Akap7 Degrades 2-5A Mediators of The

AKAP7 DEGRADES 2-5A MEDIATORS

OF THE INTERFERON ANTIVIRAL RESPONSE

ELONA GUSHO

Bachelor of Science in Biology

Fatih University, Istanbul, Turkey

June 2009

Submitted in partial fulfillment of the requirements for the degree

DOCTOR OF PHILOSOPHY IN REGULATORY BIOLOGY

CELLULAR AND MOLECULAR MEDICINE SPECIALIZATION

at the

CLEVELAND STATE UNIVERSITY

December 2015

We hereby approve this dissertation for Elona Gusho Candidate for the Doctor of Philosophy in Regulatory Biology degree for the Department of Biological, Geological and Environmental Sciences and the CLEVELAND STATE UNIVERSITY College of Graduate Studies

______Dissertation Chairperson, Robert H. Silverman, Ph.D.

______Department & Date

______Dissertation Committee Member, Sailen Barik, Ph.D.

______Department & Date

______Dissertation Committee Member, Donna M. Driscoll, Ph.D.

______Department & Date

______Dissertation Committee Member, Anton A. Komar, Ph.D.

______Department & Date

Student’s Date of Defense: September 17, 2015

DEDICATION

To my parents,

Vjollca and Vangjo Gusho for their unconditional love and support in every

step of my life. Thank you!

Per prinderit e mi,

Vjollca dhe Vangjo Gusho. FALEMINDERIT per dashurine, sakrificat

besimin, dhe mbeshtetjen tuaj ne cdo hap te jetes time!

ACKNOWLEDGMENT

Today as I am writing my PhD thesis; I look back at the path of this unique, beautiful and challenging journey and am very grateful to many people who have supported and motivated me to get here.

First I would like to express my infinite gratitude to my mentor, Dr. Robert

Silverman for being such a great educator and leader in the beautiful and challenging road of scientific research. Thank you for all you have taught me, your advices, your patience, and your trust in me. Thank you very much for giving me the opportunity to be a member of your laboratory. I hope that one day my contribution to science will be as significant as yours!

I would also like to extend my gratitude to my dissertation committee members: Dr. Anton Komar, Dr. Donna Driscoll, and Dr. Sailen Barik for their time and helpful feedback throughout my PhD education. Your questions, suggestions and comments have made a significant impact in my research project and my progress as a PhD candidate. Thank you!

I am very thankful to Dr. George Stark and Dr. Aimin Zhou for accepting to be part of my committee. It is my pleasure to have the opportunity to have their feedback.

My training as a PhD student was remarkably supported by present and past members of Silverman laboratory, an environment from which I have learned a lot. I would like to thank Dr. Jaydip Das Gupta and Dr. Arindam Chakrabarti for teaching me many techniques in the early days of my PhD training; Dr. Babal Jha for his help with the HPLC in this project and for the interesting scientific discussions in the lab; Christina Gaughan for being very friendly and helpful with many techniques; and for always taking great care of the lab; Dr. Shuvojit Banerjee for sharing his scientific ideas and technical expertise whenever I needed it; and Dr. Beihua Dong for always being so kind and knowledgeable.

A significant part of this training has also been our collaboration with Dr.

Susan Weiss and her laboratory members at the University of Pennsylvania. Our joined meetings have been a wonderful experience of sharing exiting data and scientific ideas. I am very thankful to Dr. Susan Weiss, Dr. Rong Zhang and Dr.

Joshua Thronbourgh for the collaboration in this project.

I would also like to thank the faculty members of the Regulatory Biology program at Cleveland State University, and in particular Dr. Girish Shukla for his generous advices and support. In addition, I would also like to thank all the members of the Cancer Biology department at the Lerner Research Institute,

Cleveland Clinic for being very helpful and friendly. I am also very thankful to the faculty of Biology Department in Fatih University, Istanbul; an environment and city from which I carry beautiful memories. I wish to acknowledge Preca College, in my home country Albania, for the high quality education they provide to the new generations.

During my studies I have met a lot of wonderful people and made lots of new friends. I would like to thank: Arishya Sharma, Bakytzhan Bakhautdin, Esen Goksoy Bakhautdin, Gaelle Muller, Gaurav Choudhary, Maria Barton, Payel

Chatterjee and Turkeyah Alswillah. In particular I would like to thank my best friend and soul sister Amina Abbadi, with whom we have shared our successes and failures, the best and the worst events of our lives in the past few years. My heartfelt thanks go also to: Aiola Ambo, Ardita Gusho, Denitsa Pirinova

Sokolova, Elona Murataj, Kriselda Collaku, Nina Shkurti, Uzma Jamil and

Vangjola Gjika. I wish you all best of luck in making your own dreams come true!

And finally, I dedicate this work to my family: to my parents, who have always emphasized to me the importance of education for a better future, even if this meant 10 years of being apart. To my dear brother Ondi and sister-in-law

Rezarta for the support and courage they have given me throughout the PhD training years. It is also because of you that I am here today! And to my dear nephew, Noel, who has brought so much joy in our hearts from the day he was born. I would also like to thank two very special people: my aunt Mirjana and uncle Thomas Bicolli for their love and care. Thank you for making me feel at home, even though thousands of miles away from home. My thanks are extended to my dear cousins: Kleida, Aisha, Rajli and Peter for being both family and friends to me; and to two lovely girls: Emma and Olivia.

I am blessed to have you all in my life. Thank you so much!

“Wherever the art of medicine is loved, there is also a love of humanity.”

Hippocrates

AKAP7 DEGRADES 2-5A MEDIATORS

OF THE INTERFERON ANTIVIRAL RESPONSE

ELONA GUSHO

ABSTRACT

Higher vertebrates have evolved innate immunity genes, many of which function in the interferon (IFN) induced antiviral response. Type I IFNs are produced in response to viral infections and induce expression of several hundred IFN stimulated genes, including genes for the 2’,5’-oligoadenylate (2-5A) synthetase (OAS). The OAS/RNase L pathway is one of the principal mediators of IFN antiviral response. RNase L cleaves viral and cellular ssRNAs inhibiting viral replication. Viral-encoded 2’,5’-phosphodiesterases (2’,5’-PDEs) allow these viruses to evade the OAS/RNase L by degrading 2-5A, activators of RNase L.

Moreover, mammalian 2’,5’-PDEs exist that are believed to limit RNase L activity.

Sustained RNase L activity after viral clearance may lead to cell death and hence tissue damage. Thus, we hypothesized that eukaryotic members of the 2- histidine (2H)-phosphoesterase family might have a similar function to related viral 2’,5’-PDEs (murine coronavirus ns2 and rotavirus VP3). This study identified a homologous mammalian 2’,5’-PDE, a member of the A-kinase anchoring protein (AKAP) family, that cleaves 2-5A. Recombinant, purified mouse AKAP7 degraded 2-5A with kinetics similar to viral PDEs, while a mutant of two conserved histidine residues was catalytically inactive. Similiarly, in intact cells vii

the expression of wild type but not mutant AKAP7 significantly reduced 2-5A levels after stimulation of OAS by transfection of synthetic dsRNA, polyI:polyC.

The PDE activity was confirmed by generation of cell lines in which AKAP7 was stably depleted. To determine if AKAP7 could substitute for a viral 2′,5′-PDE,

AKAP7 cDNA was inserted into an MHV genome with an inactivated mutant ns2 gene. The AKAP7 PDE domain restored the infectivity of the ns2 mutant MHV in bone marrow macrophages and in livers of infected mice. Interestingly, the PDE domain of truncated AKAP7 localized to the cytoplasm whereas full-length

AKAP7 was observed only in the nuclei. We hypothesized that there might be additional related cellular 2’,5’-PDEs that degrade 2-5A, also based on structure homology. However, the cellular 2H-phosphoesterases, CGI-18, Leng9 and

USB1 failed to degrade 2-5A, and apparently have different functions. We suggest that viral acquisition of AKAP7 PDE domain might have occurred during evolution, allowing diverse RNA viruses to antagonize the OAS/RNase L pathway.

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TABLE OF CONTENTS

ABSTRACT ...... vii

LIST OF FIGURES ...... xii

LIST OF ABBREVIATIONS ...... xiv

CHAPTERS

CHAPTER I: INTRODUCTION ...... 1

1.1 Interferons, an overview ...... 1

1.1.1 IFN classification ...... 2

1.1.2. IFN signaling ...... 3

1.1.3. The IFN-I antiviral response ...... 6

1.2 The OAS/RNase L pathway ...... 8

1.2.1 OAS sensors of foreign dsRNA ...... 11

1.2.2 Synthesis of 2’-5’-linked oligoadenylates by OAS ...... 13

1.2.3 RNase L, a ribonuclease with multiple biological functions ...... 15

1.2.3.1 The structure of RNase L ...... 16 1.2.3.2 RNase L, an endoribonuclease with many roles ...... 19

1.3 Inhibition of OAS/RNase L by cellular and viral proteins ...... 24

1.3.1 Cellular inhibitors of OAS/RNase L ...... 24

1.3.2 Viral inhibitors of OAS/RNase L ...... 27

1.3.2.1 Viral proteins that bind RNase L ...... 28 1.3.2.2 Viral encoded dsRNA binding proteins ...... 28

1.4 The 2H phosphoesterase superfamily of proteins that degrade 2-5A ...... 30

1.4.1 The 2H-Phosphoesterase superfamily ...... 30

1.4.2 The Eukaryotic-viral Lig-T family of proteins ...... 31

1.4.2.1 The viral Lig-T 2H-phospoesterases that degrade 2-5A: antagonism of OAS/RNase L by viral phosphodiesterases ...... 31 1.4.2.2 The eukaryotic Lig-T 2H-Phosphoesterases ...... 37

1.5 The A-Kinase Anchoring Protein 7 ...... 39

1.5.1 AKAPs: a superfamily of proteins with multiple functions ...... 39

1.5.2 AKAP7 as a PKA anchoring protein……………………… ...... 40

1.5.3 AKAP7 is a 2H-Phosphoesterase……………………… ...... 44

CHAPTER II: MURINE AKAP7 HAS A 2’,5’ PHOSPHODIESTERASE DOMAIN THAT CAN COMPLEMENT AN INACTIVE MURINE CORONAVIRUS ns2 GENE * ...... 47 2.1 Introduction ...... 48

2.2 Materials and Methods ...... 51

2.3 Results ...... 60

2.4 Discussion ...... 86

CHAPTER III: FURTHER CHARACTERIZATION OF AKAP7 AND OTHER CELLULAR CANDIDATE 2’,5’-PDEs ...... 93

3.1 Introduction ...... 93

3.2 Materials and Methods ...... 95

3.3 Results ...... 99

3.4 Discussion ...... 110

CHAPTER IV: FUTURE PROSPECTIVE ...... 114

CHAPTER V: CONCLUSIONS ...... 118

5.1 Overall summary and conclusions ...... 118

Bibliography ...... 120

LIST OF FIGURES

FIGURE PAGE

Figure 1.1 The IFN signaling to ISGs ...... 5

Figure 1.2 The OAS/RNase L pathway ...... 10

Figure 1.3 The structure of 2-5A ...... 14

Figure 1.4 The structure of RNase L bound to 2-5A ...... 18

Figure 1.5 Viral antagonists of OAS/RNase L ...... 36

Figure 1.6 AKAP7 isoforms...... 43

Figure 1.7 Mammalian AKAP7 is structurally homologous to viral ns2 of MHV and VP3 of rotavirus ...... 45

Figure 2.1 AKAP7 rapidly degrades 2-5A with similar kinetics as MHV strain A59 ns2 and human rotavirus strain WA VP3-CTD ...... 62

Figure 2.2 AKAP7 degrades 2-5A in intact cells ...... 65

Figure 2.3 Construction of chimeric viruses expressing AKAP7 protein ...... 67

Figure 2.4 Cellular localization of the AKAP7s in different cell lines ...... 70

Figure 2.5 Expression of the N-terminal truncation or the CD of AKAP7 restores the replication of ns2 mutant in vitro ...... 74

Figure 2.6 Expression of the N-terminal truncation or the CD of AKAP7 restores the replication of ns2 mutant in vivo ...... 77

Figure 2.7 Conservation of 2’,5’-phosphodiesterases among distantly

related taxa ...... 78

xii

Figure 2.8 Protein multiple sequence alignment (MSA) of mAKAP7γ and eukaryotic homologs ...... 80

Figure 2.9 Phylogenetic relationship between 2’,5’-phosphodiesterases of distantly related taxa ...... 85

Figure 3.1 Candidate cellular 2’,5’-PDEs do not degrade 2-5A...... 101

Figure 3.2 USB1 PDE does not degrade 2-5A ...... 103

Figure 3.3 AKAP7 and AKAP7 CD have minimal effect on EMCV

viral titers ...... 106

Figure 3.4 AKAP7, ns2 and VP3-CTD have minimal effect on EMCV

viral titers ...... 107

Figure 3.5 Depletion of AKAP7 and PDE12 increases 2-5 accumulation and RNase L activity ...... 109

xiii

LIST OF ABBREVIATIONS

2-5A 2’5’- linked oligoadenylate

ADP Adenosine diphosphate

AKAP A-kinase anchoring protein

AMP Adenosine monophosphate

ANK Ankyrin repeat domain

ASC Activating signal cointegrator

ATP Adenosine triphosphate

CD Central domain

CH25H Cholesterol-25-hydroxylase

CPD Cyclic phosphodiesterase

CTD C-terminal domain dsRNA Double-stranded RNA eIF Eukaryotic translation intiation factor

EMCV Encephalomyocarditis virus

GAF Gamma interferon activated factor

HIV Human immunodeficiency virus

HPC Hereditary prostate cancer

IAV Influenza A virus

IFITIM Interferon inducible transmembrane

IFN Interferon

IFNAR Interferon α receptor

xiv

IFNGR Interferon-γ receptor

IFN-I Type I interferon

IFN-II Type II interferon

IFN-III Type III interferon

IRF Interferon regulatory factor

ISG Interferon stimulated gene

ISGF3 Interferon stimulated gene factor 3

JAK Janus kinase

JNK c-Jun N-terminal kinase kd Knockdown

KEN Kinase-extension-nuclease

Ko Knockout

MDA Melanoma differentiation-associated gene

MEF Mouse embryonic fibroblast

MHV Mouse hepatitis virus

Mx Myxovirus resistance

NK Natural killer cell

NLS Nuclear localization signal ns Non structural

OAS Oligoadenylate-synthetase pIC Polyinosinic:polycytidylic acid

PAMPS Pathogen-associated molecular patterns

PDE Phosphodiesterase

PK Protein kinase

PKA Protein kinase A

PKR Protein kinase R

PRR Pattern recognition receptors

RIG Retinoic acid- induced gene

RLI Rnase L inhibitor

RNASE Ribonuclease

STAT Signal transducers and activators of transcription

TRIM Tripartite motif

TYK Tyrosine kinase

VP Viral protein

ZAP Zinc-finger antiviral protein

xvi

CHAPTER I

INTRODUCTION

1.1 Interferons, an overview

Discovery of the interferon (IFN) family of cytokines represented one of the most fundamental advances in molecular biology (Isaacs and Lindenmann,

1957). Today, 58 years after the identification of IFN as an antiviral agent secreted from cells exposed to heat-inactivated influenza virus we now know that

IFNs are a diverse family of cytokines, some of which have demonstrated clinical utility against certain viral infections and malignancies, multiple sclerosis and chronic granulomatous disease.

The innate immune response mediated by IFNs is the first line of defense against pathogen infection. This response is driven in the host cell by the pattern recognition receptors (PRRs) through recognition of the foreign ligand known as pathogen-associated molecular patterns (PAMPs). PRRs consist of several

families of receptors which are secreted, membrane-bound or localized in the cytoplasm. Upon exposure to PAMPs, PRRs initiate intracellular signaling cascades resulting in expression of chemokines and cytokines genes, including

IFN genes that together orchestrate a cellular protective state against pathogens.

1.1.1 IFN classification

IFNs are classified based on their sequence homology and the receptors that they bind and signal through. Type I IFN (IFN-I) is the largest family consisting in humans of 17 genes: 13 IFN-α genes and a single gene each for

IFN-β, IFN-ω, IFN-ε, IFN-δ, and IFN-τ. All IFN-I members signal through the IFN-

α receptor (IFNAR), a heterodimer of two polypeptides, IFNAR1 and IFNAR2

(Figure 1.1). A broad range of cell types express IFNAR and hence respond to

IFN-I signaling. IFN-I will be more extensively described in the next section.

Type II IFN (IFN-II or IFN-γ) consists of a sole cytokine encoded by a single gene (Naylor et al., 1983), and is mostly produced by natural killer (NK) and T cells. IFN-γ is a significant agent of innate and adaptive immunity involved in mechanisms against both pathogens and tumors. The IFN-γ receptor, composed of IFNGR1 and IFNGR2 (Figure 1.1), is expressed in a wide range of cell types. Mutations in IFN-γ and its receptor genes have been associated with higher susceptibility to viruses and intracellular bacteria (Schroder et al., 2004) while IFN-γ production controlled tumor development in mice treated with carcinogens (Kaplan et al., 1998, Shankaran et al., 2001).

Three genes belong to Type III IFNs (IFN-III): IFN-λ1 (IL-29), IFN-λ2(IL-

28A) IFN-λ3(IL-28B) (Kotenko et al., 2003, Sheppard et al., 2003). In a recent study a fourth member IFN-λ4 was also described to originate from a deletion frameshift in IFN-λ3 (Prokunina-Olsson et al., 2013) The receptor for IFN-λ signaling is a heterodimer complex consisting of IFNLR1 and IL-10R2 (Figure

1.1). IFNLR1 is expressed mostly in epithelial cells (Sommereyns et al.,

2008)restricting IFN-λ signaling to these cell types. IFN-λ signaling is associated with a host antiviral response. In addition, IFN-λ has also been studied in therapeutic applications. In a pre-clinical study, hepatitis C patient treatment with

PEGylated IFN-λ1, showed an antiviral effect as well as minimal side effects

(Miller et al., 2009).

