plasmids lacking EBNA1expression were used as a negative control for

replication, since EBNA1 is required to bind to the oriP element to facilitate replication. As

expected, oriP plasmid lacking EBNA1 did not replicate regardless of the presence of PML

48

NBs. In contrast, oriP-plasmids expressing EBNA1 did replicate, and this replication was

shown to be at most only slightly affected by the presence of PML NBs.

I observed that the CNE2-shPML cells were dividing more slowly than WT cells

following transfection, an important factor that was not considered in the initial experiment. In

order to accurately compare the amount of plasmid replication between CNE2 and CNE2-

shPML cells, it is important to ensure that each cell line has undergone the same number of cell

doublings. I then determined the appropriate transfection conditions to ensure the cells were

proliferating better and performed a cell proliferation assay to determine the cells doubling

times. For my experiment, I ensured that both CNE2 and CNE2-shPML cells were allowed to

undergo the same number of cell doublings. This discrepancy between CNE2 and CNE2-shPML

replication most likely accounted for the previous observation that less oriP-plasmid replication

occurred in CNE2-shPML cells. Although I have shown that PML NBs do not contribute much,

if at all, to EBV latent replication, previous data does implicate PML NBs as having an

important role in suppressing EBV lytic reactivation (Sivachandran et al., 2012b).

4.3 USP7-null cells have increased amounts of PML NBs and PML protein

Everett et al (1997) made the original observations that USP7 partially associates with

PML NBs in cells. The first functional link between USP7 and PML came from later

observations that the HSV-1 protein ICP0 requires an interaction with USP7 to disrupt PML

NBs in some cell backgrounds (Parkinson and Everett, 2000). With respect to latent EBV

infection, Sivachandran et al (2008) found that USP7 is used by EBNA1 to disrupt PML NBs.

Taken together, these results implicated USP7 in the normal regulation (i.e. not in the context of

a viral infection) of PML NBs. Indeed, Sarkari et al (2011) then demonstrated that USP7 alone

negatively regulates PML NBs. They found that silencing USP7 led to an increase in PML

protein and NBs, whereas overexpressing USP7 reduced the number of PML NBs.

USP7 is also a known regulator of the p53 tumour suppressor. This regulation, however,

is quite complex, since USP7 silencing results in p53 destabilization and a complete USP7

knock down causes p53 stabilization (Li et al., 2002; Li et al., 2004). Due to this complex

regulation of p53, I analyzed PML NBs in USP7-null cells to determine if completely knocking

down USP7 would yield a different result than what has been observed in USP7 silencing

experiments. Consistent with previous experiments, I have found that the levels of PML proteins

and NBs are greatly increased in USP7-null cells.

49

The mechanisms involved in USP7’s regulation of PML NBs remain elusive. Although a

scenario whereby USP7 stabilizes an E3 ubiquitin ligase that negatively regulates PML (e.g.

how USP7 stabilizes MDM2, which negatively regulates p53) is plausible, it is very unlikely

given that USP7’s DUB activity is dispensable for PML regulation (Li et al., 2004; Sarkari et

al., 2011). A more likely explanation is that USP7 recruits other negative regulators of PML to

PML NBs, or that USP7 directs PML proteins to the proteasome. In support of these scenarios,

USP7 has been shown to be able to interact with the E3 ligases MDM2 and MARCH7 (Li et al.,

2002; Nathan et al., 2008) and to associate with the 26S proteasome (Besche et al., 2009;

Bousquet-Dubouch et al., 2009). Moreover, my findings suggesting that USP7 can directly

interact with PML proteins (see section 4.5) further supports the possibilities that USP7 could

recruit negative regulators directly to PML, or USP7 could directly recruit PML proteins to the

proteasome.

With respect to future experiments, the HCT116 USP7-/- cell line that was utilized has

limited potential. Multiple attempts were made to transfect them with USP7 and EBNA1

expression plasmids; however, the cells became very sick upon transfection (data not shown).

An adenovirus delivery system was also utilized; however, the adenovirus itself affected PML

NB levels (data not shown). For these reasons, no future experiments will be done in the USP7-

null cells. Further experiments to investigate the mechanisms by which USP7 interacts with and

regulates PML proteins will be discussed in section 4.6.

4.4 Serine 393 of EBNA1 plays a critical role in EBNA1-mediated PML NB disruption

EBNA1-mediated PML NB disruption is dependent on the interaction with the host

protein CK2, a kinase known to be important in the normal regulation of PML NBs. CK2

phosphorylates PML proteins, triggering their polyubiquitination and subsequent proteasome-

mediated degradation (Scaglioni et al., 2006). Importantly, EBNA1 increases the association of

CK2 with PML proteins, resulting in an increase in PML phosphorylation by CK2

(Sivachandran et al., 2010). Another student in our laboratory (Jennifer Cao) performed alanine

scanning mutagenesis of EBNA1 on amino acids 387-394, a region previously shown to be

required for the EBNA1-CK2 interaction (Sivachandran et al., 2010). Serine 393 was shown to

be critical for the EBNA1-CK2 interaction (Cao et al, submitted). Moreover, S393 is the only

residue in the CK2-binding region that is a known phospho-site (Duellman et al, 2009; Kang et

al, 2011) and, while mutation of neighbouring serines partially affected CK2 binding, mutation

of S393 consistently reduced CK2 binding to undetectable levels (Cao et al, submitted).

50

Interestingly, an S393D mutation (which reconstitutes the negative charge of a phosphate) could

not rescue CK2 binding, whereas an S393T mutation (which reconstitutes the CDK phosphosite

and was shown to be phosphorylated) partially restored CK2 binding (Cao et al., submitted).

In order to determine the effects of serine 393 on EBNA1 function, I examined the only

known function of the EBNA1-CK2 interaction, the ability of EBNA1 to induce the degradation

of PML NBs. I found that EBNA1 S393A cannot disrupt PML NBs, whereas EBNA1 S393T

preserves the ability to disrupt PML NBs almost to the level of WT EBNA1. This data directly

correlates with the ability of each mutant to bind CK2 and also shows that the ability of EBNA1

to disrupt PML NBs can be regulated by EBNA1 phosphorylation. The importance of the

EBNA1-CK2 interaction is clearly exemplified in these findings.

4.5 EBNA1 and USP7 preferentially interact with PML isoform IV in vitro

The findings that EBNA1 and USP7 were both negative regulators of PML NBs

suggested that these proteins might interact with PML proteins to mediate their functions.

Indeed, both EBNA1 and USP7 co-immunoprecipitated PML proteins from cells, and moreover,

a preferential interaction with PML isoform IV was observed (Sivachandran et al., 2008; Sarkari

et al., 2011). When investigating the effects of these proteins in cells that express individual

PML isoforms (and therefore NBs composed of only one isoform), both EBNA1 and USP7

primarily affected PML IV NBs, with a lesser effect on all other PML isoforms (Sarkari et al.,

2011; Sivachandran et al., 2012b). Although EBNA1 and USP7 primarily affected PML IV,

partial effects were seen with PML I. Exon 8a is common between PML I and IV and might

account for the partial effects observed. It is likely that exon 8 (specific to PML IV and adjacent

to exon 8a) is particularly important, as EBNA1 and USP7 still preferentially bind and affect

PML IV. To determine if EBNA1 and USP7 could directly interact with PML proteins, or

whether the association observed was mediated through other proteins, I explored these

interactions in vitro.

I investigated the interactions of purified EBNA1 and USP7 with PML I and IV, due to

the observations that have indicated the importance of these isoforms. I used PMLVI as a

negative control for binding, since this isoforms lacks almost the entire PML C-terminus

(including exons 8 and 8a). Using high levels of EBNA1 and USP7, I initially found that these

proteins could interact with all PML isoforms examined. Concurrently with this experiment, and

again using high levels of EBNA1 and USP7, I tested the ability of these proteins to bind PML

when they were mixed together before incubating them with PML. I found that EBNA1 bound

51

PML less effectively in the presence of USP7, whereas USP7 bound PML equally as well with

or without EBNA1. These results suggest that EBNA1 and USP7 could be competing for the

same binding site on PML and that USP7 has a higher affinity for binding than EBNA1.