1.1.2 IFN signaling

The IFN response starts upon IFN binding to its high-affinity receptors triggering a cascade of downstream signaling events leading to activation of IFN stimulated genes (ISGs). Between the IFNs, their receptors and the ISGs stands the Janus kinase (JAK) and signal transducers and activators of transcription -

(STAT) pathway. The JAK-STAT pathway consists of the tyrosine protein kinases: JAK1, JAK2 and TYK2 (tyrosine kinase 2) that are pre-associated with the cytoplasmic portions of IFN receptors and STATs which are phosphorylated by the JAKs and migrate to the nucleus as part of transcription factor complexes to turn on transcription of the ISGs. Upon IFN binding to the specific IFN receptor, the JAK kinases phosphorylate the receptor leading to recruitment and 3

phosphorylation of the STATs. The phosphorylated STAT heterodimers

(STAT1/STAT2 for IFN-I and IFN-III) bind to the IFN regulatory factor (IRF) IRF9 to form IFN-stimulated gene factor 3 (ISGF3). While STAT1 homodimers in IFN-II signaling form gamma IFN activated factor (GAF). Once formed, these complexes translocate to the nucleus and activate transcription of ISGs (Figure

1.1). ISGs comprise a family of hundreds of genes that have a wide range of roles including targeting pathogen infections at different stages of its replication or inducing cell death (Der et al., 1998).

Figure 1.1 The IFN signaling to ISGs. Type I-III IFNs bind their specific receptors and signal through JAK/STAT to activate expression of several hundred ISGs. From: (Sadler and Williams, 2008)

1.1.3 The IFN-I antiviral response

IFN-I signaling plays a critical role in activating a protective state of the host cell during viral infections. The genes encoding type I IFNs are located on chromosome 9p in humans and chromosome 4 in mice (Pestka et al., 2004).

Expression of these genes is turned on upon recognition of non-self agents through their PAMPs. Most viral PAMPs consist of nucleic acids such as: double stranded RNA (dsRNA), uncapped 5’-triphosphorylated RNA, or cytoplasmic

DNA. All the members of the IFN-I family signal through IFNAR, as a consequence mice deficient in this receptor were unresponsive to IFN-I and dramatically more susceptible to viral infections (Muller et al., 1994). ISGs activated by type I IFN exceed 300 genes, some of which are involved in antiviral signaling cascades (Der et al., 1998). Due to the ability of viruses to antagonize the host antiviral mechanisms, the activated gene products initiate pathways that target the virus at different stages of its replication. Some of these proteins are also known as “antiviral effectors” (Sadler and Williams, 2008) and they restrict viral entry, translation, replication, or assembly. Some examples of IFN induced antiviral proteins that target viral entry include: the IFN-inducible transmembrane

(IFITIM), tripartite motif (TRIM) and cholesterol-25-hydroxylase (CH25H).

Myxovirus resistance (Mx) proteins are involved in inhibition of intracellular trafficking of subviral particles. These consist of two members: Mx1 and Mx 2 (in mice) or Mx A and Mx B (in humans). The mouse Mx locus was identified as a gene that increased resistance to influenza A virus in A2G mouse

strain (Lindenmann, 1962). Mx1 protein was shown to be responsible for this effect (Arnheiter and Meier, 1990). Mx1 forms ring-like structures that surround viral nucleocapsid and lead to degradation or impairs its intracellular trafficking

(Haller and Kochs, 2011). Mx 2 protein was recently reported as an inhibitor of

HIV infection that prevents viral DNA integration (Goujon et al., 2013, Kane et al.,

2013, Liu et al., 2013).

Protein kinase R (PKR) is an extensively studied ISG encoding a serine/threonine protein kinase activated by dsRNA. It was first described in cells treated with IFN and infected with vaccinia virus (Metz and Esteban, 1972) or treated with IFN and dsRNA (Kerr and Brown, 1978). PKR inhibits viral replication by phosphorylating one of the key components of translation machinery: the eukaryotic translation initiation factor 2α (eIF2-α). eIF2-α phosphorylation inhibits translation of both viral and cellular mRNA hence inhibiting viral propagation. A recent study revealed a novel function of PKR being involved in maintaining integrity of IFN-β mRNA (Schulz et al., 2010).

Therefore, PKR restricts protein synthesis while also stabilizing the type I IFN response.

Other IFN-inducible antiviral effectors include IFN-stimulated gene 15

(ISG15), IFN-induced protein with tetratricopeptide repeats (IFIT), zinc-finger antiviral protein (ZAP) and the oligoadenylate synthetase (OAS)/RNase L

(OAS/RNaseL) pathway. ISG15 is a very potent antiviral effector. It is highly

induced by IFN-I during viral infections and it was first isolated from cells treated with IFN-β (Blomstrom et al., 1986). Mice deficient in ISG15 were susceptible to infections with Sindbis, herpes and influenza viruses, (!!! INVALID CITATION !!!,

Lenschow et al., 2007). ISG15 is mainly involved in a process known as

ISGylation, (similar to ubiquitination) where it binds to the target proteins and modifies them leading to a change in function (Zhang and Zhang, 2011). The substrate proteins can have either a viral or host origin.

Overall, the host innate immunity programs pathways specifically target viral infections at different stages. My studies focus on the OAS/RNase L pathway, an IFN induced pathway that senses dsRNA and inhibits viral replication by degrading single-stranded (ss) RNA.

1.2 The OAS/RNase L pathway

OAS/RNase L is an IFN-I induced antiviral pathway. OAS genes are ISGs but are constitutively expressed in cells. The levels of OAS proteins increase significantly during a viral infection as a result of IFN signaling. The IFN may come from neighboring cells as an alert for pathogen presence (paracrine signaling) or it may come from the infected cells themselves (autocrine signaling). OAS proteins are PRRs that recognize viral dsRNA in the cytoplasm

of virus infected cells. In response to dsRNA, most OAS proteins synthesize a unique series of oligoadenylate compounds characterized by an unusual 2’-5’ linked phosphodiester bond from ATP; the compounds are known as 2-5A

(Figure 1.3). Inactive, monomeric RNase L dimerizes upon binding to 2-5A, to its active ribonuclease form (Dong and Silverman, 1995). The active protein cleaves ssRNA preferentially after (3’) of UU or UA (Wreschner et al., 1981). The ssRNA substrates have either viral or cellular origins. The end result of this RNA cleavage is inhibition of viral replication and spread. 2-5A turnover is controlled by phosphodiesterases and possibly phosphatases. Each of the components of this pathway will be described in more detail in this section. (The pathway is schematically described in Figure 1.2)

Figure 1.2 The OAS/RNase L pathway

From: (Silverman and Weiss, 2014a)

1.2.1 OAS sensors of foreign dsRNA.

Many types of viruses produce dsRNA in the cytoplasm. This unusual form of nucleic acid is recognized as non-self by host PRRs such as OAS. The most studied functions of OAS proteins are their ability to sense dsRNA and synthesize 2-5A.

Human OAS1-3 belongs to the template-independent nucleotidyl transferase superfamily characterized by a conserved motif in the active site, containing three conserved asparatic acid residues (Holm and Sander, 1995,

Steitz, 1998). There are four human OAS genes: OAS1, OAS2, OAS3 and OAS- like (OASL) (Justesen et al., 2000). The OAS1-3 genes are located in chromosome 12q24.1 and OASL in 12q24.2 and they encode for six isoforms:

OAS1 (p40 and p46), OAS2 (p69 and p71), OAS3 (p100) and OAS L (p59)

(Chebath et al., 1987a). While murine OASs are encoded by eight Oas1 genes, two Oasl and single genes of Oas 2 and Oas 3 (Eskildsen et al., 2002). The

OAS1-3 family members share structure homology as well as their ability to synthesize 2’-5’ linked phosphodiester bonds. Characterization of the OAS unit in porcine OAS1 revealed two structural domains: the N-terminal domain consisting of 5 antiparallel β-strands and a C-terminal α-helix (Hartmann et al., 2003).

OAS1-3 consist of 1-3 OAS units, respectively, as their names suggest. These proteins contain a positively charged binding grove where dsRNA binds

(Hartmann et al., 2003). OASL differs from the other members in its ubiquitin-like

C-terminus (Hartmann et al., 1998) and it was initially described as an OAS that lacks catalytical activity (Rebouillat et al., 1998). A recent study of OASL

structure showed a binding site for dsRNA that is involved in RIG-I signaling

(Ibsen et al., 2015)

OAS1 has a preference for longer dsRNA (Desai and Sen, 1997) and it is activated by more than 20,000 fold upon dsRNA binding which leads to conformational change at the N-terminal and brings the 3 active site residues

(D75, D77 and D148) closer together (Donovan et al., 2013). OAS1 has an antiviral activity against viruses such as EMCV, Dengue virus and West Nile virus

(Chebath et al., 1987b). Polymorphisms in the OAS1 gene are related to higher virus infectivity for the splice variant p46 (Li et al., 2009). On the other hand, several viruses have adapted inhibition mechanisms against OAS1, some of which include: human cytomegalovirus and adenovirus VA (I) (Tan et al.).Similarly, OAS3 is an IFN induced antiviral protein that synthesizes 2-5A that activates RNase L (Ibsen et al.). All three OAS3 domains can bind dsRNA, but only the C-terminal domain possesses catalytical activity (Donovan et al.). OAS2 is also enzymatically active, synthesizing 2-5A in response to dsRNA stimulation.

Besides ATP, OAS use a variety of acceptor substrates including NAD, ADP- ribose, A5’p45’A and tRNA to synthesize 2’-5’ phosphodiester bond by the addition of 2’-AMP residues (Ferbus et al., 1981, Naylor et al., 1983).

1.2.2 Synthesis of 2’-5’ linked oligoadenylates by OAS

The OAS1-3 proteins have a positively charged dsRNA binding grove, which upon binding to dsRNA undergoes structural changes leading to shifts of the C- and N-termini and causing assembly of the active site (Hartmann et al.,

2003). Two ATP molecules bind to active OAS to start a 2-5A chain. The mechanism used to form the 2’-5’ phosphodiester bond has been reported to be similar to that used by polyadenosine polymerase forming 3’-5’ bond, the phosphoryl transfer reaction utilizes two magnesium ions (Steitz, 1998). The donor ATP is added to the 2’ AMP of the acceptor molecule releasing pyrophosphate. The product, referred to as 2-5A, is a 5’ triphosphorylated oligonucleotide with general formula: px5’A(2’p5’A)n. where x=1-3 and n>2 (Kerr and Brown, 1978).

2-5A was discovered as a low molecular weight inhibitor of protein synthesis in extracts of cells treated with IFN and incubated with dsRNA (Kerr and Brown, 1978). The most common species identified in IFN treated cells are the trimeric and tetrameric forms (Knight et al., 1980). And the trimeric form: p3A3 is the principal activator of RNase L (Figure 1.3).

2-5A is a critical second messenger of the pathway; it controls the enzymatic activity of RNase L, an antiviral endoribonuclease.

Figure 1.3 The structure of 2-5A

From: (Kerr and Brown, 1978)

1.2.3 RNase L, a ribonuclease with multiple biological functions

RNase L is an endoribonuclease characterized as a 2-5A dependent

RNase that was cloned by screening a cDNA expression library with radiolabelled 2-5A (Zhou et al., 1993). It is endogenously expressed in all mammalian tissues, typically in its monomeric and inactive form (Zhou et al.,

2005). Evolutionary studies have showed that it is highly conserved in different species from reptiles to mammals (Schroder et al., 2004). RNase L levels are significantly induced by IFN treatment in mice tissues but there is no such evidence in humans (Krause et al., 1985). In addition, RNase L levels are also regulated at the post-transcriptional level by eight AU-rich elements located in the

3’-UTR region, controlling its mRNA stability (Li et al., 2007). RNase L overexpression had a down-regulating effect on huR, an RNA binding protein, and in mouse fibroblasts lacking RNase L the huR mRNA stability and cell growth increased (Al-Ahmadi et al., 2009).

2-5A is a highly specific and sensitive (subnanomolar) activator of RNase

L. This binding leads to RNase L dimerization (Dong and Silverman, 1995) into its active ribonuclease form to degrade ssRNA preferably after UU or UA generating 5’-hydroxyl (5’-OH) and 2’,3’-cyclic monophosphate (2’,3’

Smith et al., 1981); (Wreschner et al., 1981, Cooper et al., 2014). The ssRNA can be viral RNA or cellular RNA such as ribosomal RNA (rRNA) or mRNA

(reviewed (Floyd-Smith et al., 1981, Silverman et al., 1983, Kaplan et al., 1998).

Indeed, the rRNA cleavage pattern is one of the hallmarks of RNase L activity in virus-infected cells (Silverman et al., 1983). The cleavage products can be self or non-self RNA which in turn have the ability to amplify the antiviral IFN response

(Malathi et al., 2010, Malathi et al., 2007). Due to its ability to degrade both viral and cellular RNA to inhibit viral replication, RNase L is involved in diverse processes.

1.2.3.1 The structure of RNase L

RNase L is composed of 3 major domains, from N-to-C termini: a regulatory ankyrin repeat domain (ANK), a pseudo protein kinase (PK) domain and a ribonuclease (RNASE) domain. The PK and RNASE domains together are also referred to as kinase-extension-nuclease (KEN) domain which is homologous to IRE1, a kinase and endonuclease involved in mRNA splicing in the unfolded protein response (Sidrauski and Walter, 1997, Korennykh et al.,

2009). The ANK consists of 9 ankyrin repeats, ankryn 2, 4 and 9 are involved in

2-5A binding (Han et al., 2014, Tanaka et al., 2004).

Two recent publications revealed a detailed crystal structure of porcine and human RNase L respectively, of the dimers bound to 2-5A and ADP/ATP

(Han et al., 2014, Huang, 2014). The ANK and PK domains are responsible for binding to 2-5A (Figure 1.4). In both structures, the PK domain is characterized by a larger C-terminal lobe compared to the N-terminal one. In addition, the C-

lobe lacks the activation segment of a typical kinase instead containing a four- residue shunt that causes an unusual placement of helix αG hence compromising the substrate recognition site of a kinase. The PK domain of

RNase L is therefore not capable of phosphorylation and is classified as a pseudokinase. The RNase domain consists of nine α-helixes and is similar to

IRE1 with a difference in its helix α1’. The dimer formation of an active RNase L requires interaction of ANK repeats 1 to 4 from one protomer and ANK 9 from the other, and a junction of the PK C-lobe with RNase domain (Figure 1.4A). A back- to-back dimer in the presence of ADP/ATP leads to assembly of the active porcine RNase site (Huang, 2014).

Similar dimer formation was also reported for human RNase L (Han et al.,

2014). This work identified a single residue, H672 in the KEN site as important for the RNase activity, consistent with a prior report on H672 (Dong et al., 2001).

Figure 1.4 The structure of RNase L bound to 2-5A

From: (Huang, 2014)

A. Surface view of 2-5A binding to two ANK and one PK domain

B. The groove formed between two RNase L protomers, the 2-5A binding

site

1.2.3.2 RNase L, an endoribonuclease with many roles

Viral and cellular RNA degradation. Different types of viruses produce ssRNA during their replication, which may be the viral genome in case of ssRNA viruses, in addition to mRNAs. RNase L is involved in the direct cleavage of these viral ssRNAs. An example of genomic cleavage is the picornavirus encephalomyocarditis virus (EMCV). EMCV RNA levels were significantly reduced in presence of RNase L, independent of IFN treatment (Li et al., 1998).

Viral ssRNA linked to double stranded regions are replication intermediates which are also targeted by RNase L (Nilsen and Baglioni, 1979). In theory, the advantage of such targeted viral cleavage would be that it does not interfere with normal cell function. More focused studies on specific viruses, such as influenza

A virus (IAV) have identified specific sites of RNase L cleavage (Shankaran et al., 2001). IAV with a deleted ns1 gene encoding a protein that sequesters dsRNA produced degradation of viral mRNAs for PB1, PB2 and PA which make up the viral polymerase and contribute to viral replication (Shankaran et al.,

2001).

Damage to host replication machinery may be key to inhibition of viral replication and spread. Active RNase L degrades both 28S and 18S rRNA in ribosomes in intact cells (Wreschner et al., 1981, Silverman et al., 1983). A variety of host mRNAs have also been identified as cleavage targets of RNase L in microarray studies (Domingo-Gil et al., 2010, Malathi et al., 2005, Kaplan et al., 1998). In addition to rRNA cleavage, RNase L also targets ribosomal protein

mRNAs through a RNase L-ribonucleoprotein complex formation to reduce protein synthesis in the infected cell (Andersen et al., 2009). A diverse number of genes targeted by RNase L were also identified in microarray studies of vaccinia virus infected HeLa cells. The down-regulated mRNAs are involved in processes such as cell growth (RUNX2 and ESR2) or mitochondrial homeostasis (COX5A and MIPEP; Domingo-Gil et al., 2010). Interestingly, some other mRNAs were up-regulated after activation of OAS/RNase L. These mRNAs are mostly related to cell growth arrest (GADD45B and KCTD11), apoptosis (CUL2, TNFAIP8L2 and PDCD6) or IFN action (IFN stimulated gene IFI6) (Domingo-Gil et al., 2010).

Role of RNase L in IFN induction and the innate immunity response.

As described above, the OAS-RNase L is a IFN-I induced pathway that blocks viral replication. However this system also has an effect on inducing IFN production hence amplifying the host antiviral response. The self or non-self

RNA cleavage products of active RNase L are characterized by 2’,3’-cyclic phosphate and 5’-OH groups (Floyd-Smith et al., 1981, Wreschner et al., 1981,

Han et al., 2004, Cooper et al., 2014) (Wreschner et al., 1981). These small

RNAs act as PAMPs and are recognized by PRRs, the retinoic acid–induced gene (RIG-I) and melanoma differentiation-associated gene (MDA5). As a result, small self-RNA produced after RNase L activation caused increase in IFN-β production. Also, in vivo data in mice lacking RNase L showed less IFN-β production when infected with EMCV virus (Malathi et al., 2007). Similarly, hepatitis C virus (HCV) RNA degraded by RNase L activated RIG-I (Malathi et

al., 2010). The effect of OAS/RNase L on IFN production may also be cell-type specific. Mouse embryonic fibroblasts (MEF) express lower levels of OAS and

RNase L and produce more IFN-β when treated with synthetic dsRNA, polyI:polyC (pIC) or infected with EMCV. In contrast. macrophages have high levels of RNase L and therefore pIC treatment inhibits protein synthesis and causes apoptosis. Consequently RNase L-/- macrophages produced more IFN-β when treated with pIC or when they were infected with virus compared to similarly treated wild type macrophages (Banerjee et al., 2014).