However, since PML NBs are complex structures consisting of many PML proteins (and

therefore many potential binding sites), it is also possible that EBNA1 and USP7 could bind to

different PML binding sites in vivo. Since high levels of EBNA1 and USP7 were used in this in

vitro experiment, the conditions of the experiment most likely do not represent the actual ratios

of EBNA1 and USP7 at PML NBs, and could explain the competitive binding observed. It is

also likely that the high levels of protein used in these experiments drove a binding reaction with

PML, not representative of in vivo interactions, which could account for the binding observed to

PML VI. I therefore titrated back the amounts of EBNA1 and USP7 to reassess their PML

interactions and to determine if I could recreate in vitro what we observe in cells.

Upon titrating back the amounts of EBNA1 used, I demonstrate that EBNA1 first loses

binding to PML VI and then to PML I. Moreover, even at the lowest concentration of EBNA1

tested I can still detect binding to PML IV, an observation that recapitulates what is seen in

cells. Upon titrating back the amounts of USP7, I found that USP7 first loses binding to PML I

and then to PML VI. Although this does not directly correlate with what is seen in cells, a

possible explanation for this discrepancy is that much more PML VI was retained on the FLAG

resin than PML I. Importantly, at the lowest amounts of USP7 tested, binding to both PML I and

VI is lost while binding to PML IV remains intact. Overall, my findings demonstrate that

EBNA1 and USP7 can directly interact with PML proteins and implicate PML IV as the most

important isoform for the EBNA1 and USP7 interaction.

4.6 Future directions

4.6.1 Determining how EBNA1 and USP7 interact with PML

I have shown that both EBNA1 and USP7 exhibit a binding preference for the PML

isoform IV in vitro, consistent with our previous data indicating that EBNA1 and USP7

preferentially co-IP PML IV and preferentially disrupt PML IV NBs. Further characterization

and mapping of the binding regions important for EBNA1/USP7’s interaction with PML IV will

be done by using the in vitro approach discussed above. First, the competition experiment

between EBNA1 and USP7 will be performed under the optimized conditions in which EBNA1

and USP7 exhibited PML IV binding specificity. Next, the binding assays will be conducted

with PML IV proteins with specific C-terminal deletions (e.g. exons 8 and 8a) to identify PML

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sequences bound by EBNA1 and USP7. Finally, the binding assays will be conducted with

EBNA1 and USP7 mutants to identify the sequences that mediate the PML interaction.

To support the in vitro binding data, the localization of both EBNA1 and USP7 WT and

mutant proteins to PML NBs will be investigated in vivo. It was recently shown that the Herpes

simplex 1 (HSV-1) ICP0 protein associates with PML NBs via two mechanisms involving two

different regions of ICP0: first, through PML I-specific interactions; and second, through an

interaction with the SUMOylated sites on PML that are common to all PML isoforms (Cuchet-

Lourenço et al., 2012). Taking into account the ability of the HSV-1 protein ICP0 to associate

with PML NBs in two different ways, it will be important to determine if EBNA1 is also able to

associate with PML NBs both specifically and non-specifically. Comparing the ability of

EBNA1 mutants to localize to WT PML NBs and PML IV NBs will reveal if EBNA1 has

multiple mechanisms of associating with NBs.

I have previously examined the localization of a variety of EBNA1 mutant proteins in

cells expressing WT PML NBs, and the EBNA1 452-641 mutant is the only protein tested that

does not localize to PML NBs (data not shown; full length EBNA1 is amino acids 1-641).

EBNA1 ∆40-376 can still localize to PML NBs, and therefore it is likely that the regions

mediating PML association are 1-40 or 376-452. EBNA1 mutants that have parts of the N-

terminus added back to the EBNA1 452-641 construct, specifically in regions 1-40 and 376-452,

will be screened for ability to restore localization. Once the EBNA1 regions mediating

localization have been determined with WT PML NBs, localization of the same mutants in cells

containing NBs only consisting of PML IV and NBs composed of PML IV mutant NBs will be

examined. The localization of USP7 (WT and mutants) to PML NBs (WT, IV, and IV-mutant

NBs) will also be examined. The in vitro binding and in vivo localization experiments provide a

comprehensive approach to determining the regions of EBNA1, USP7, and PML responsible for

the interactions observed.

4.6.2 Determining the role of autophagy in PML NB degradation

Upon observing EBNA1 localization to PML NBs in the presence of the proteasomal

inhibitor MG132, I found that EBNA1 was still able to disrupt PML NBs (Figure 4-1). This

finding suggests that EBNA1 may be inducing PML degradation via another mechanism of

protein turnover, perhaps in addition to proteasomal turnover. A likely candidate is the

autophagy pathway, particularly since the degradation of EBNA1 (at least in immune cells) is

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Figure 4-1: EBNA1 can induce PML NB disruption in CNE2Z cells in the presence of a proteasomal inhibitor (MG132). Cells were transiently transfected with a plasmid expressing EBNA1 (oriP-EBNA1) and treated with MG132 proteasomal inhibitor at 5 hours post-transfection and samples were fixed at 10 hours post-transfection. Cells were then stained with primary antibodies against EBNA1 (red) and PML (green), and counter stained with DAPI (blue). Secondary antibodies used to stain EBNA1 and PML were GAR 555 and GAM 488, respectively.

54

through autophagy, and thus EBNA1 could be bringing PML proteins with it through this

pathway (Paludan et al., 2005).

Autophagy, which literally means “self-eating”, can sequester proteins and organelles in

autophagosomal vesicles and fuse them to lysosomes for degradation, thereby providing an

alternative route for turnover of stable and defective cellular proteins (Figure 4-2). Autophagy is

initiated by the formation of a double-membrane autophagosome that surrounds cargoes. A

number of autophagy proteins (Atgs) are involved in the formation of two ubiquitin-like

conjugation systems involving Atg12 and the microtubule-associated protein 1 light chain 3

(LC3), that mediate autophagosome formation (Behrends et al., 2010).

Although proteasomal and autophagy-mediated lysosomal degradation use distinct

components, these pathways are not mutually exclusive. Inhibition of the proteasome can

stimulate autophagy activity, possibly acting as a compensatory regulation mechanism (Pandey

et al., 2007). Moreover, selective autophagy also requires polyubiquitination of cargo, whereby

the adaptor proteins (e.g. the sequestosome 1 (p62/SQSTM1) or the neighbor of BRCA1 gene

(NBR1)) bind both ubiquitin and LC3 and target ubiquitinated proteins to autophagosomes for

degradation (Pankiv et al., 2007). In this process, single proteins and various cellular structures

such as protein aggregates, peroxisomes, ribosomes, and mitochondria can be specifically

engulfed by autophagosomes (Kraft et al., 2009).

I hypothesize that EBNA1 is able to facilitate the degradation of PML proteins through

autophagy-mediated mechanisms. Therefore, the potential involvement of autophagy in

EBNA1-mediated PML degradation will be determined as follows:

The ability of EBNA1 to disrupt PML NBs in the presence of autophagy inhibitors

Bafilomycin, NH4Cl, and 3-methyladenine (3MA) will be investigated first. Bafilomycin and

NH4Cl block lysosomal acidification, whereas 3MA blocks autophagosome formation. These

inhibitors will be tested in CNE2 cells, a biologically relevant NPC cell line that is EBV-

negative, following EBNA1 overexpression by transient transfection. The inhibitors will also be

tested in CNE2E cells, which have been engineered to stably express EBNA1, and therefore

EBNA1 transfection is not required. The number of PML NBs and the amount of PML protein

will be examined by IF microscopy and western blotting, respectively. Second, the ability of

EBNA1 to disrupt PML NBs when key autophagic components are silenced (i.e. Atg12) will be

determined in both CNE2 and CNE2E cells. In CNE2 cells, Atg12 will be silenced followed by

transfection with an EBNA1 or control plasmid. Again, PML NBs will be examined by IF

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Figure 4-2: Schematic of autophagy. Various stimuli, such as starvation or immune signals, trigger autophagy through the class III PI3K complex with Beclin 1 or the class I PI3K Atg1 complex that activate downstream autophagy-related genes (Atgs). Autophagy is initiated by the formation of a double-membrane autophagosome that surrounds cargoes. A number of autophagy proteins (Atgs) are involved in the formation of two ubiquitin-like conjugation systems involving Atg12 and the microtubule-associated protein 1 light chain 3 (LC3), that mediate autophagosome formation. Cleavage of LC3 to LC3-I by Atg4 is followed by the lipidation of LC3-I by Atg7 and Atg3 that form the transient linkage between LC3-I and phosphatidylethanolamine to make LC3-II. This conversion of LC3-I to LC3-II during autophagosome formation allows for the incorporation of LC3 into the autophagosomal membrane. Maturation of the autophagosome into an autolysosome with a single membrane, involves fusion with a lysosome such that the sequestered contents can now be degraded (Adapted from Lin et al., 2010).