OAS/RNase L is also involved with other components of innate immunity such as the inflamasome. The RNase L cleavage products of RNA produced by virus infection activated the NLRP3 inflammasome and induced proinflammatory cytokine IL-1β (Chakrabarti et al., 2015).

Effects of RNase L on cell survival and homeostasis. Elimination of the infected cell is an effective way of restricting the spread of viral infection.

Sustained RNase L mediated RNA cleavage inhibits protein synthesis during viral infections and causes apoptosis (Castelli et al., 1997, Castelli et al., 1998,

Zhou et al., 1997, Diaz-Guerra et al., 1997). The process involves release of cytochrome c from mitochondria to cytoplasm and caspase-3 activation (Rusch et al., 2000). Other factors involved in RNase L induced cell death include degradation of mitochondrial mRNA (Le Roy et al., 2007) and activation of c-Jun

N-terminal kinase (JNK) (inhibition of JNK suppressed RNase L induced apoptosis) (Li et al., 2004).

Autophagy is a mechanism by which the cell recycles organelles and proteins to maintain homeostasis during cellular stress such as during viral infection (Dreux and Chisari, 2010). RNase L has a direct effect on virus induced autophagy. Autophagy was induced in cells treated with 2-5A and it was impaired in cells lacking RNase L and infected with EMCV or VSV. The antiviral effect of

RNase L against EMCV was enhanced when autophagy was inhibited, possibly due to activation of cytokines during early stages of infection (Chakrabarti et al.,

2012). The mechanism of autophagy induction by RNase L involves activation of

PKR and JNK (Siddiqui and Malathi, 2012). Furthermore, several other roles of

RNase L have been identified due to its ability to degrade cellular RNA. RNase L is implicated in inducing senescence as a result of ribosomal protein mRNA degradation (Andersen et al., 2007, Andersen et al., 2009).

RNase L in cancer. The ability of RNase L to control cell growth and induce apoptosis supports the possibility of RNase L being a tumor suppressor, as proposed by many studies. Prostate cancer is the second most common cancer in men and clusters within families is one of the risk factors for the disease. Genetic studies in patients with familial prostate cancer have mapped

RNase L gene to Hereditary Prostate Cancer (HPC1) locus at chromosome 1q25

(Carpten et al., 2002, Rokman et al., 2002). A germline mutation was identified at

R462Q of the PK-domain which decreased the ribonuclease activity by 3 fold

(Xiang et al., 2003). Homozygosity (QQ) increased the risk of prostate cancer by

2 fold (Casey et al., 2002). Mutations in RNASEL have also been linked to risk

other types of cancer, including head and neck, uterine, cervix, breast (Madsen et al., 2008), pancreatic (Bartsch et al., 2005) and hereditary non-polyposis colorectal cancer (Kruger et al., 2005).

Patients with inflammatory bowel disease have a higher risk of colitis- associated-cancer. Studies in mice showed that RNase L has a protective function against in vivo induced colitis and contributed to faster recovery by initiating an innate immune response through IFN-β production (Long et al.,

2013). Another novel function of RNase L is related to retrotransposons, mobile genetic elements have been associated with cancer (Wilkins and Gale, 2010).

The expression of wild type RNase L in human ovarian carcinoma cell lines restricted the mobility of an engineered LINE-1 retrotransposon (Zhang et al.,

2014).

Antibacterial defense. The protective role of RNase L in the host cell extends also to bacterial infections. Mice deficient in RNase L had higher bacterial levels and mortality rates compared to wild type mice when infected with bacteria such as Bacillus anthracis and Escherichia coli. In addition, expression of RNase L in the infected mice induced higher levels of proinflammatory cytokines and regulated levels of cathepsin-E, an endolysosomal protease involved in degrading lysosome-associated membrane proteins required for elimination of phagocytosed bacteria (Li et al., 2008).

Overall, that study showed a direct effect of RNase L in innate immunity against bacteria. The antibacterial function also correlates with inhibition of RNase L

activity by enteropathogenic Escherichia coli (EEC). EEC inhibited RNase L and

IFN-β production through its effectors in a type III-secretion system manner

(Long et al., 2014).

The host has developed several pathways to target and control virus replication. As reviewed above the OAS/RNase L is one of the mechanisms that inhibits infection at different stages and eliminates the infected cell. Several viruses have evolved mechanisms to counteract this host pathway and achieve successful replication, while the host also controls sustained enzymatic activation of RNase L possibly to prevent tissue damage.

1.3 Inhibition of OAS/RNase L by cellular and viral proteins

1.3.1 Cellular inhibitors of OAS/RNase L

OAS/RNase L is a regulated mechanism for eliminating or processing ssRNA, typically in virus-infected cells. Its induction comes as a result of IFN-I production (inducing OAS proteins) and viral dsRNA sensing by OAS proteins. At the same time there are also several proteins that control the activity of the pathway. In this section, I will describe the regulation and control of OAS/RNase

L by proteins of cellular and viral origin.

One of the earliest studied inhibitors of OAS/RNase L is: (as also its name suggests): is RNase L- Inhibitor (RLI or ABCE1). This inhibition was suggested to involve heterodimer formation to monomeric RNase L, reducing 2-5A binding

(Bisbal et al., 1995). It belongs to the ATP binding cassette proteins and is also named as the ATP binding cassette E1 (ABCE1). ABCE1 gene is localized in chromosome 4q31 (Diriong et al., 1996) and is not an IFN inducible gene (Bisbal et al., 1995). The protein is induced by synthetic dsRNA or viral infections such as EMCV or HIV (Martinand et al., 1999, Martinand et al., 1998b, Martinand et al., 1998a). ABCE1 is also involved in HIV-1 capsid assembly in cells, where it is a component of intermediates that assist in Gag assembly [Reviewed (Lingappa et al., 2014)]. Other functions of ABCE1 related to ATP binding involve ribosomal recycling and translation regulation (Pisarev et al., 2010). In addition, ABCE1 is also proposed to be implicated in malignancy. Gene expression profiling showed higher levels of ABCE1 in melanoma cells compared to normal cells (Heimerl et al., 2007). Also, studies in which ABCE1 levels have been depleted report an anti-proliferative and pro-apoptotic effect in cancer cells such as: small cell lung cancer, esophageal carcinoma cells and (Huang et al., 2014). Still, there is no clear scientific evidence that ABCE1 is directly related to the general antiviral or pro-apoptotic role of RNase L. Interestingly, ABCE1 is the only known cell encoded inhibitor of the pathway suggested to be involved in direct RNase L binding. Other inhibitors of the OAS/RNase L have been identified that inhibit the pathway at the 2-5A level.

Degradation of 2-5A is a mechanism that some viruses and the host cell use to limit activation of RNase L, and is the subject of my thesis. Considering the structure of 2-5A, degrading it to a form that cannot activate RNase L could occur through the action of either phosphodiesterases (PDEs) or phosphatases.

The 5’ nonphosphorylated 2-5As (referred to as core 2-5A) have little or no

RNase L activation ability. Early evidence of PDE activity against 2-5A was reported by Williams at al 1978 and Torrence et al 1983. One of the cellular candidates to degrade 2-5A is the 2’-phosphodiesterase (2’-PDE) also called phosphodiesterase 12 (PDE12), which belongs to the exonuclease- endonuclease phosphatase (EEP) domain superfamily. In vitro, PDE12 degraded the phosphodiester bond of trimeric 2-5A generating ATP and AMP (Kubota et al., 2004b). Further characterization of the protein showed that it also degrades other forms of 2-5A such as the tetrameric, pentameric and hexameric core releasing 5’ AMP residues (Poulsen et al., 2012a). Similar to other EEP members, the catalytic domain of PDE12 is in the C-terminus and contains an α- helix/β-strand structure (Wood et al., 2015). PDE12 knockdown by siRNA reduced vaccinia virus replication by approximately 50% (Kubota et al., 2004b).

Recently deletion of PDE12 gene by TALEN technology produced cells that required higher levels of EMCV (65 fold) to induce the same viral induced cytopathic effect as the control cells (Wood et al., 2015).

Moreover, PDE12 has been reported to localize in mitochondria and have an effect on mitochondrial mRNA turnover. It has a mitochondrial signal peptide at the N-terminal and both recombinant as well as endogenous protein localized

in this organelle. Besides from 2’,5’-PDE activity, PDE12 also has a 3’-5’ exoribonuclease activity (Poulsen et al., 2011b). Due to this enzymatic activity,

PDE12 controls mitochondrial gene expression by removing the poly A tail from mitochondrial mRNA (Rorbach et al., 2011). It still remains a question whether

PDE12 is directly involved in regulating antiviral and pro-apoptotic effects of

RNase L in vivo or if its function is mostly related to mitochondrial gene expression, where this protein resides. Further work needs to be done to answer these questions. Based on its nuclease activity, another cellular protein was also proposed as a possible 2’,5’-PDE: the ectonucleotide pyrophosphatase/phosphodiesterase 1 (ENPP1). ENPP1 resides in the plasma membrane with an extracellular active site (Poulsen et al., 2012a). In the same report, enzymatic studies showed that ENPP1 has the ability to degrade 2-5A.

1.3.2 Viral inhibitors of OAS/RNase L

Viruses have evolved several mechanisms to counteract and antagonize innate immunity pathways such as OAS/RNase L in order to successfully replicate and spread. With OAS/RNase L being one of the principal antiviral mechanisms, many viruses encode proteins that interfere with the pathway and inhibit activation of RNase L. What’s interesting is that these viruses belong to different families, a fact that also confirms the broad range antiviral activity of

RNase L.

1.3.2.1 Viral proteins that bind RNase L

There are several viral proteins that directly bind RNase L to prevent its endoribonuclease activity. Some of group C enteroviruses such as poliovirus contain a phylogenetically conserved RNA region that binds to and competitively inhibits RNase L activation, also known as ciRNA (Townsend et al., 2008a). The conserved structures consist of conserved H-H kissing loops that are key to

RNase L inhibition (Townsend et al., 2008b). Another picornavirus such as

Theiler’s encephalomyelitis virus also encodes a L* protein, from an alternative reading frame to L protein, which binds the ankyrin domain of murine RNase L, as mentioned above this domain is involved in 2-5A binding

1.3.2.2 Viral encoded dsRNA binding proteins

The NS1 protein of influenza A virus (IAV) is a critical component of IAV infectivity and replication. It contains an N-terminal RNA-binding domain with a nuclear localization signal (NLS) and a C-terminal effector domain. This protein manipulates host cell innate immunity responses in several ways such as by inhibiting IRF3, NFκB, PKR activation and inhibiting transcription of antiviral genes Reviewed: (Engel, 2013, Krug, 2015). Additionally, NS1 inhibits

OAS/RNase L by binding to dsRNA, the activator of OAS. The arginine in position 38 is critical for this binding. Viral titers were significantly decrease in a

R38A ns1 mutant virus, in cells expressing RNase L as compared to cells lacking

RNase L where there was no difference between titers of wild type and mutant

viruses (Min and Krug, 2006). This implies that prevention of OAS and RNase L activation are major functions of the NS1 protein. RNase L inhibits IAV replication by cleaving the genomic viral RNA in locations coding important viral proteins involved in replication (Cooper et al., 2015). Similarly, another viral protein which counteracts OAS/RNase L and other innate immunity responses by dsRNA binding is E3L of vaccinia virus (Xiang et al., 2002, Beattie et al., 1995,

Rivas et al., 1998). Virus replication was significantly impaired by PKR and

RNase L when E3L was mutated. Importantly, absence of E3L protein also increased activation of IRF3 and IFN-β independent of RNase L, PKR or Mx1

(Xiang et al., 2002).

The protein of gene 7 of transmissible gastroenteritis coronavirus has deveopled a different mechanism against innate immunity. It associates with protein phosphatase 1 to inhibit eIF2 phosphorylation and RNA degradation which correspond to PKR and RNase L activity, respectively (Cruz et al., 2011a).

In addition to the viral antagonists of OAS/RNase L mentioned above, recent studies in Dr. Susan Weiss and Dr. Robert Silverman laboratories have identified a novel mechanism of the 2H-Phosphoesterase family of proteins that degrade 2-5A.

1.4 The 2H phosphodiesterase superfamily of proteins that degrade 2-5A

1.4.1 The 2H-Phosphoesterase superfamily

The 2H-Phosphoesterases are a family of proteins characterized by two conserved histidine residues in their active site (two H-Φ-[ST]-Φ motifs, where Φ is a hydrophobic residue) separated by an average of 80 residues) (Mazumder et al., 2002). Some 2H phosphoesterases have 2’,3’ cyclic phosphodiesterase

(CPD) activity. They were originally described in yeasts where the phosphodiesterase is involved in tRNA intron splicing (Culver et al., 1994,

Phizicky et al., 1992). Further characterization of these proteins in yeast and plants showed that they contain two conserved histidines (for this reason they are named 2H-phosphoesterases) and serine/threonine motifs that are important for their catalytical activity (Hofmann et al., 2000, Nasr and Filipowicz, 2000,

Hofmann et al., 2002). The phosphodiesterase mechanism of these proteins seems to involve interaction of ta conserved His with a water molecule to break down the cyclic phosphate bond (Nasr and Filipowicz, 2000), while the conserved Ser/Thr are proposed to be involved directly in stabilizing the substrate (Hofmann et al., 2000).

Although initially described in yeasts, there are more members added to the 2H-superfamily of proteins in other eukaryotes and even viruses. They are all characterized by conserved 2H and threonine in 86% of the motifs. A subclassification of the new members divided them into 12 families. Many of these novel proteins share significant homology to Escherichia coli LigT enzyme,

making the Lig-T like groups of archaeo-bacterial or eukaryotic-viral proteins

(Mazumder et al., 2002).

1.4.2 The Eukaryotic-viral Lig-T family of proteins

The members of this family of proteins are very diverse in their origins and functions. Below, I will describe the known properties and functions of some of the members.

1.4.2.1 The viral Lig-T 2H-phospoesterases that degrade 2-5A: antagonism of OAS/RNase L by viral phosphodiesterases

The members of the viral Lig-T proteins are encoded by two viruses that are evolutionary unrelated. They are the nonstructural protein 2 (ns2) encoded by group 2a coronaviruses such as: human coronavirus OC43, mouse hepatitis virus (MHV), and enteric HEC4408; and the viral protein 3 (VP3) of group A rotavirus (reviewed in Silverman and Weiss, 2014). They are characterized by the two conserved His-X-Thr/Ser motifs in the phosphodiesterase active site.

Recently, these proteins with homologous PDE activities have been identified as antagonists of the OAS/RNase L (Figure 1.5A).

MHV, a coronavirus with a positive RNA strand genome, is an important pathogen in mice. The ns2 of MHV is a 30kDa protein (Figure 1.5B), encoded by the open reading frame 2a gene, localizes primarily in cytoplasm (Zoltick et al.,

1990). Initial evidence with the A59 strain of MHV showed that missense mutations affecting either or both of the conserved two histidines of ns2 significantly reduced viral pathogenicity in liver of wild type, but not of RNase L-/- mice. Although ns2 was important in liver pathogenesis and replication of the virus in this organ, no effect was observed in brain (Roth-Cross et al., 2009).

Indeed, ns2 catalytic site mutants H46A and H126R of A59 and a ns2 deletion mutant JHM.WUd ns2 could not replicate in livers of infected mice. The mutant viruses did not have a different effect from the parental virus on IFN production, but instead the mutant viruses were significantly more sensitive to IFN signaling in macrophages/microglia only and not in other cell types, making this a cell-type specific effect (Zhao et al., 2011). Although initially proposed to have a cyclic nucleotide phosphodiesterase (CPD) activity, ns2 did not have such an effect on any of the cyclic nucleotide substrates such as: ADP-ribose, 1”,2” cyclic phosphate, 2’,3’cAMP or 3’,5’ cAMP raising the possibility that ns2 acted on a different type of substrate (Zhao et al., 2012b).

Furthermore the sensitivity of ns2 mutant MHV to IFN signaling was shown to be unaffected by the absence of ISGs such as: ISG15, IFIT1, IFIT2 or

PKR but instead it required OAS/RNase L pathway activation. The mutant ns2H126R virus could replicate to the same levels as wild type virus in RNase L-/- deficient BMMs. The LigT family of eukaryotic-viral proteins, which includes ns2, belongs has an ancestral member in Escherichia coli, LigT, which has a 2’-5’ ligase as well as 2’,5’-PDE activity (Mazumder et al., 2002). The activator of

RNase L, 2-5A, is characterized by 2’-5’ bonds hence it was proposed that ns2 might degrade 2-5A. This effect was confirmed in vivo, as viral titers of ns2H126R virus were around three log10 units lower than those of wild type virus in B6 mice; while they remained the same in mice lacking RNase L. A similar effect was not observed in lack of PKR expression, demonstrating that the difference was directly related to RNase L. Expression of ns2 protein in cells inhibited rRNA cleavage (a hallmark of RNase L activation) and 2-5A accumulation upon IC transfection. Similiarly, the levels of 2-5A in BMMs infected with wild type A59 virus were much higher compared to the levels of 2-5A from cells infected with ns2H126R virus, confirming that ns2 was an antagonist of RNase

L activity by degrading 2-5A. In vitro studies of 2-5A integrity in presence of ns2 showed that the viral protein degrades 2-5A into ATP and AMP. As previously mentioned that ns2 was essential for liver pathogenesis in wild type mice where the replication of ns2H126R was very low but replication and pathogenicity of the mutant virus was restored in mice lacking RNase L. This replication difference due to ns2 and RNase L inhibition was also accompanied by virus-mediated necrosis, apoptosis and inflammation (hepatitis). Therefore, MHV encodes the

2H-PDE ns2 protein to degrade 2-5A hence antagonize the antiviral factor

RNase L in order to successfully replicate (Zhao et al., 2012a). These studies on ns2 introduced a novel mechanism for how a viral protein, member of the LigT family of 2H-phosphoesterases, could antagonize OAS/RNase L to evade innate immunity.