56

microscopy and western blotting. Both the autophagy inhibition and autophagy silencing

experiments will be conducted in the presence or absence of proteasomal inhibitors, since

EBNA1 may be able to shuttle PML proteins down both degradation pathways. These

experiments will also be repeated in AGS cells, a gastric carcinoma cell line that is also

biologically relevant to EBV, to demonstrate this phenotype in multiple cell lines. Third, nuclei

vs microsome fractionation experiments will be performed to determine if there is a shift of

PML protein localization from the nucleus to the cytoplasm during EBNA1 overexpression (+/-

autophagy inhibitors). Paludan et al (2005) have previously demonstrated an accumulation of

EBNA1 in the cytosol, particularly in lysosomal vesicles (microsomes), upon inhibition of

acidification.

Since PML NBs are localized inside the nucleus, PML proteins would have to gain

access to the cytoplasm via nuclear export in order to be degraded through canonical autophagy,

or PML proteins would need to be pinched off from the nucleus via nucleophagy (autophagy of

the nucleus), followed by lysosomal fusion in the cytoplasm. Nucleophagy was first described in

yeast as a process wherein small pieces of the nucleus are pinched off into the yeast degradative

vacuole (Roberts et al., 2003). Nucleophagy has recently been observed in mammalian cells

(human and mouse macrophages, fibroblasts, and epithelial cells); in this instance, HSV-1

infection triggers a form of autophagy involving four-layer autophagosomes emerging from the

nuclear envelope (English et al., 2009; Radtke et al., 2013).

The p62/SQSTM1 or NBR1 adaptor proteins can function in the nucleus and could

therefore be recruiting polyubiquitinated PML protein to the cytoplasm. Notably, the

p62/SQSTM1 adaptor recognizes K63 linked polyubiquitin, which has been shown to be

involved in protein regulation by selectively targeting proteins to autophagosomes for

degradation (Matsumoto et al., 2011) (whereas K48 linked ubiquitin is usually involved in

targeting proteins for proteasomal-mediated turnover). The ability of EBNA1 to disrupt PML

NBs will be examined when p62 or NBR1 have been silenced with siRNAs in CNE2 cells,

followed by EBNA1 overexpression (+/- proteasomal inhibitors). The differences in K63 and

K48 linked polyubiquitin chains on PML proteins will also be determined with antibodies

specific to these linkages. This experiment will be done by overexpressing EBNA1 in the

presence of autophagy or proteasomal inhibitors, to block degradation, and western blotting for

the ubiquitin chains. Next, EBNA1-mediated PML degradation will be examined in the presence

of nuclear export inhibitors (Leptomycin B), as this may abrogate potential PML protein export

and subsequent degradation.

57

Finally, the localization of EBNA1 and PML proteins to autophagic structures will be

investigated. Following the co-transfection of EBNA1 and GFP-tagged LC3-II (a marker of

autophagic structures due to its incorporation into autophagosome membranes), cells will be

stained for EBNA1 and PML and visualized by IF microscopy. Cells will also be stained for

LAMP-1, as an alternative to LC3-II, for visualizing lysosomes, as EBNA1 has already been

reported to co-localize with LAMP-1 upon inhibition of lysosomal acidification (Paludan et al.,

2005). Visualizing EBNA1/PML proteins localizing to LC3-II stained autophagosomes will be

essential to confirm the role of autophagy in PML degradation; additionally, visualizing

EBNA1/PML budding off from the nucleus will be essential to confirm the involvement of

nucleophagy in this process.

EBNA1 requires binding to the host cellular protein USP7 to induce PML degradation.

Additional studies demonstrated that USP7 itself is a negative regulator of PML proteins and

that this negative regulation is not dependent on USP7’s deubiquitination activity (Sarkari et al.,

2011). The mechanism by which USP7 regulates PML proteins remains unclear; however, it is

likely USP7 is recruiting other negative regulators to PML NBs. I will, therefore, examine the

role of autophagy in USP7’s ability to regulate PML proteins with the same experiments

outlined above.