Another viral member of the 2H-phosphoesterase family is the VP3 protein of rotavirus A (RVA) (Mazumder et al., 2002, Zhang et al., 2013). Rotaviruses have segmented dsRNA genomes and are a common pathogen in young children, causing gastroenteritis. Rotaviruses account for ~450,000 deaths worldwide every year, largely in underdeveloped countries in which vaccination rates are low (Tate et al., 2012). Even though evolutionary unrelated to ssRNA genome coronaviruses, the C-terminal domain (CTD) of VP3 shares a structure homology of the two His-X-Thr/Ser motif in ns2 of MHV. The CTD consisting of around 143 amino acids, was shown to antagonize RNase L activity in a similar manner to ns2 (Zhang et al., 2013). The purified recombinant VP3-CTD protein in vitro degraded the trimeric form of 2-5A, 2’,5’-p3A3, into ATP and AMP. While when expressed in intact cells, VP3 significantly reduced 2-5A levels induced by transfection of synthetic dsRNA, polyI:polyC. Because of the lack of reverse genetic system for rotaviruses, the VP3-CTD was inserted in a chimeric MHV in place of the nonessential ns4 gene. The ns2 of the chimeras was mutated in its ns2H126R form, which as described above is a catalytically inactive form. The VP3-

CTD chimeric A59 was able to restore replication in BMMs as well as in vivo, in levels comparable to wild type virus, while chimeras expressing mutant catalytic histidines H718A and H797R of VP3-CTD were unable to replicate. As expected there was no difference in the replication of mutant viruses in cells lacking RNase

L, confirming that RNase L inhibition was the target of rotavirus VP3 (Zhang et al., 2013)(Figure 1.5A).

VP3 is a ~98kDa structural protein also shown to have guanylyltransferase and methyl-transferase activities in its N-terminal domain and the PDE in the C- terminal domain (as described above) (Figure 1.5B). Biochemical and bioinformatics studies on structure of this protein have characterized the PDE domain. This domain consists of an α/β concave fold of 2 α-helices and 7 β- sheets, in VP3-CTD of strain SA11 of simian RVA. The groove is positively charged and made of β2 and β5 strands with each containing the His-X-Thr motifs (Brandmann and Jinek, 2015). Similarly, the VP3-CTD of another strain simian RRV also contained 3 α-helices and 7 β-strands with the conserved motifs in β2 and β5. The groove is positively charged and accommodates the negatively charged 2-5A substrate. In rotavirus A VP3-CTD, the conserved Hix-x-Thr motifs consists of histidines H718A and H797A, as well as threonines T720 and T799.

T720, H797 and T799 form hydrogen bonds with substrate while the catalytical histidines stabilize the ligand. As predicted, mutational studies determined essential roles for the catalytic histidines H718 and H797 (Ogden et al., 2015).

Figure 1.5 Viral antagonists of OAS/RNase L

Modified from: (Zhang et al., 2013)

A. Viral 2H-phosphoesterases ns2 and VP3 antagonize RNase L

B. Ns2 and VP3 contain a 2H PDE domain

1.4.2.2 The eukaryotic Lig-T 2H-Phosphoesterases

The eukaryotic-viral LigT family members are suggested to be involved in

RNA metabolism, similar to their ancestral precursor proteins in prokaryotes. This family is typified by the cellular protein CGI-18 (Jung et al., 2002, Mazumder et al., 2002, Liang et al., 2008). Also known as activating signal cointegrator (ASC1)

1 complex subunit 1 (ASCC1) due to being part of ASC1, this protein is mainly involved in transcription regulation by enhancing nuclear factor kappa-B and activator protein 1 (Jung et al., 2002). Another cellular 2H-phosphoesterase as predicted by structure homology in Mazumder at al 2002 is the leukocyte receptor cluster member 9 (Leng 9). A member of the CCCH zinc finger protein family involved in macrophage activation (Liang et al., 2008). Leng9 contains a predicted RNA binding domain between CCCH and 2H-phosphoesterase domain

(Mazumder et al., 2002).

In addition a newly recognized member, the exoribonuclease USB1, was suggested to be also a LigT 2H phosphoesterase. Crystal structure of this protein showed that it contains two antiparallel strands β3 and β8 with each of them having a His-X-Ser motif (Hilcenko et al., 2013). Moreover, USB1 was also identified by a DALI server search based on the structure similiarities to rotavirus

VP3 protein PDE domain (Brandmann and Jinek, 2015). This evolutionarily conserved protein is a 3’-5’ exonuclease involved in U6 small nuclear RNA (a component of splicosome) processing, by controlling the length of U6 oligoU tail and generating a 3’ terminal phosphate (Mroczek and Dziembowski, 2013, 37

Hilcenko et al., 2013). Moreover, the histidines of the conserved 2H PDE domain were essential for this phosphodiesterase function in vitro and in vivo (Hilcenko et al., 2013). USB1 protein is encoded by the gene C16orf57, mutations of which have been linked to poikiloderma with neutropenia, an autosomal recessive skin disease (Volpi et al., 2010, Mroczek and Dziembowski, 2013, Hilcenko et al.,

2013). The other typical member of the cellular LigT family, as predicted and picked by structural comparison to CGI-18 is a member of the superfamily of A- kinase anchoring proteins (AKAP), which will be extensively described in the next section as it is the main subject of my thesis research.

The viral 2H PDEs have been shown to be antagonists of the OAS/RNase

L. On the other end, there is very little evidence concerning the functions of the above mentioned cellular PDEs. In particular prior to my studies AKAP7 was not known to function in controlling RNase L activity.

1.5 The A-Kinase Anchoring Protein 7

1.5.1 AKAPs: a superfamily of proteins with multiple functions

The AKAP is a large and diverse family of proteins with the general function of tethering the regulatory subunits of cAMP-dependent protein kinase A

(PKA) to different cellular compartments, bringing them closer to their substrates.

The PKA holoenzyme consists of two catalytic (C) and two regulatory (R) subunits where cAMP binds. AKAPs were initially discovered in agarose affinity columns as contaminants that bound to the R subunits (Theurkauf and Vallee,

1982). Today these signal transduction proteins are shown to be involved in a wide variety of functions and are therapeutic targets for several diseases

[Reviewed (Welch et al., 2010, Esseltine and Scott, 2013)]. The family consists of

43 members, some of which have more than one isoform due to alternative splicing events. Still all these proteins are functionally related by the ability to bind and anchor PKA. The binding occurs between an α-helical region in AKAPs and the R subunit of PKA, mostly RII subunit which has a higher affinity to the anchoring proteins (Fink et al., 2001, Hausken et al., 1994, Herberg et al., 2000,

Carr et al., 1991). By tethering PKA to a variety of targets, AKAPs are important components of diverse processes such as: cardiac excitation-contraction coupling, insulin secretion, renal homeostasis, reproduction, neuronal synaptic plasticity (Fink et al., 2001, Rosenmund et al., 1994, Klussmann et al., 2000).

Hence AKAPs are important targets of different diseases and conditions

[Reviewed: (Troger et al., Esseltine and Scott, 2013)]. Simultaneously, AKAPs

also bind other molecules such as kinases, phosphatases and phosphodiesterases. Binding of AKAPs to PDEs is related to their ability to control cAMP signaling activity and distribution [Reviewed (Stangherlin and

Zaccolo, 2011)].

Among the 43 members of the family and their isoforms, the A-kinase anchoring protein 7 (AKAP7) is the sole protein which has a 2H PDE domain with

2 conserved His-X-Thr/Ser motifs and is a member of the viral-eukaryotic LigT family.

1.5.2 AKAP7 as a PKA anchoring protein

The AKAP7 (also known as AKAP18) protein consists of four isoforms as a result of alternative splicing which are widely distributed in many tissues and organs (Trotter et al., 1999). The smaller isoforms: α and β, of 15kDa and 18kDa respectively, contain a PKA-R binding motif and a membrane targeting region.

While the larger isoforms γ and δ (37kDa and 42kDa, respectively) contain the

PKA R binding motif, a nuclear localization signal (NLS) motif as well as a PDE domain with 2 conserved histidines (Figure 1.6 ). Evolutionary analyses suggested that the ancestral forms of AKAP7 are AKAP7α for the short isoforms, which is conserved from yeast to humans; and AKAP7γ for the longer ones.

AKAP7 δ is proposed to have formed as a result of an insertion of a 19- nucleotide exon upstream of exon 5 in the γ isoform from cow to humans

(Johnson et al., 2012). AKAP7 γ and δ PDE domain is a 2H-phosphoesterase

(Mazumder 2002) and my study focuses only in these two isoforms.

Interestingly, AKAP7 γ isoform was the the first recognized AKAP that binds with a higher affinity to the RI subunit while also localizing to both nuclei and cytoplasm in oocytes (Brown 2003). In addition to PKA, AKAP7 possibly also regulates in tethering protein kinase C in different cellular compartments (Redden et al., 2012). The AKAP7 isoforms are involved in a variety of processesdue to their ability to bind the RII of PKA via its RII-binding domain. They are mostly shown to be important in cardiac contraction relaxation coupling. AKAP7δ is part of a supra molecular complex in heart sarcoplasmic reticulum which controls the adrenergic calcium (Ca+) re-uptake (Lygren et al., 2007). Indeed the AKAP7α was initially identified by co-purification with skeletal muscle L-type Ca+ channels

(Gray et al., 1997). AKAP7α anchors PKA to cardiac calcium channels through a leucine zipper domain interaction with the channels to regulate Ca+ current

(Hulme et al., 2002, Hulme et al., 2003). Moreover, AKAP7δ is proposed to regulate PKA phosphorylation in rat heart by directly binding to protein phosphatase 1 (PP1) and its inhibitor I-1 protein (Singh et al.). Surprisingly, studies in cardiomyocytes with a deleted version of AKAP7 exon 7 (which is responsible for the leucine zipper domain and the 3’UTR) responded normally to stimulation by a β-adregenic agonist. In addition, the mutant mice were phenotypically normal (Jones et al., 2012). That study suggested an alternative function for AKAP7 in addition to/instead of PKA regulation.

Although well described as a PKA-binding domain, prior to my studies little was known about the PDE domain in AKAP7 and its function. Structural homology classified this protein as a member of the ligT family of viral-eukaryotic

2H-phosphoesterases.

Figure 1.6 AKAP7 Isoforms

Modified from (Jones et al., 2012)

1.5.3 AKAP7 is a 2H-Phosphoesterase

Structural comparison to the CGI-18 as well as viral 2H PDEs such as

VP3, and ns2 classify AKAP7 a member of the LigT family (Mazumder et al.,

2002). X-ray crystallography identified a PDE central domain (CD) of AKAP7γ/ δ as four α-helixes and eight β-sheets. The β2 and β5 run antiparallel and each of them contains a conserved His-x-Thr motif (a characteristic of LigT family of 2H- phospoestrases as described above). The groove formed by the CD bound

5’AMP and 5’CMP and the binding involved hydrogen bonds to the two conserved motifs in the base of the groove. Although 2H PDEs are suggested to be CPDs, AKAP7 CD did not bind to any of the cyclic substrates and lacked CPD activity (Gold et al., 2008). Hence, it was possible that AKAP7 was a specific

PDE for some other type of substrate.

Importantly, structural homology was confirmed between AKAP7 PDE,

MHV ns2 and rotavirus VP3 (Gold 2008). The folded structure of AKAP7 CD was a model template for RVA VP3-CTD (Zhang et al., 2013) (Figure 1.7). A DALI database search for structure homology (Holm and Rosenstrom, 2010) selected

AKAP7 CD as a homolog of rotavirus VP3. The catalytic sites of both proteins were shown to contain the two conserved His-x-Thr motifs in equivalent positions

(Ogden et al., 2015, Brandmann and Jinek, 2015).

Figure 1.7 Mammalian AKAP7 is structurally homologous to viral ns2 of

MHV and VP3 of rotavirus

Modified from (Zhang et al., 2013)

Structure homology of AKAP7 to viral antagonists of OAS/RNase L suggested the possibility that cellular AKAP7 could also have a similar effect on

2-5A integrity. In my thesis work, I have identified murine AKAP7 as the first mammalian encoded 2H PDE that degrades 2-5A, the activator of antiviral

RNase L. I have also tested other candidate cellular PDEs for effects on 2-5A stability. From my experimental work, AKAP7 was the only member of the family which degrades 2-5A in vitro. Hence this work may suggest a role of AKAP7 in antiviral immunity and other functions of RNase L. Identification of cellular antagonists of OAS/RNase L could in the future guide attempts to enhance the antiviral response or to control cell survival. At the same time, these data suggest an evolutionary process in which host genes are acquired by viruses for the purpose of evading innate immunity.

CHAPTER II

MURINE AKAP7 HAS A 2’,5’-PHOSPHODIESTERASE DOMAIN THAT

CAN COMPLEMENT AN INACTIVE MURINE CORONAVIRUS ns2 GENE *

Elona Gusho1,2, #, Rong Zhang3, #, Babal K. Jha1, #, Joshua M. Thornbrough3,

Beihua Dong1, Christina Gaughan1, Ruth Elliott3, Susan R. Weiss3, and Robert H.

Silverman1

1Department of Cancer Biology, Lerner Research Institute, Cleveland Clinic,

Cleveland, Ohio, USA

2Department of Biological, Geological and Environmental Sciences, Cleveland

State University, Cleveland, Ohio, USA

3Department of Microbiology, Perelman School of Medicine, University of

Pennsylvania, Philadelphia, Pennsylvania, USA

#These authors were equal contributors

*Appeared in mBIO as (Gusho et al., 2014)

2.1 Introduction

Host antiviral pathways triggered by type I interferons (IFNs), are self- limiting so that after virus is eliminated the host can restore normal cellular and tissue functions (Alexander and Hilton, 2004). Many types of viruses also prevent activation of host antiviral pathways [reviewed in (Randall and

Goodbourn, 2008)]. The 2’,5’-oligoadenylate (2-5A) synthetase (OAS)/RNase L system is one of the principal mediators of the IFN antiviral response [reviewed in

(Silverman, 2007, Chakrabarti et al., 2011, Silverman and Weiss, 2014b, Sadler and Williams, 2008)]. Recently, we reported that two homologous viral proteins from unrelated viruses, the coronavirus mouse hepatitis virus (MHV) strain A59 ns2 and group A rotavirus strain SA11 VP3, express 2’,5’-PDE activities that antagonize the antiviral activity of RNase L by degrading 2-5A [px(5’A2’p)n5’A, x=1 to 3; n=2 or greater](Zhao et al., 2012b, Zhang et al., 2013). Ns2 and VP3 are eukaryotic-viral LigT-like family members that include both viral and cellular proteins of diverse origins, some of which possess cyclic nucleotide phosphodiesterase (CPD) activity (Figure. 2.1A) (Mazumder et al., 2002). Lig-T proteins are named for the prototypical archaeo-bacterial tRNA ligating enzyme

LigT with reversible 2’-5’ RNA ligase activity (Arn and Abelson, 1996) and are part of a larger superfamily of 2H phosphoesterases characterized by the presence of a pair of conserved His-h-Thr/Ser-h motifs (h, typically a hydrophobic residue) (Mazumder et al., 2002, Snijder et al., 2003, Roth-Cross et al., 2009).

However, while MHV ns2 has 2’,5’-phosphodiesterase (PDE) activity, it apparently lacks CPD activity based on its inability to cleave 2',3' cAMP, 3',5'

cAMP, and ADP-ribose 1",2" cyclic phosphate (Zhao et al., 2012b). Mutation of the active site of ns2 blocked MHV replication in liver thereby preventing hepatitis in wild type (wt) mice, but not in Rnasel-/- mice (Zhao et al., 2012b).

Furthermore, the group A rotavirus (strain SA11) VP3 C-terminal domain (CTD) was able to restore replication and virulence of a chimeric ns2 mutant MHV in mice (Zhang et al., 2013), showing that these two virally-encoded activities are functionally equivalent. Homologous PDEs encoded by other 2a betacoronaviruses, toroviruses and group A rotaviruses suggest this is a general mechanism of host antagonism necessary for replication of many RNA viruses

(Mazumder et al., 2002, Snijder et al., Zhang et al., 2013, Silverman and Weiss,

2014b) (unpublished data).

A-kinase anchoring proteins (AKAPs) are a family of scaffolding proteins that bind the regulatory (R) subunits of protein kinase A (PKA) to localize, coordinate and regulate cAMP signaling during diverse processes including cardiac excitation-contraction coupling, neuronal synaptic plasticity, sperm motility, insulin secretion and renal homeostasis [reviewed in (Welch et al., 2010,

Mauban et al., 2009, McConnachie et al., 2006, Diviani et al., 2011)]. The muscle-specific AKAP, mAKAP, partners with a cAMP-specific phosphodiesterase, PDE4D3, which limits activation of PKA (Dodge et al., 2001).

Several other AKAPs also bind to different cyclic nucleotide PDEs [reviewed in

(Diviani et al., 2011)]. However, among more than 43 known AKAP family members (Welch et al., 2010), only long isoforms of AKAP7 (also known as

AKAP15 or 18) have a central domain (CD) with two characteristic His-h-Thr/Ser- h motifs and predicted structural homology to the viral 2’,5’-PDEs, MHV ns2 and rotavirus VP3 (Figures 2.1A and 2.7) (Roth-Cross et al., 2009, Gold et al., 2008,

Mazumder et al., 2002). Therefore, we explored the possibility that instead of binding an extrinsic PDE similar to other AKAPs, AKAP7 might have an intrinsic

PDE activity.

There are four splice variants of AKAP7, two short forms of 15 and 18 kDa

( and ) that have a membrane targeting region and the PKA binding motif but lack the NLS and CD and two long forms of 37 and 42 kDa ( and ) which contain (from the N- to the C- teminus) a nuclear localization signal (NLS), the

CD, and the AKAP helix and leucine zipper that bind R subunits of PKA and also

Ca+2 and Na+ channels (Gold et al., 2008) A recent report described a cre/loxP knockout in mice of AKAP7 exon 7 from all four AKAP7 splice variants(Jones et al., 2012). AKAP7 exon 7 encodes the C-terminal modified leucine zipper domain that binds RI/RII PKA subunits, Ca+2 and Na+ channels, as well as the 3’-

UTR. The AKAP7 deficient animals were, however, phenotypically normal. In particular, cardiomyocytes from AKAP7 deficient mice responded normally to adrenergic stimulation, leading the authors to suggest that another AKAP isoform performs this function (Jones et al., 2012).

Here we investigated the possible role of AKAP7 in an alternative biochemical function to regulating cAMP signaling, namely, regulating 2-5A 50

signaling to RNase L. We determined that the AKAP7 CD is a 2’,5’-PDE that rapidly degrades 2-5A. As a result, replication of an ns2 mutant MHV was restored by the AKAP7 CD or by an N-terminal truncated AKAP7 that retains the

CD, both of which localized to the cytosol, also the site of viral replication.

However, full length AKAP7 localized to the nucleus and failed to restore replication of ns2 mutant MHV. These studies identify a novel biochemical function of an AKAP family member and suggest that the AKAP7 CD maybe be the evolutionary precursor of viral 2’,5’-PDEs that enable some viruses to evade the antiviral activity of type I IFNs by preventing activation of RNase L (Zhao et al., 2012b, Zhang et al., 2013). In addition, our findings suggest that localization of 2-5A degrading enzymes near sites of viral RNA synthesis may negate the antiviral activity of RNase L.

2.2 Materials and methods

Cell lines and mice. Murine 17Cl-1, L2 fibroblast, and BHK-MHVR cells were cultured as described previously (Roth-Cross et al., 2009, Yount et al.,

2002). Human 293T (ATCC) and human Hey1B (Baumal et al., 1986) cells were cultured in DMEM and RPMI media, respectively, both with 10% FBS. Primary

BMMs were generated from the hind limbs of C57BL/6 (B6) or Rnasel-/- mice and cultured as described previously (Caamano et al., 1999). C57Bl/6 (B6) mice were purchased from the Jackson Laboratory. Rnasel-/- mice (B6, ten

generations of backcrossing)(Zhou et al., 1997, Leitner et al., 2003) were bred in the University of Pennsylvania animal facility under an approved IACUC protocol.

Antibodies. The following antibodies were used: mouse anti-ns2 monoclonal antibody (provided by Dr. Stuart Siddell, University of Bristol, UK), mouse monoclonal anti-FLAG epitope (M2, Sigma), sheep anti-mouse IgG, HRP- linked whole antibody (GE Healthcare), horse anti-mouse IgG, HRP-linked antibody and goat anti-rabbit IgG, HRP-linked antibody (Cell Signaling), Alexa

Fluor 488 goat anti-mouse IgG, Alexa Fluor 594 goat anti-rabbit IgG secondary antibodies and Alexa Fluor 488 goat anti-rabbit IgG (Invitrogen), mouse anti- glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (US Biological), monoclonal anti-β-actin (Sigma-Aldrich, A1978), and custom-made affinity- purified rabbit polyclonal anti-AKAP7CD antibodies against peptide A:

CQLLNEDEVNIGTDALLELK; and peptide B: KKQSNGYYHCESSIVIGEK

(Biosynthesis). Peptide A is 100% homologous to murine AKAP7 (except for the

N-terminal cysteine residue) whereas peptide B is 100% homologous to both mouse and human AKAP7.

Plasmids. For protein purification, the murine AKAP7 full-length coding sequence (clone MmCD00295344 from the PlasmID DF/HCC Resource Core at

Harvard University (NCBI reference sequence NP_061217.3) was transferred from entry vector pENTR223.1 to a destination vector pDest-pGEX-6P-1 using

Gateway cloning technology (Life Technologies) resulting in pGEX-mAKAP7.

The histidine residues at position 93 and 185 were mutated with QuikChange II

XL Site-Directed Mutagenesis Kit (Agilent) to generate pGEX-AKAP7 (H93A;

H185R). Plasmid pMAL-ns2 was described previously(Zhao et al., 2012b). The cDNA encoding the C-terminal domain (CTD) of human rotavirus WA strain VP3

(AA 691-835) (Virus Sequence Database: JX406749) was optimized for bacterial expression, synthetically made (GeneScript, Piscataway, NJ) and cloned in pMal parallel vector and expressed as a maltose binding protein fusion protein with a

TEV cleavable tag, as we previously described for SA11 VP3-CTD (Zhang et al.,

2013). For Hey1b transient transfections, the murine AKAP7 full-length coding sequence was PCR amplified from entry vector pENTR223.1 and cloned into mammalian expression vector pCAGGS, generating pC-AKAP7. The histidine residues at position 93 and 185 were mutated to generate pC-AKAP7 (H93A;

H185R), and a FLAG tag sequence for the C-terminus was added. The CD region of murine AKAP7 (corresponding to amino acids 48-253) was PCR amplified from the full-length construct with a Flag epitope sequence added to the C-terminus and cloned into pCAGGS generating pC-AKAP7 CD. For 293T stable cells lines, lentivirus vector pLentiCMV-Puro-DEST, from Eric

Campeau(Campeau et al., 2009), and pCMV-VSV-G expressing VSV G envelope protein and pCMV-dR8.2 packaging plasmid, from Robert

Weinberg(Stewart et al., 2003), were obtained from Addgene. The murine

AKAP7 full-length cDNA insert was transferred from entry vector pENTR223.1 into destination vector pLentiCMV-Puro-DEST with Gateway cloning technology.

The CD region of murine AKAP7 (see above) was PCR amplified from the full-

length construct with a Flag epitope sequence added to the C-terminus and cloned into entry vector pENTR2B (Gateway, Life Technologies). The insert was transferred to destination vector pLentiCMV-Puro-DEST with Gateway technology. All of the clones were verified by nucleotide sequencing.

Viruses. Wt A59 and mutant ns2 virus ns2H126R were described previously(Zhang et al., 2013). The chimeric AKAP7 viruses were constructed based on the infectious cDNA clone icMHV-A59 (provided by Ralph S. Baric,

University of North Carolina at Chapel Hill)(Zhang et al., 2013, Yount et al.,

2002). Chimeric AKAP viruses all encode the mutant ns2H126R gene. To insert the

AKAP7 sequences in place of ns4, the BsmBI restriction site in AKAP7 gene was first removed by PCR-based site-directed mutagenesis with no change in coding sequence. The full-length AKAP7 (encoding 314 amino acids), AKAP7 with an N- terminal 47 amino acid deletion (AKAP7∆NTD, amino acids 48-314), and the CD of

AKAP7 (AKAP7CD, amino acids 48-253) were amplified and digested with SalI and NotI and cloned into icMHV-A59-fragment G as previously described for

VP3-CTD(Zhang et al., 2013). The mutant AKAPCD H185 had the histidine at position 185 exchanged for arginine (H185R, CAC to CGC), and cloned into the icMHV-A59 fragment G using the same strategy as for the wild type genes. All clones were confirmed by DNA sequencing. The full-length A59 genome cDNA was assembled and the recombinant viruses were recovered as previously described (Sperry et al., 2005, Yount et al., 2002, Zhang et al., 2013). Briefly, wild-type A-E plasmids, F plasmid with single mutation (H126R) in ns2 (Zhang et

al., 2013), and G plasmids with AKAP7 insertions were digested with appropriate restriction enzymes and the viral genome fragments were assembled in vitro to produce a full-length genome infectious DNA. Viral RNA transcripts were generated from the DNA by using the mMessage mMachine T7 transcription kit

(Ambion). The viral genome transcripts combined with the in vitro transcripts of viral nucleocapsid gene were electroporated into the BHK-MHVR cells with a Bio-

Rad Gene Pulser II electroporator. When virus cytopathology was observed, cell lysates were combined with the supernatant and virus plaque purified and amplified on 17Cl1 cells for use ( by Rong Zhang and Susan Weiss).

2’,5’-PDE activity assays. Proteins were expressed from the pGEX or pMAL constructs in the BL21DE3pLysS strain of E.coli (Life Technologies). Wt or mutant AKAP7 was cleaved from the GST fusion proteins bound to glutathione- sepharose beads with 30 units of human rhinovirus – HRV3C Protease

(PreScission Protease, GE Healthcare). The cleaved proteins were analyzed by

SDS-PAGE and staining with GelCode Blue Stain Reagent (Thermo Scientific).

The proteins were further purified by ion exchange chromatography using monoQ GL100 column on an AKTA purifier UPC (GE Healthcare Life Sciences).

The buffer in the pooled fractions was exchanged with assay buffer (20 mM

HEPES, pH7.2; 10 mM MgCl2, and 1 mM DTT) in Centriprep Centrifugal Filter

Devices (Millipore; 3000 MWCO #4302). MHV ns2 was expressed in bacteria and purified as described previously (Zhao et al., 2012b). The plasmid encoding

WA VP3-CTD was expressed in bacteria and the WA VP3-CTD was purified

essentially as described earlier for SA11 VP3-CTD (Zhang et al., 2013). Purified proteins (1.5 M of ns2, VP3, AKAP7, AKAP7H93A;H185R) were incubated with 10

M of (2’-5’)p3A3 as described previously (Zhao et al., 2012b) in assay buffer at

o o 22 C or 37 C. Degradation of the (2’-5’)p3A3 was determined by FRET-based

RNase L activity assays, in comparison to a standard curve with different concentrations of (2’-5’)p3A3(Thakur et al., 2005), or with high performance liquid chromatography (HPLC) as we described previously (Molinaro et al., 2006).

Degradation of 2-5A in pIC-transfected cells. Hey1B cells were plated at 4x105 cells/well in six well plates for 20 hour. Plasmid DNA (1 g) per well

(empty vector (pCAGGS), AKAP7CD, AKAP7 and FLAG tagged-AKAP7H93A;H185R, were transfected using lipofectamine 2000. After 16 hours cells were mock transfected or transfected with poly(rI):poly(rC) (pIC) (EMD Biosciences) at 1

µg/well for 3 hours. The cells were washed with PBS, lysed in buffer (50 mM Tris-

HCl, pH 7.2 ; 0.15 M NaCl, 1% NP-40, 200 µM sodium orthovanadate; 2 mM

o EDTA; 5 mM MgCl2; 5 mM DTT) and heated to 95 C for 7 minutes. Lysates were centrifuged for 10 minutes at 14,000g at room temperature and supernatants were passed through microcon centrifugal filters with a molecular weight cutoff of

3kDa (Millipore Corporation) for 45 minutes at 11000xg. Levels of 2-5A were determined by RNase L-based FRET assays in comparison to a standard curve of authentic (2’-5’)p3A3 as described (Thakur et al., 2005).

Viral growth kinetics. Viral growth curves were carried out by infecting

B6 or Rnasel-/- BMMs with each virus at an moi of 1 PFU/cell. After 1 hour incubation, the cells were washed with PBS and cultured with DMEM supplemented with 10% FBS. Culture supernatants were collected at 6, 9, 12 and 24 hours post-infection, and virus titers were determined by plaque assays on L2 cells(Hingley et al., 1994) (by Rong Zhang and Susan Weiss ).

RNA chip analysis. For analysis of rRNA integrity, total cellular RNA was isolated at 10 hours post-infection with the RNeasy kit (QIAGEN). RNA was quantified by Nanodrop analyzer and equal quantities of RNA were resolved on

RNA chips using an Agilent 2100 BioAnalyzer (Xiang et al., 2003) (by Rong

Zhang,and Susan Weiss).

Immunofluorescence assays. 17Cl-1 cells or B6 BMMs were infected with each virus at an moi of 1. At 9 hours post infection, cells were fixed with 4% paraformaldehyde in PBS followed by blocking with 2% BSA and 0.5% Triton X-

100 in PBS, incubated with the primary antibodies for one hour and then with secondary antibodies for 1 hour. Cell were stained with DAPI (Molecular Probes,

Eugene, OR), and visually analyzed by using an Olympus IX81 inverted fluorescence microscope and associated Slidebook 5.0 software (by Rong Zhang and Susan Weiss).

293T cells were transfected using Lipofectamine 2000 with empty lentivector, or lentivector containing cDNA to AKAP7 or AKAP7CD, together with

plasmid expressing VSV G envelope protein (from a pCMV vector) and pCMV dR8.2 packaging plasmid. Cells were selected with 3 µg/ml puromycin to generate stably expressing cell lines. 293T cell lines were plated on sterile cover slips at 2x105 cells/ well in six well plates. After 24 hours, the cells were washed with PBS, fixed with 4% paraformaldehyde in PBS for 20 min, blocked with 0.3%

Triton X-100, 1% BSA in PBS for 2 hours and incubated with primary anti-

AKAP7CD antibody peptide A at a 1:100 dilution at 4oC for 16 hours. The cells were washed three times with PBS and probed with secondary Alexa Fluor 488 goat anti-rabbit antibody at 1:500 dilution and stained with VECTASHIELD

Mounting Media with DAPI. Images were collected using an HCX PL APO

63X/1.4NA oil immersion objective on a Leica SP2 confocal microscope (Leica

Microsystems, GmbH, Weltzar, Germany).

Western blotting. Proteins in cell lysates were separated in 12.5% SDS polyacrylamide gels, and transferred to polyvinylidene difluoride (PVDF) membranes. Membranes were blocked and probed with antibodies described above and developed using Amersham ECL Western Blotting Detection Reagent

(GE Healthcare) and x ray film (Figures. 2.2 and 2.4C) or using using Western lightning Plus-ECL enhanced chemiluminescence substrate (Perkin Elmer), and proteins detected under an Intelligent dark box II (Fujifilm) (Figure. 2.3B). As controls for loading and transfer, the blots were probed with anti-glyceraldehyde

3-phosphate dehydrogenase (GAPDH) or -actin.

Animal Experiments. Four-week-old B6 or Rnasel-/- mice were anesthetized with isoflurane (IsoFlo; Abbott Laboratories), and inoculated intrahepatically (i.h.) with each virus (2000 PFU/mouse) in 50 µl of PBS containing 0.75% BSA. At 5 days post infection the mice were sacrificed and perfused with PBS. The livers were removed, homogenized, and titrated by plaque assays on L2 cells as previously described (Gombold et al., 1993). All mouse experiments were reviewed and approved by the University of

Pennsylvania IACUC. (by Rong Zhang and Susan Weiss).

Statistical Analysis. Two-tailed t-test was performed to determine statistical significance and the p values are shown in Figure 4. Data were analyzed with GraphPad Prism software (GraphPad Software, Inc., CA).

2.3 Results

Murine AKAP7 rapidly degrades 2-5A with similar kinetics as MHV ns2 and rotavirus VP3. Alignment of murine AKAP7 to its eukaryotic homologs suggests that it is a member of an ancient family of 2H phosphoesterases that extend from plants to humans (Figure. 2.8). To determine if murine AKAP7 is a

2’,5’-PDE, cDNA encoding the full length AKAP7 (subsequently, “AKAP7” unless stated otherwise) was expressed in bacteria, purified and tested for its ability to cleave the trimeric species of 2-5A, (2’-5’)p3A3, in vitro. Wild type (wt) AKAP7,

AKAP7 mutated in both conserved histidines (H93A;H185R), and, for comparison, MHV strain A59 ns2 and human rotavirus strain WA VP3-CTD

(Figure. 2.1A) were incubated with (2’-5’)p3A3 at 37 ˚ C. 2-5A levels were measured by activation of RNase L in vitro in comparison to a standard curve of

2-5A dilutions using a previously described fluorescence resonance energy transfer (FRET) method (Thakur et al., 2005). Ns2, VP3 and AKAP7 (each at 1.5

M) rapidly degraded (2’-5’)p3A3 (10 M) such that after 1 minute incubation at

o 37 C less than 30% of the input (2’-5’)p3A3 remain intact (Figure. 2.1B). Little or no detectable 2-5A remained after 10 minutes of incubation with any of the three proteins. In contrast, AKAP7 mutated in the two conserved histidine residues lacked the ability to degrade 2-5A (Figure. 2.1B).

To demonstrate that AKAP7 has a 2’,5’-PDE activity that cleaves one 5’-

AMP at a time from the 2’,3’-terminus of 2-5A, we performed incubations at a 60

lower temperature, 22oC. As we reported previously (Zhao et al., 2012b), ns2 degrades (2’-5’)p3A3 to (2’-5’)p3A2 and 5’-AMP and then the diadenylate (2’-

5’)p3A2 is degraded to 5’-AMP and 5’ATP (Figure. 2.1C). AKAP7 also removed one 5’-AMP at a time from trimeric 2-5A, and therefore it is also a bona fide 2’,5’-

PDE (Figure.2.1D). As expected, active site mutant AKAP7H93A;H185R failed to degrade (2’-5’)p3A3 (Figure.2.1E). Quantitation of these results showed that

AKAP7 and ns2 displayed comparable kinetics (Figure. 2.1F).

Figure 2.1 62

Figure 2.1 AKAP7 rapidly degrades 2-5A with similar kinetics as MHV strain

A59 ns2 and human rotavirus strain WA VP3-CTD

(A) Diagrams of MHV strain A59 ns2, human rotavirus strain WA VP3-CTD, mouse AKAP7 , and mutant AKAP7 H93A;H185R. NLS, nuclear localization signal;

PKA-R BD, protein kinase A RI and RII subunit binding domain; black filled regions represent undefined domains (B) Degradation of (2’-5’)p3A3 in vitro at

37oC by the different purified proteins (indicated) as determined by FRET assays.

Control, no protein added. Results are an average of three biological replicates and the error bars are the standard deviations (S.D.). (C-E) Purified (2’-5’)p3A3

(10 M) was incubated with 1.5 M purified ns2, AKAP7 or AKAP7H93A;H185R , respectively, at 22oC. At the times indicated (to the right), the reactions were stopped. The substrate, (2’-5’)p3A3 and its degradation products (2’-5’)p3A2, 5’-

AMP and 5’-ATP (as indicated) were separated by HPLC. Elution times are shown on the x-axes. (F) The percentage of intact (2’-5’)p3A3 remaining was determined by areas under the peaks from the chromatograms using 32 Karat™ software (Beckman Coulter, Inc.) and is plotted as a function of time.