The normal regulation of PML proteins and PML NBs has been thought to be proteasome-

mediated, and since proteasome- and autophagy-mediated degradation of proteins are not

mutually exclusive, it will be important to determine if a population of PML proteins are turned

over by autophagy in the absence of EBNA1. This will be investigated by silencing key

autophagy components (siAtg12, sip62) or using autophagy inhibitors (3MA, Bafilomycin,

NH4Cl) on wild-type CNE2 cells to investigate whether there is an increase in PML NBs by IF

and western blotting. Importantly, PML proteins exist as six nuclear isoforms, which may have

differential regulation. To address this possibility, the autophagy inhibitors and siRNAs, as well

as proteasomal inhibitors, will be used in our cell lines that express individual PML isoforms to

determine if the different isoforms are regulated by different pathways.

Recently, I have started investigating the role of the autophagy degradation pathway in

EBNA1-mediated PML NB disruption. To this end, I have shown that treating CNE2E cells

with either a proteasomal inhibitor (MG132) or an autophagy inhibitor (3-methyladenine)

causes a subtle increase in PML NB number and protein levels. However, when both inhibitors

are used simultaneously, an additive affect is observed with respect to an increase in PML NB

58

B C

0  2  4  6  8  10  12  14  16  18  

Control   3MA   MG132   3MA  &  MG132  

PML  NB  #  

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Figure 4-3: An autophagy inhibitor (3MA) and a proteasomal inhibitor (MG132) additively block PML protein degradation in CNE2E cells. (A) CNE2E cells (stably expressing EBNA1) were treated with a control (DMSO), 3-methyladenine (3MA), MG132, or a combined drug treatment with both 3MA and MG132. Samples were fixed at 10 hours post drug treatment. Cells were then stained with primary antibodies against EBNA1 (red) and PML (green), and counter stained with DAPI (blue). Secondary antibodies used to stain EBNA1 and PML were GAR 555 and GAM 488, respectively. (B) The average number of PML NBs was determined by counting NBs in 50 cells of each condition for one experiment. (C) The amount of PML protein was examined by western blot analysis; actin was used as a loading control.

60

Figure 4-4: Inhibiting autophagy in CNE2 cells causes an increase in PML NBs. Cells were treated with 3-methyladenine (3MA) to inhibit autophagy and fixed at 24 hours post drug treatment. Cells were then stained with primary antibodies against PML, and counter stained with DAPI. The secondary antibody used to stain PML (green) was GAM 488.

61

number and protein levels (Figure 4-3). I have also seen an increase in PML NBs with 3-

methyladenine treatment in wild-type CNE2 cells in the absence of EBNA1/EBV (Figure 4-4).

4.7 Concluding remarks

PML NBs have an imperative role in a variety of tumor suppressive and antiviral

functions and govern appropriate levels of cell proliferation and survival in response to stimuli.

The turnover of PML proteins is an important determinant in controlling PML-associated

functions and, thus, elucidating mechanisms of PML regulation is important. A variety of

viruses have evolved mechanisms to induce the disruption of PML NBs in order to facilitate

viral infection; one such virus is Epstein-Barr virus, which utilizes the EBNA1 protein to

mediate the degradation of PML NBs in both latent and lytic infection. EBNA1 requires binding

to the host protein USP7 for this function, and USP7 itself negatively regulates PML.

Expanding our knowledge of how EBNA1 and USP7 are targeted to, associate with, and disrupt

PML NBs is essential to determine how these proteins regulate PML. I have provided useful

data to this end; specifically, I have confirmed USP7’s negative regulation of PML in USP7-null

cells, and I have shown that the EBNA1 393 phosphosite is crucial for EBNA1-mediated PML

disruption. I have also demonstrated that EBNA1 and USP7 directly associate with PML

proteins and exhibit a binding specificity for PML IV. Additionally, proteasome-mediated

turnover of PML proteins has been the accepted avenue of PML degradation. However, recently

I have found that EBNA1 is able to facilitate PML NB degradation in the presence of

proteasomal inhibitors. EBNA1 turnover in B cells has been shown to occur through autophagy;

however, it remains to be determined whether EBNA1 processing is through autophagy in

epithelial cells. Investigating the potential involvement of autophagy in both EBNA1-mediated

PML protein degradation and normal PML turnover could reveal a novel mechanism of PML

regulation. In summary, investigating the regulation of PML proteins by EBNA1 and USP7 has

proved to be a useful tool in understanding the complexities of PML biology.

62

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