AKAP7 degrades 2-5A in intact cells transfected with double stranded (ds) RNA. To determine if AKAP7 was able to degrade 2-5A in intact cells, cDNAs for full length AKAP7, AKAP7 CD and mutant AKAP7H93A;H185R (with a C-terminal flag epitope) were transiently expressed in the human ovarian carcinoma cell line, Hey1B(Baumal et al., 1986). Hey1B cells were selected for these experiments because endogenous AKAP7 was undetectable as determined by Western blot analysis with a rabbit polyclonal antibody against an

AKAP7 CD peptide (Figure. 2.2A). Ectopic expression of the different AKAP7 proteins was confirmed by immunoblotting (Figure.2.2A). At 20 hours post- transfection with either empty vector or with the AKAP7 cDNAs, cells were transfected for an additional 3 hours with the synthetic dsRNA, poly(rI):poly(rC)

(pIC), a potent activator of OAS(Kerr et al., 1977). 2-5A was undetectable in control cells transfected with vector alone as determined by FRET (lower limit of detection was about 15 fmol/106 cells) (Figure. 1H). However, pIC caused high levels of 2-5A (38 pmol/106 cells) to accumulate in the vector controls cells. In contrast, expression of AKAP7 CD reduced pIC-induction of 2-5A by almost 200- fold (to 0.2 pmol/106 cells). The full length AKAP7 reduced pIC-induced levels of

2-5A by about 12-fold (to 3 pmol/106 cells), while mutant full length

AKAP7H93A;H185R failed to deplete pIC-induced levels of 2-5A (26 pmol/106 cells remaining). These findings demonstrate that AKAP7 (full length and CD) effectively degraded 2-5A in intact pIC-transfected cells.

Figure 2.2 AKAP7 degrades 2-5A in intact cells

(A) Western blot analysis of AKAP7 proteins stably expressed in human Hey1B cells as determined by probing with a rabbit polyclonal antisera against AKAP7 peptide B (Materials and Methods). (B) Levels of 2-5A in Hey1B cells expressing different AKAP7 proteins (or vector control cells) transfected with pIC, as determined by FRET assays (Materials and Methods). Results are averages of three biological replicates. Error bars represent S.D.

Viral expression of AKAP7 in bone marrow-derived macrophages

(BMM). To determine if AKAP7 could functionally replace ns2, we further exploited an MHV-A59 reverse genetic system(Yount et al., 2002, Zhao et al.,

2012b, Zhang et al., 2013). We hypothesized that the expression of AKAP7 from a chimeric ns2 mutant MHV would restore viral replication in B6 BMM. Four

MHV-AKAP7 chimeric viruses, all based on the ns2H126R mutant MHV, were constructed (Figure. 2A) by inserting coding sequences for full-length

AKAP7 (ns2H126R-AKAP7), an N-terminal domain (NTD) truncation of AKAP7

(ns2H126R-AKAP7ΔNTD), the AKAP7 CD (ns2H126R-AKAP7CD), or a mutant AKAP7

CD (ns2H126R-AKAP7CD H185R) into the MHV nonessential ns4 gene (Figure. 2.3A).

BMM (Rnasel-/-) were infected with each chimeric virus, as well as with control wt A59 and ns2H126R, for 10 hours at an moi of 1. All viruses expressed similar levels of ns2 and AKAP7 proteins of the expected size in infected BMM as determined by Western blotting (Figure. 2.3B).

Figure 2.3 Construction of chimeric viruses expressing AKAP7 proteins.

A. Schematic diagram of the mutant ns2 and chimeric AKAP7 viruses. The

AKAP7cDNAs for flag-tagged full-length, N-terminal truncation, CD and its histidine mutant were inserted in place of ns4. The numbers above the boxes indicate the location of histidine residues critical for the function of ns2 or

AKAP7 proteins (histidine-to-arginine mutations are shown in the boxes). The location of the predicted NLS is also indicated. (B) Verification of the expression of MHV proteins ns2 and AKAP7 proteins. Rnasel-/- BMM were infected (1 PFU/cell) for about 10 hours, total cellular proteins were harvested and subjected to SDS-PAGE and Western blot analysis. Cellular GAPDH was monitored as a loading and transfer control.

Intracellular localization wt and mutant AKAP7 proteins. Full length

AKAP7 has an NLS sequence at its N-terminus which was deleted in the other

AKAP7 constructs (Figures. 2.1A & 2.3A). We compared the subcellular localization of the AKAP7 proteins expressed by each of the chimeric viruses.

Indirect immunofluorescence analysis with antibody against an AKAP7 CD peptide was carried out in 17CL-1 fibroblasts and BMM, neither of which expresses detectable level of endogenous AKAP7. The full-length AKAP7 was located in the nuclei of both 17Cl-1 and B6 BMM, as was previously reported

(Figures. 2.4A & B)(Brown et al., 2003). The N-terminal deletion (ΔNTD) including the NLS caused the truncated AKAP7 protein to localize exclusively to the cytoplasm of 17Cl-1, but to both the nuclei and cytoplasm of BMM (Figures.

2.4A&B, and data not shown). The CD of AKAP7 and its mutant (also missing the NLS) localized to the cytoplasm of 17Cl-1 cells but, similar to the NTD

AKAP7, localized to both the cytoplasm and nuclei of BMM. Likewise, the ns2 protein was clearly cytoplasmic in 17Cl-1 cells but present in both nuclei and cytoplasm of BMM.

To confirm these findings, human 293T cells were stably transfected with cDNA expressing full-length AKAP7 or the AKAP7 CD, or, as a control, the empty vector. Western blots showed that the 293T cells also failed to express detectable levels of endogenous AKAP7, whereas both AKAP7 and AKAP7 CD were clearly present in the transfected cells (Figure. 2.4C). As expected, full- length AKAP7 localized to the nuclei, whereas AKAP7 CD, which lacks the NLS, 68

localized to the cytoplasm (Figure. 2.4D). These studies show that expressed full-length AKAP7 is nuclear but deletion of its NLS causes some or all of the truncated protein to localize to the cytoplasm depending on the cell type.

Figure 2.4

Figure 2.4 Cellular localization of the AKAP7s in different cell types.

(A) 17Cl1 cells and (B) B6 BMMs were infected with chimeric MHV viruses expressing the full-length AKAP7, N-terminal truncation of AKAP7 (AKAP7∆NTD),

CD of AKAP7 (AKAP7CD) or its mutant (AKAP7CD H185R), and stained for the expression of ns2 and AKAP7 with monoclonal antibodies against ns2 or against murine AKAP7 CD, peptide A. DAPI was used to visualize nuclei. (C) 293T cells expressed AKAP7 or AKAPCD from lentivirus vectors(indicated). Western blot detection of AKAP7 and AKAP7 CD by immunoblotting with anti-AKAP7 CD antibody (peptide B) and with antibody against -actin as a loading and transfer control are shown. (D) 293T cell lines expressing AKAP7 or AKAP7 CD (as in panel C) were stained with a polyclonal antisera against murine AKAP7 CD peptide A. Nuclei were visualized with DAPI staining.

AKAP7 polypeptides that localize to the cytoplasm restore the replicative capacity of ns2 mutant MHV in BMM of wt B6 mice. To determine the effect of AKAP7 on viral replication, BMM from wt B6 or Rnasel-/- mice were infected with AKAP7 chimeric viruses or with control wt A59 and mutant ns2H126R.

H126R As expected, the replication of ns2 was severely impaired (by three log10 units) when compared to the wt A59 in B6 BMM (Figure. 2.5A), consistent with our previous studies(Zhao et al., 2012b, Zhang et al., 2013, Zhao et al., 2011).

Intriguingly, chimeric ns2H126R-AKAP7 was unable to rescue replication in B6

BMM demonstrating that the expression of full length AKAP7 does not confer a wild type level of replication in B6 BMM. In contrast, viruses expressing either the N-terminal deleted AKAP7 (ns2H126R-AKAP7ΔNTD) or AKAP7 CD (ns2H126R-

AKAP7CD) replicated to similar levels as wt A59 showing that either of these truncated forms of AKAP7 can compensate for the inactive ns2H126R PDE.

However, when catalytic histidine residue number 185 in the AKAP7 CD was mutated, replication of the chimeric virus ns2H126R-AKAP7CD H185R was impaired to a similar extent as with the ns2H126R mutant. As expected, however, all of these chimeric viruses replicated efficiently, with similar kinetics and to a similar extent as wt A59 in BMM derived from Rnasel-/- mice (Figure. 2.5B). These results show that expression of the N-terminal truncation, or the CD of AKAP7, both of which lack the NLS and are present in the cytoplasm, can restore the function of ns2 in the mutant virus, whereas full length AKAP7, which localizes to the nuclei, cannot. In addition, the ability of AKAP7 CD to rescue ns2 mutant MHV is dependent on a catalytic histidine residue.

Expression of the N-terminal deleted AKAP7 or the CD of AKAP7 inhibits RNase L-mediated rRNA degradation induced by the ns2 mutant virus in B6 BMM. To further investigate whether the restoration of ns2 mutant virus replication by AKAP7 CD or N-terminal deleted AKAP7 expression is due to antagonism of RNase L activity, we analyzed the integrity of rRNA in BMM infected with each recombinant virus. RNase L cleaves rRNA in intact ribosomes resulting in a characteristic set of discrete rRNA cleavage products(Silverman et al., 1983, Wreschner et al., 1981). BMMs infected with each virus were lysed at

10 hours post infection, RNA was extracted and analyzed by RNA chip (2.2

Methods)(Xiang et al., 2003). The chimeric viruses expressing either the AKAP7

N-terminal truncated protein or the CD prevented rRNA degradation by RNase L, similarly to wt A59, while cells infected with the chimeras expressing full length

AKAP7 degraded RNA to a similar extent as the ns2H126R mutant or the double mutant ns2H126R-AKAP7CD H185R (Figure. 2.5C). These findings show that an

AKAP7 polypeptide containing the PDE domain but lacking the NLS prevents activation of RNase L in the cytoplasm, whereas full-length AKAP7, which localizes to nucleus, or mutant AKAP7 CD, does not.

Figure 2.5

Figure 2.5 Expression of the N-terminal truncation or the CD of AKAP7 restores the replication of ns2 mutant in vitro. Growth kinetics of chimeric

AKAP7 viruses were determined on BMMs from (A) B6 mice and (B) Rnasel-/- mice. BMMs were infected with each virus (as indicated) at an moi of 1. Samples of the cultured supernatant were taken at the indicated time points, and viral titers were determined by plaque assays. The data are from one representative experiment of at least two, each performed in triplicate. (C) AKAP7 CD and N- terminal truncated proteins inhibit rRNA degradation during viral infection. B6

BMMs were infected (MOI of 1), harvested at 10 hours post-infection and the integrity of total cellular RNA was analyzed on RNA chips (Agilent).

Expression of the AKAP7 CD enhances replication of ns2 mutant virus in liver. We have previously shown that ns2 mutant is highly attenuated for replication and pathogenesis in the liver of B6 mice but it replicated and induced hepatitis to a similar extent as wt A59 in Rnasel-/- mice(Zhao et al.,

2012b, Zhao et al., 2011, Roth-Cross et al., 2009). To determine if expression of

AKAP7 is able to compensate for an inactive ns2 and confer liver replication in vivo, B6 and Rnasel-/- mice were infected intrahepatically with viruses expressing AKAP7 CD or its mutant as well as A59 and ns2H126R. The virus titers in liver were determined at day 5 post-infection, the peak day for viral replication in this organ. As expected, ns2H126R virus replicated minimally in B6 mice but recovered to wt A59 virus replication levels in Rnasel-/- mice (Figure. 2.6A). The chimeric virus ns2H126R-AKAP7CD, replicated in B6 mice to a titer of 104 PFU/g tissue, while the isogenic AKAP7 mutant virus ns2H126R-AKAP7CD H185R failed to replicate above the level of detection (p< 0.001) (Figure. 2.6B). Both of the chimeric AKAP7 viruses replicated equally in Rnasel-/- mice, to a similar level as ns2H126R-AKAP7CD in B6 mice. These results suggested that expression of the active AKAP7 CD promotes the replication of ns2 mutant virus in liver of wt B6 mice. Furthermore, virus replication in the liver required the active site residue, histidine-185. Taken together, our findings suggest that the AKAP7 CD can restore the replication of mutant ns2 virus in vivo as a result of its 2’,5’-PDE activity that cleaves 2-5A thereby preventing RNase L activation in the cytoplasm.

Figure 2.6 Expression of the N-terminal truncation or the CD of

AKAP7 restores the replication of ns2 mutant in vivo

Four-week-old B6 or Rnasel-/- mice were infected 2000 PFU/mouse intrahepatically with (A) A59 and ns2H126R or (B) ns2H126R –AKAP7CD and the double mutant ns2H126R –AKAP7CD H185R. At day 5 post-infection, mice were sacrificed, and viral titers in the liver were determined by plaque assays. The dashed line represents the limit of detection, and error bars represent standard errors of means (n=5). Asterisks indicate that differences are statistically significant (**, p < 0.05). Data are derived from one representative experiment

Figure 2.7

Figure 2.7 Conservation of 2’,5’-phosphodiesterases among distantly related taxa. Selected mammalian AKAP7 (mouse NP_061217.3, rat

NP_001001801.1, human NP_057461.2), coronavirus ns2 (mouse MHV

P19738.1, human HCoV-OC43 AAT84352.1) and group A rotavirus VP3 C- terminal domain (simian RVA-SA11 AFK09591.1, human RVA-Wa AFR77808.1) proteins. Alignment wasperformed with the Constraint-based Multiple Alignment

Tool (COBALT) (Papadopoulos and Agarwala, 2007) set on default parameters and subsequently evaluated with T-Coffee Core to assess local reliability of the alignment by pairwise methods utilizing multiple algorithms (Mfast_pair,

Mmafft_msa and Mmuscle_msa) (Notredame et al., 2000). To convey confidence in the alignment, scores indicate the percent agreement between all MSA algorithms utilized in the analysis for each protein and for the consensus sequence. Additionally, color-coding indicates positional confidence of each aligned residue based on agreement between MSA algorithms with blue indicating the least and red indicating the greatest confidence in the alignment.Conservation of sequence is indicated by Clustal MSA symbols in the consensus line (cons) with “*”indicating fully conserved, “:” strong conservation of properties and “.” weak conservation of properties. Catalytic HxT motifs highlighted (blue boxes) (by Joshua M Thornbrough and Susan Weiss).

Figure 2.8 Protein multiple sequence alignment (MSA) of mAKAP7γ and eukaryotic homologs.

MSA of Mus musculus (mouse), Rattus norvegicus (rat), Homo sapiens (human),

Pan troglodytes (chimpanzee), Macaca mulatta (macaque), Nomascus leucogenys (gibbon), Callithrix jacchus (marmoset), Oryctolagus cuniculus

(rabbit), Bos taurus (cow), Ovis aries (sheep), Pantholops hodgsonii (antelope),

Orcinus orca (whale), Trichechus manatus latirostris (manatee), Melopsittacus undulatus (parakeet), Gallus gallus (chicken), Anolis carolinensis (lizard),

Alligator mississippiensis (alligator), Danio rerio (zebra fish), Ectocarpus

siliculosus (brown algae), Zea mays (corn). NCBI reference sequences

NP_061217.3, NP_001001801.1, NP_057461.2, XP_518739.2,

XP_001103953.2, XP_003255757.1, XP_002747004.1, XP_002714864.1,

NP_001095736.1,XP_004011367.1, XP_005967580.1, XP_004263875.1,

XP_004368981.1, XP_005154795.1,XP_004940279.1, XP_003215759.1,

XP_006259840.1, XP_005173856.1, CBN75660.1 and DAA52752.1, respectively. Alignment was performed with the T-Coffee Expresso, which incorporates known protein database structures to aid alignment, with all MSA options selected (pcma_msa, mafft_msa, clustalw_msa, dialigntx_msa, poa_msa, muscle_msa, probcons_msa, t_coffee_msa, amap_msa, kalign_msa, fsa_msa, mus4_msa) to ensure high confidence in the alignment To convey confidence in the alignment, scores indicate the percent agreement between all MSA algorithms utilized in the analysis for each protein and for the consensus sequence. Additionally,color-coding indicates positional confidence of each aligned residue based on agreement between MSA algorithms with blue indicating the least and red indicating the greatest confidence in the alignment. Conservation of sequence is indicated by Clustal MSA symbols in the consensus line (cons) with “*” indicating fully conserved, “:” strong conservation of properties and “.” Weak conservation of properties. Boxes indicate predicted nuclear localization signals (blue) (Kosugi et al., 2009, Kosugi et al., 2008), catalytic HxT motifs (yellow) and PKA RII-binding domain (black) (Hou et al.,

2011). ( by Joshua M Thornbrough and Susan Weiss )

Figure 2.9 Phylogenetic relationship between 2’,5’-phosphodiesterases of distantly related taxa. Protein sequences of Vertebrate AKAP7, rotavirus VP3, torovirus polyprotein 1a (pp1a) and coronavirus ns2 were analyzed by fast minimum evolution with a maximum sequence difference of 0.9 using the Grishin distance model. The tree is based on a COBALT alignment derived from a PSI

Blast search with 3 iterations of Murine hepatitis virus strain A59 ns2 protein

NCBI P19738.1. ( by Joshua M Thornbrough and Susan Weiss )

2.4 Discussion

Control of viral infections by regulation of 2-5A turnover. Our findings establish the AKAP7 CD as a 2’,5’-PDE that is able to substitute for a viral enzyme, MHV ns2, with the same activity. 2’,5’-PDEs that cleave 2-5A to ATP and AMP and phosphatases that remove the 5’-terminal phosphates on 2-5A prevent perpetual activation of RNase L after the viral infection is cleared and thereby limit RNA damage to the host(Williams et al., 1978, Trujillo et al.,

1988)[reviewed in (Silverman and Weiss, 2014b)]. In vitro, enzymes with 5’- phosphatase activity, such as alkaline phosphatase, can remove the 5’- triphosphate moiety of 2-5A thus eliminating, or greatly reducing, the ability of the core 2’,5’-oligoadenylate to activate RNase L(Dong et al., 1994). In addition, porcine coronavirus transmissible gastroenteritis virus (TGEV) gene 7 protein has been reported to dephosphorylate 2-5A through its interactions with protein phosphatase PP1(Cruz et al., 2011b). Two mammalian PDEs, PDE12 (2’-

PDE)(Kubota et al., 2004a, Poulsen et al., 2011a, Rorbach et al., 2011) in the exonuclease–endonuclease–phosphatase family of deadenylases and ectonucleotide pyrophosphatase/phosphodiesterase 1 (ENPP1)(Poulsen et al.,

2012b) have been shown to degrade 2-5A in vitro. PDE12 is a mitochondrial protein with both 2’,5’- and 3’,5’-PDE activities that removes poly(A) tails from mitochondrial mRNAs (Rorbach et al., 2011, Poulsen et al., 2011a). ENPP1 has a catalytic domain that is extracellular, and is therefore essentially an extracellular enzyme, and in addition to degrading 2-5A also cleaves

phosphodiester bonds in 3’,5’ RNA, DNA and cAMP (Poulsen et al., 2012b). In contrast to mitochondrial and extracellular locations of PDE12 and ENPP1, respectively, OAS proteins localize to either the cytoplasm or to nuclei, also sites of virus replication (Hovanessian et al., 1987, Chebath et al., 1987a). Our findings here demonstrate that AKAP7 is an additional host enzyme with 2’,5’-

PDE activity. Among the host 2’,5’-PDEs described to date, only AKAP7 is a 2H- phosphoesterase family member with homology to the viral enzymes of the same class.

A prior study with a commercial AKAP7 antibody (from ProteinTech) showed widespread expression of AKAP7 in different mouse organs including heart, brain, skeletal muscle, kidney and lung(Jones et al., 2012). Using the same antibody, we were able to detect low levels of AKAP7 in Hey1b cells and

BMM. However, AKAP7 mRNA is not induced by wt A59 or ns2 mutant infection

(unpublished data). Thus AKAP7 may be functional at a low expression level, may be expressed at higher level in some tissues (presently under investigation) and/or it may be induced by cytokines or other soluble mediators not induced by

MHV.

Cytoplasmic expression of the AKAP7 PDE rescues ns2 mutant MHV.

AKAP7 is one of a relatively few members of the AKAP family that localizes to the nucleus(Shanks et al., 2012, Brown et al., 2003) \l "_ENREF_full length AKAP7 traffics to nuclei, the AKAP7 central PDE domain typically

localizes to the cytoplasm, or to both cytoplasm and nuclei, after the N-terminal

NLS is deleted(Figure. 2.4) (Brown et al., 2003). 2-5A produced in the cytoplasm during viral infections might be expected to transit nuclear pores leading to its degradation by the endogenous nuclear AKAP7. However, expression of full length AKAP7 failed to rescue ns2 mutant MHV or to prevent RNase L activation in the infected wt BMM. In contrast, the AKAP7 CD rapidly degraded 2-5A preventing activation of RNase L and restoring the ability of an ns2 mutant MHV to replicate in vitro and in vivo. MHV RNA replication occurs completely in the cytoplasm of transformed cell lines (Wilhelmsen et al., 1981) and the local requirement for degradation of 2-5A may not be so surprising. Nilson and

Baglioni (Nilsen and Baglioni, 1979), proposed, based in part on their experiments with encephalomyocarditis virus, that microdomains (localized accumulation) of 2-5A occur at the sites of viral double-stranded replication intermediates (RI) where OAS binds and is activated causing localized activation of RNase L. Thus 2’,5’-PDE activity may be required in the same subcellular compartment as viral RNA replication to effectively antagonize RNase L activation as measured by virus rescue as well as protection of rRNA. These findings raise the possibility of an additional, yet-to-be identified isoform of

AKAP7 that retains the CD but localizes to the cytoplasm. Alternatively, AKAP7 could be relevant to viruses that replicate in the nuclei, but not to viruses such as coronaviruses or rotaviruses that replicate in the cytoplasm. It remains to be determined, however, whether the endogenous, nuclear AKAP7 has any role in viral infections. Alternatively, it is also possible that AKAP7 functions to degrade

nuclear 2-5A or 2-5A-like molecules that might be produced during non-viral cellular responses to stress(Reid et al., 1984). However, full length AKAP7 was able to degrade 2-5A in cells transfected with pIC, although less effectively than

AKAP7 CD, perhaps due to some leakage of the overexpressed AKAP7 into the cytoplasm (Figure.2.2B). Moreover, the intracellular localization of transfected pIC and the MHV RI are likely to be different. In addition, post-translational modifications, such as phosphorylation, could possibly cause AKAP7 to relocate to different intracellular sites as suggested for the AKAP species, (Shanks et al.,

2012).

The titers of ns2 mutant chimeric viruses with AKAP7 genes inserted in the place of MHV gene 4 are all significantly lower than those with the natural gene 4 sequences (Figures. 2.6A & B). This is similar to our findings with VP3-

CTD/MHV chimeric viruses which reached similar liver titers as the AKAP7-CD chimeras(Zhang et al., 2013). The reduction in viral titers is most likely due to attenuation associated with disturbing the genome in the region of gene 4, which is not required for viral replication or virulence(Ontiveros et al., 2001). This explanation is supported by our finding that a virus expressing wt ns2 and mutant

VP3-CTD from the gene 4 position is similarly attenuated as virus expressing mutant ns2 and wt VP3-CTD from the gene 4 position (data not shown).

Because the replication of these chimeric viruses in the liver is so much less robust than that of wt A59, it was not possible to determine whether expression of foreign PDEs fully confers hepatitis on the ns2H126R mutant.

Coronavirus replication is reported to occur only in the cytoplasm of infected cells (Wilhelmsen et al., 1981) and to involve the rearrangement of cellular membranes into double-membrane vesicles and convoluted membranes, the sites of viral RNA replication (Hagemeijer et al., 2012, Knoops et al., 2008).

As expected, in infected murine fibroblasts coronavirus proteins are localized into the cytoplasm as shown in Figure 2.4A. Most, if not all, studies of localization of

MHV encoded proteins or membrane rearrangements have been carried out in transformed cell lines and there is little information on infection of primary cells such as BMM. Interestingly, full length AKAP7 retains its exclusively nuclear localization in BMM whereas the N-terminal deleted AKAP7 and AKAP7 CD as well as ns2 were present in both cytoplasm and nuclei (Figure. 2.4B). Future studies will be directed at comparing subcellular localization of proteins and replication complexes in MHV-infected primary cells to those in transformed cell lines. Nevertheless, rescue of the ns2 mutant phenotype occurs only when at least some of the AKAP PDE is localized to the cytoplasm.

Relationship between viral and cellular eukaryotic-viral LigT-like members of the 2H phosphoesterase superfamily. Degradation of 2-5A appears to be a general strategy of many RNA viruses for preventing activation of RNase L that would otherwise block viral replication. Group 2a betacoronaviruses, which have plus RNA strand genomes, and the group A rotaviruses, with segmented dsRNA genomes, are unrelated RNA viruses with different replication strategies, yet both encode the related 2’,5’-PDEs, ns2 and

VP3, respectively (Zhao et al., 2012b, Zhang et al., 2013). Here we present the first evidence that a viral pathogen for humans, the rotavirus strain WA (Wyatt et al., 1980), also encodes a functional 2’,5’-PDE (Figure. 2.1B). In addition, many related viruses encode predicted or confirmed 2’,5’-PDEs including additional 2a betacoronaviruses human OC43 and HEC4408, bovine BCV, porcine hemagglutinating virus, the torovirus and coronavirus superfamily member, equine torovirus (Berne) and, group A, B and G rotaviruses [reviewed in

(Silverman and Weiss, 2014b)]. While the evolutionary origin of these viral 2’,5’-

PDEs are unknown, we suggest that the host AKAP7 2’,5’-PDE domain coding sequence might have been captured through RNA recombination during viral infections in the distant past. The phylogenetic relationship of different species of

AKAP7, VP3 and ns2 based on protein sequence data were explored by means of fast minimum evolution (Figure. 2.9). While this analysis shows a close relationship among different species of AKAP7, VP3 and ns2, it does not reveal whether this relationship is the result of divergent or convergent evolution

(Figure. 2.9). Coronaviruses mRNAs are transcribed from their genomes by a discontinuous process believed to involve switching of template by the viral replicase complex. During this process there is a high rate of homologous recombination, reportedly up to 25% observed during in vitro replication and in vivo [reviewed in (Lai, 1990)]; this high frequency template switching could result in low frequency copying of host mRNA. Indeed, sequences homologous to the

5’ end of MHC class I coding regions are found on the 5’ end of the MHV HE gene, adjacent to the 3’ end of the ns2 orf and it was previously speculated that

this was a result of homologous recombination between the MHV genome and host RNA (Luytjes et al., 1988). We suggest that AKAP7 PDE sequences could have been acquired by a similar process and subsequently evolved into ns2. In support of this idea, an alignment of the amino acid sequences of different isoforms of AKAP7, ns2 and VP3 show extensive homology that extends beyond the His-h-Thr/Ser-h motifs (Figure. 2.7). Subsequent to randomly acquiring the

AKAP7 CD coding sequence, the virus would have a selective advantage resulting in retention of the gene.

CHAPTER III

FURTHER CHARACTERIZATION OF AKAP7 AND OTHER

CELLULAR CANDIDATE 2’,5’-PDEs

3.1 Introduction

The type I IFN innate immune effectors make up the first barrier against virus infections. The OAS/RNase L pathway directly degrades viral ssRNA and cellular ssRNA to inhibit viral replication and spread. Several viruses have evolved different mechanisms to inhibit this pathway. The viral proteins ns2 of

MHV and VP3-CTDof rotavirus antagonize OAS/RNase L by degrading the 2-5A activator into ATP and AMP (Zhang et al., 2013, Zhao et al., 2012b). Similarly, we showed that a structurally homologous cellular protein AKAP7 also degrades

2-5A when expressed in ovarian carcinoma cell lines or when the bacterial purified protein is incubated with 2-5A. This function is depended on the two conserved histidines, as mutations affecting these amino acids render the protein

catalytically inactive. The AKAP7 central domain (CD) complemented the ns2 function in an ns2 mutant chimeric virus (Gusho et al., 2014) and Chapter 2 of this thesis. These newly identified 2’,5’-PDEs belong to the LigT phosphoesterase family of enzymes which are characterized by two conserved

His-x-Ser/Thr motifs, similar to bacterial 2’,5’ ligase which also acts as a phosphoesterase (Mazumder et al., 2002). This family consists of additional mammalian members such as LENG9 and CGI-18 which typifies the family and the recently described USB1 (Mazumder et al., 2002, Hilcenko et al., 2013).

It is unknown whether these proteins are similar to AKAP7 in the ability to degrade 2-5A (described in Chapter 2). Here I investigated the possibility that other cellular members of the 2H phosphoesterase family could have a similar effect on 2-5A, as the structurally homologous protein AKAP7. The data suggest that none of the three tested PDEs has an effect on 2-5A levels mediated by polyI:polyC (pIC) transfection. In addition, I have also further characterized the endogenous AKAP7 in intact cells. There were higher levels of 2-5A when expression of the endogenous AKAP7 is reduced. Furthermore, expression of

AKAP7 or its central domain (CD) had only minimal effects on EMCV virus replication. It increased replication of EMCV by 2-4 fold. These studies suggest that to date, AKAP7 is the only cellular member of the 2H phosphoesterase family which degrades 2-5A and hence may have an effect on RNase L activity.

3.2 Materials and methods

Antibodies. The primary antibodies used in the western blots were: mouse monoclonal anti-FLAG epitope (M2; Sigma); custom made AKAP7 antibody against peptide B (KKQSNGYYHCESSIVIGEK); anti-HA- Tag mouse monoclonal antibody (Cell Signaling, 2362); anti-PDE12 antibody (Abcam, ab87738); AKAP7 rabbit polyclonal antibody (Proteintech, 12591-1-AP), monoclonal anti-β-actin (Sigma-Aldrich, A1978); GAPDH loading control antibody

(Pierce, MA5-15738). The secondary antibodies used were: anti-rabbit IgG,

HRP-linked antibody (Cell Signaling, 7074); anti-mouse IgG, HRP-linked antibody (Cell Signaling, 7076); Biotin-SP AffiniPure Donkey Anti-Rabbit IgG

(JacksonImmunoResearch, 711-065-152). In addition Streptavidin-HRP (Cell

Signaling, 3999) was also used.

Cell Lines. Human ovarian carcinoma Hey1B cells were cultured in RPMI

1640 media (Baumal et al., 1986) while human alveolar adenocarcinoma A549 and mouse fibroblast L929 cells were cultured in Dulbecco’s modified Eagle’s medium. Media was supplemented with 10% heat inactivated fetal bovine serum and 1% penicillin-streptomycin.

Plasmids. The cDNAs of human wild type USB1 and the H208A mutant

USB1 (used for Hey1B transfection) were cloned in pCAGGS vector with an HA- tag added at the C-terminal end (cloning by Yize Li, Dr. Susan Weiss

Laboratory). While murine AKAP7 full length and AKAP7 CD was cloned as 95

described previously (Section 2.2). Viral proteins ns2 and VP3-CTD were cloned in pCAGGS as previously described (Zhang et al., 2013, Zhao et al.,

2012b).Human CGI-18 and Leng9 were PCR amplified from and pOTB7 clone

HsCD00331501 and pENTR223 clone HsCD00288777 respectively (PlasmID

DF/HCC Resource Core at Harvard University) and cloned in p3X-FLAG-CMV10 vector (Sigma). To generate Hey1B stable cell lines using lentivirus production, the plasmids used were: pCMV-VSV-G, expressing vesicular stomatitis virus G envelope protein and pCMV-dR8.2 packaging plasmid (Stewart et al.,

2003)(Addgene); human AKAP7 shRNA (NM_004842.2-729s21c1), human

PDE12 shRNA (NM_177966.5-1111s21c1) and pLKO.1-puro non-target shRNA control plasmid (Sigma). All the plasmids were sequence verified by nucleotide sequencing.

2-5A production assays. Hey1B cells were transfected with vector control, murine AKAP7 CD, human: USB1, mutUSB1, CGI-18 and Leng 9 followed by pIC transfection as previously described (Section 2.2). The cells were lysed in buffer (50 mM Tris-HCl, pH 7.2 ; 0.15 M NaCl, 1% NP-40, 200 µM sodium orthovanadate; 2 mM EDTA; 5 mM MgCl2; 5 mM DTT) followed by an incubation at 95oC for 7 minutes. Next, the lysates were centrifuged for 15 min at

13,000 rpm. An RNase L-based FRET assay was used to determine 2-5A levels.

A standard curve of authentic (2’-5’)p3A3 was used to quantify the 2-5A concentrations (Thakur et al., 2005).

Stable cell lines.Hey1B cells. 293T cells were plated in 10 cm dishes at

30% confluency. After 16h the cells were transfected with pLKO.1-puro empty vector control or the vector containing AKAP7 shRNA or PDE12 shRNA and the packaging plasmid,pCMV-dR8.2 as well as pCMV-VSV-G using Lipofectamine

2000 . The cellular supernatant was collected and used to infect ovarian carcinoma Hey1B cells with polybrene at 8µg/ml. Hey1B cells were selected with puromycin at 3ug/ml concentration to obtain stable cell lines with depleted levels of AKAP7 and PDE12. While the 293T stable cell lines expressing murine

AKAP7 and its CD were obtained as described in section 2.2.

Viral plaque assays. Hey1B cells were transiently transfected using

Lipofectamine 2000 with 2µg plasmid DNA of pCAGGS empty vector,

AKAP7,VP3-CTD or ns2. After 20h the cells were infected with EMVC virus at

0.01MOI in plain RPMI media. The media was replaced with 10% FBS containing

RPMI after 1h. The supernatants from the infected cells were collected at 10h post infection and used in plaque assay in L929 cells to determine viral titers.

293T stable cell lines were also infected by EMCV virus and analyzed in plaque assays in a similar manner.

RNA chip analysis. Hey1B cells were transfected with pIC at 1µg/ml concentration using Lipofectamine 2000. Total RNA was isolated using RNeasy kit (Qiagen) and analyzed on RNA chips in an Agilent 2100 Bioanalyzer.

Statistical analysis. A paired two tailed t-test was used to determine statistical significance (GraphPad Prism software). P values shown in Figure 3.1.

Western blotting. The cells were lysed in RIPA buffer containing protease inhibitor cocktail and separated in 12.5% SDS-polyacrylamide gels. The proteins were transferred to PVDF membranes in a semi-dry transfer and blocked for 1 hour with 5% milk in Tris-buffered saline, 0.1% Tween 20 (TBST), at room temperature. Next, the membranes were probed with primary antibodies

(described above) for 16 hours at 4˚C followed by secondary antibodies at

1:5000 dilution in 5% Bovine Serum Albumin (BSA). Amersham ECL Western

Blotting detection reagent (GE Healthcare) was used to develop the membranes.

We were unable to detect expression of human or mouse endogenous AKAP7 in cell lines or mouse tissues. This could be due to low expression of the protein in cells or low specificity of the available antibodies against AKAP7. To enhance the signal, a biotin-streptavidin method was used. After cell lysis, the proteins were separated in polyacrylamide gel and transferred to PVDF membrane as described above. Next, the membrane was blocked in 2% non-fat dry milk in

TBST (blocking buffer) at room temperature for 1 hour. The membrane was probed overnight at a 1:500 dilution in blocking buffer with AKAP7 primary antibody (Proteintech,12591-1-AP). Next, it was incubated with 1:5000 Biotin-SP

AffiniPure Donkey Anti-Rabbit followed by Streptavidin-HRP each for 1 hour in blocking buffer at room temperature. The membrane was washed with TBST 3

times (10min each) between the blockings. Finally the membrane was developed using Amersham ECL Western Blotting detection reagent.

3.3 Results

Candidate 2’,5’-PDEs CGI-18 and Leng9 do not degrade 2-5A in cells transfected with dsRNA. Studies on the viral and cellular 2’,5’-PDEs have led to identification of a new set of enzymes which have the ability to break down the phosphodiester bonds of 2-5A, and as a result control the ribonuclease activity of

RNase L. There is a possibility that other enzymes, members of the Lig T family,also characterized by the conserved His-x-Ser/Thr motifs, can have a similar function (described in Mazumder 2002). To test this hypothesis, human

CGI-18 and Leng9 were cloned in p3xFLAG vectors with a Flag epitope was added at their N terminal sequences. The constructs were transiently expressed in ovarian carcinoma Hey1B cell lines (Figure 3.1A). After 20 h, 2-5A production was induced by synthetic dsRNA transfection of polyI:polyC (pIC) for an additional 3.5h. As previously described in Chapter II, I used a FRET assay to measure the levels of 2-5A induced by pIC (Figure 3.1b). There was no significant difference in 2-5A concentrations between the empty vector control

(12.5 nM) and the other 2’,5’ candidate PDEs: human CGI18 at 10.1nM and

Leng9 10.8nM ( as suggested by statistical analysis). In contrast, when the

AKAP7 central domain (CD) was used as a positive control and there was

around 4 fold less 2-5A when this 2’,5’-PDE was expressed (Figure 3.1C).

These results suggested that although these proteins are all members of the LigT family of phosphodiesterases with homologous conserved Hix-x-Thr/Ser motifs,

CGI-18 and Leng9 do not appear to degrade 2-5A. Instead they may be specific to some other substrates.

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B C

Figure 3.1 Candidate cellular 2’,5’-PDEs do not degrade 2-5A. Hey1B cells were transfected with plasmid DNA followed by pIC transfection for 3.5h. (A)

Western blot analysis showing expression of the transfected plasmids (B) 2-5A standard curve for FRET assay (C) 2-5A concentrations as determined by

FRET assay. (CD p value 0.0004 )( CGI-18 p value 0.0690) (Leng9 p value

0.2500)(***, P< 0.005 )

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USB1 does not affect 2-5A levels in cells. USB1 is a newly identified member of the LigT family, which is involved in U6 oligoU tail processing

(Hilcenko et al., 2013). This protein is also characterized by the two conserved histidine motifs typical of 2H phosphoesterases including ns2, VP3-CTDand

AKAP7 (Zhao et al., 2012a, Zhang et al., 2013, Gusho et al., 2014). To determine whether these histidines in USB1 have an effect on the phosphodiester bonds of 2-5A, a point mutation was introduced in the protein.

The histidine at position 208 was mutated to alanine (by Yize Li, Dr. Susan Weiss

Laboratory). The wild type and mutant USB1 were cloned in pCAGGS vector with a C-terminal HA tag. Both constructs were transiently expressed in Hey1B cells

16h prior to pIC transfection for 3.5h. The FRET assay data on 2-5A concentrations showed that USB1 had no reduction effect on 2-5A levels, in contrast to AKAP7 CD. This suggested that similar to other members of the family CGI-18 and Leng9, USB1 as well does not have a catalytic effect on the activator of RNase L. These data, show that from the tested members of the LigT family, AKAP7 is the only one that when expressed in cells in degraded 2-5A.

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Figure 3.2 USB1 PDE does not degrade 2-5A. Hey1B cells were transfected with plasmid DNA followed by pIC transfection for 3.5h. (A) Western blot analysis showing expression of the transfected plasmids (B) 2-5A standard curve for FRET assay (C) 2-A concentrations as determined by FRET assay.

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AKAP7 and its Central Domain have a minimal effect on EMCV virus titers. One of the major roles of RNase L is its antiviral activity.. Several studies have shown the effect of this endoribonuclease on EMCV virus (Li et al., 1998,

Huang, 2014). .In addition, previous data shown in Chapter II confirmed that

AKAP7 degrades 2-5A activators of RNase L. To test a possible role of AKAP7 in antiviral activity of RNase L against EMCV virus, I used 293T stable cell lines expressing AKAP7 generated by lentivirus infection and puromycin selection.

The stable cells constitutively expressed an empty vector control, murine AKAP7 or murine AKAP7 CD (Figure 2.4C). The cells were infected with EMCV virus at

0.01 MOI for 10 hours (1 viral replication cycle). And the supernatants collected were used in plaque assay on indicator L929 cells to determine the EMCV viral titers (Figure 3.3B). Each plaque represents one infectious viral particle and the dilutions were performed at a 10-fold coefficient.

As quantified by the graph in Figure 3.3, there was only 2-fold increase in

EMCV titers in cells expressing AKAP7 or its CD compared to cells expressing an empty vector control. 293T cells express low levels of OAS and as a result less 2-5A is being produced (data not shown). Hence, I transiently transfected the full length murine AKAP7, rotavirus VP3-CTD and MHV ns2 in Hey1B cells

(Figure 3.4C), which express higher levels of 2-5A compared to 293T cells. After

24h, the cells were infected with 0.01 MOI EMCV (Hoskins, Sanders 1957). The supernatants were collected at 10h post infection and used in a plaque assay on

L929 cells (Figure 3.4A and B). The titers of EMCV increased by 4 fold in cells expressing AKAP7, 3 fold in cells expressing VP3-CTDand only 2 fold in cells

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expressing ns2, as compared to vector control. These results in Hey1B cells were similar to those in 293T cells suggesting that the expression of AKAP7 has minimal effect on antiviral activity of RNase L against EMCV in both of these cell lines.

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Figure 3.3 AKAP7 and AKAP7 CD have minimal effect on EMCV viral titers

293T stable cell lines expressing full length and CD AKAP7 proteins were infected for10h with EMCV virus. Viral titers were determined by plaque assays on L929 indicator cells.

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Figure 3.4 AKAP7, ns2 and VP3-CTD have minimal effect on EMCV viral titers

Hey1B cells were transfected with plasmid DNA followed by EMCV infection for 10h.

(A) Plaque assay of cellular supernatants in L929 cells (B) Quantification of plaque assay, 2’,5’-PDEs have only minimal effects on EMCV titers (C) Protein expression of the transfected plasmids (ns2 not detected, FLAG folds into the ns2)

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Depletion of AKAP7 and PDE12 expression causes increase in 2-5A levels. Our previous studies had only described the effect of AKAP7 protein overexpression in cells. To test the PDE activity of the endogenous AKAP7 protein, I generated stable cell lines of human alveolar adenocarcinoma cell line

A549 using short hairpin (sh) RNA that targets human AKAP7 and for comparison a PDE12 shRNA. The stable cells were selected by puromycin treatment and the protein expression levels were tested by western blotting

(Figure 3.5A and B). A FRET assay was used to measure 2-5A production mediated by pIC transfection for 3.5 h. As predicted, the decrease of AKAP7 and

PDE12 expression caused higher 2-5A to accumulate in the cells upon pIC

(about two-fold) l. In addition, rRNA cleavage is a signature of RNase L activation

(Wreschner et al., 1981, Silverman et al., 1983). To test the integrity of the rRNA

3.5h after activation of OAS by synthetic dsRNA, total RNA was isolated and analyzed in an Agilent Bioanalyzer. Intact 18S and 28S rRNA were reduced in

AKAP7 and PDE12 depleted cells compared to vector control. Taken together these data confirm our previous studies that AKAP7 is a cell encoded 2’,5’ PDE which degrades 2-5A thus controlling RNase L activity.

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Figure 3.5 Depletion of AKAP7 and PDE12 increases 2-5 accumulation and RNase L activity. Stable Hey1B cells were transfected with pIC for 3.5h. shRNA depleted protein expression levels arre shown by western blot analysis (A) an (B). (C) rRNA cleavage assay after RNase L activation. (D) 2-5 accumulation in Hey1B cells measured by FRET assays.

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3.4 Discussion

Candidate cellular PDEs fail to degrade 2-5A in vitro

The viral/eukaryotic LigT of 2H phosphoesterase superfamily is typified by

CGI-18 and the members of this family consist of two conserved H-Φ-[ST]-Φ

(where Φ is a hydrophobic residue) separated by an average of 80 residues

(Mazumder 2002). Viral members such as ns2 and VP3-CTD as well as cellular

AKAP7 had a catalytic effect on 2-5A. As predicted by structural homology,

(described by Mazumder 2002, Morelly 2015, Ogden 2015 and Brandmann

2015) other cellular members such as CGI18, Leng9 and USB1 could also be

2’,5 ’PDEs that degrade 2-5A. However, transient expression of these proteins failed to reduce 2-5A levels, in contrast to AKAP7 full length or its central domain.

The statistical analyses suggest that the effect of CGI-18 and Leng9 on 2-

5A levels is not significant (Figure 3.1). Interestingly, expression of USB1 and its mutant in Hey1B cells had higher levels of 2-5A compared to the control, as measured by FRET assay (Figure 3.2). It is possible that this is an effect of cell death in control cells, caused by high levels of 2-5A production after pIC transfection.

To our knowledge, this is the first evidence that CGI-18, Leng9 and USB1 do not affect 2-5A levels in vitro. Further studies are needed to determine the functions of these PDEs in cells. 2-5A is an essential component of the

OAS/RNase L pathway; hence 2-5A degrading enzymes are also critical controlling factors in the OAS/RNase L pathway. Active RNase L inhibits virus

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replication but also causes cellular RNA degradation eventually leading to cell damage. As a result, identification of cellular 2’,5’-PDEs is important for recovery of the infected cells while minimizing tissue injury. Our findings raise the possibility that CGI-18, Leng9 and USB1 do not use 2-5A as a substrate. Instead they may be specific to cyclic 2’,3’ substrates or others. It still remains an open question whether the host encodes additional PDE proteins that degrade 2-5A besides AKAP7, PDE12 and ENPP1 (Wood et al., 2015, Kubota et al., 2004b).

AKAP7 has minimal effects on EMCV titers

Several studies have already established the fact that RNase L effects

EMCV replication by cleaving viral genome ( Li 1998, Martinand 1999, Zhou

1998, Huang 2014). Based on the degradation effect that AKAP7 PDE has on 2-

5A, this protein could have an effect on EMCV titers as well. Our experiments on

EMCV replication in the presence of AKAP7 full length or its CD showed only 2- fold increase compared to vector control (Figure 3.3). Similarly, when expressed in Hey1B cells AKAP7 increased EMCV titers by 4-fold, while VP3-CTDand ns2 increased by 3 and 2 fold respectively (Figure 3.4). This difference between cell lines can be due to the difference of OAS expression levels. Our data suggest that AKAP7 has a minimal effect of EMCV possibly, due to the fact that EMCV replicates in the cytoplasm while AKAP7 is mostly nuclear. Similarly AKAP7 CD, localized to cytoplasm (MHV replication site) but not nuclear full length protein rescued MHV replication in BMMs (Figure 2.5). Hence AKAP7 may have a greater effect on viruses that replicate in the nucleus such as influenza A virus

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(see Future Perspective, Chapter IV). Alternatively, RNase L may not be activated by EMCV infection in Hey1b cells, at least not at the low moi (0.01) used in these experiments. Accordingly, rRNA cleavage of EMCV infected cells showed no RNase L specific cleavage products in EMCV infected Hey1b cells, suggesting that EMCV may not activate RNase L in these cells under these conditions (data not shown). In HeLa M cells (which lack RNase L) expression of wild type RNase L was necessary to reduced EMCV titers by around 10-fold

(Huang at el 2014). Our data suggest that AKAP7 might have a minimal proviral effect. Alternatively it may rather have a protective role in the host cell by degrading 2-5A after virus clearance to inhibit cellular RNA degradation by

RNase L.

Depletion of AKAP7 and PDE12 increased accumulation of 2-5A

PDE12, belonging to the exonuclease-endonuclease phosphatase family, was one of the first identified PDEs shown to degrade 2-5A into ATP and AMP

(Kubota 2004). AKAP7, a LigT 2’,5’-PDE, is mostly nuclear and aids in tethering

PKA. In contrast, PDE12 localizes in mitochondria and is involved in degrading mitochondrial mRNA polyA 3’-tails (Rorbach et al., 2011). AKAP7 is structurally homologous to viral antagonists of OAS/RNase L, VP3-CTD and ns2. Since previously AKAP7 was only tested by overexpression studies, shRNA knockdown stable cells were generated for both AKAP7 and PDE12. As predicted, reduction of either AKAP7 or PDE12 caused an increase in 2-5A levels produced in response to synthetic dsRNA transfection. Accordingly, there

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was an increase in rRNA cleavage as a result of active RNase L. This suggested that one of the cellular functions of AKAP7 is 2-5A degradation. We observed similar effect for both AKAP7 and PDE12. Each of these PDEs has a different intracellular localization so possibly they have localization dependent functions.

There are no studies that have detected 2-5A in mitochondria, although p46

OAS1 was observed in mitochondria. (Kjaer et al.) However 2-5A has been previously reported to be found in nuclei (Nilsen et al., 1982). Due to its small size 2-5A is predicted to transit through nuclear pores.

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CHAPTER IV

FUTURE PROSPECTIVES

The results obtained from my doctoral research identified a novel role for

AKAP7. Its PDE domain degrades 2-5A, the only known natural activator of

RNase L. This endoribonuclease when activated is involved in many roles

(reviewed in Section 1.2.2.2), and as a consequence the function(s) of AKAP7 in antagonizing OAS/RNase L still remains an open question.

To address this question, I recently generated AKAP7 knockout cell lines.

The CRISPR-Cas 9 system was used for generating human alveolar adenocarcinoma A549 cell lines lacking AKAP7 expression. To identify a small guide (sg) RNAs that targets human AKAP7 DNA, I used a sequence from the

Landers database (Wang et al., 2013). The sgAKAP7_8 sequence

TGAGCGACTGGCCAAAGCAA (targeting exon 1) was inserted in pLentiCRISPR v2 (Wang et al., 2014, Shalem et al., 2014). 293T cells were used to generate lentivirus containing an empty vector or the sgRNA targeting human AKAP7. A549 cells were infected for 2 days with lentivirus from 293T 114

cells that survived puromycin treatment were split into single cell/well and grown separately. Lack of AKAP7 expression in knockout cells was tested by western blot (data not shown)

These cells will be used in future studies to test the effect of AKAP7 on antiviral activity of RNase L. Influenza A virus replicates in nucleus, where

AKAP7 also resides (Figure 2.4D). NS1 protein of this virus is an RNase L antagonist, by sequestering dsRNA from OAS, but an ns1ΔPR8 strain will be used to infect the AKAP7 knockout cells and compare viral titers to control A549 cells. Furthermore, additional viruses which are targeted by RNase L can be used in similar experiments (e.g. West Nile virus and Sindbis virus). Since RNase

L also cleaves cellular RNA, it is possible that AKAP7 might have a protective role in the cell by eliminating 2-5A after viral clearance. AKAP7 knockout cells can be used in experiments where cell death is induced by viral infection or synthetic dsRNA. To determine if 2-5A turnover by AKAP7 protects the cell, a comparison of apoptosis markers and cell survival assays will be made between the AKAP7 knockout and control cells, Overall these experiments will determine whether the antiviral function of AKAP7 is related to cell survival. Moreover, these cells can be used in other additional experiments to characterize AKAP7 function, such as regulating mRNA stability (Brennan-Laun et al., 2014).

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Hence, AKAP7 deficient cells will be a good tool that can be used to study the function of 2-5A degradation by AKAP7. Furthermore this may lead to identification of AKAP7 inhibitors, which might have an effect on activating the immune response against viruses or producing an anti-tumor effect.

Overall, AKAP7 could be a regulator of any of the processes that RNase L or products of OAS control. For example, AKAP7 might have roles in antiviral activity and cell survival (as described above). AKAP7 might also be involved in other physiological processes affected by RNase L, such as regulation of autophagy (Chakrabarti et al., 2012) . Alternatively, it may regulate IFN production through RNase L generated RNA cleavage products, indirectly modulating innate immunity (Malathi et al., 2010, Malathi et al., 2007).

In addition, it is possible that AKAP7 is a PDE for alternative forms of 2-5A than those studied here. Interestingly, two studies have shown the presence of alternate forms of 2-5A during viral infections (Hersh et al., 1984, Rice et al.,

1985). These alternative forms of 2-5A were separated by HPLC from extracts of

IFN treated cells infected with Simian Virus 40 or vaccinia virus, but were absent in pIC treated cells. The alternative forms of 2-5A are also products of OAS and could consist of 2’-5’ linked oligoadenylates linked to several different acceptor substrates such as tRNA, A5’ppp5’’A; A5’pppp5’’A, ADP-ribose or NAD+ (Justesen et al., 2000; Ferbus et al., 1981). The function of these alternative 2-5As in cells is still not known, but they do not activate the endoribonuclease activity of RNase

L (Hersh et al., 1984, Rice et al., 1985). These alternative 2-5As may rather have

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other functions in the cell. Due to its 2’,5’-PDE activity, AKAP7 could possibly be involved in recycling of these alternate 2-5A molecules and affecting their functions in the cell.

As a conclusion, AKAP7 is a 2’,5’-phosphodiesterase that is predicted to degrade

2’,5’-linked oligoadenylates involved in a variety of cellular processes; implicating

AKAP7 as a strong candidate protein that could be therapeutically targeted.

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CHAPTER V

CONCLUSIONS

5.1 Overall summary and conclusions

The data obtained from my doctoral thesis studies shows that AKAP7 is a cellular 2’,5’-PDE and is a component of the antiviral OAS/RNase L pathway.

This system is IFN induced and its activity is controlled by viral dsRNA induced production of 2-5A molecules through OAS, but also by the enzymes that degrade 2-5A. AKAP7 cleaved 2-5A into ATP and AMP at a comparable rate with the viral homologous proteins MHV ns2 and rotavirus VP3-CTD. This suggested that AKAP7 can antagonize OAS/RNase L. Indeed AKAP7 CD successfully restored the function of a mutant ns2 gene in MHV and inhibited RNase L activation by degrading 2-5A. Furthermore 2’,5’-PDE activity was also confirmed by studies in cells transfected by synthetic dsRNA to induce OAS activity. An increased expression of AKAP7 reduced the levels of 2-5A while depleting the

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endogenous protein caused higher levels of 2-5A to accumulate in cells. In addition, we also identified a mostly nuclear localization of the full length protein when expressed in stable cell lines. In an attempt to identify additional candidate cellular 2’,5’-PDEs, that may localize in the cytoplasm, we also tested other members of the viral/eukaryotic LigT family of proteins, CGI-18, Leng9 and

USB1, but these failed to degrade 2-5A when expressed in cells. This study suggests that AKAP7 may be the only eukaryotic LigT member that cleaves the phosphodiesterase bonds of the 2-5A molecules.

Overall, we propose a novel role for AKAP7. It is the first member of the

AKAP family to act as a PDE instead of binding one. AKAP7 degrades 2-5A the mediator of the innate immunity OAS/RNase L pathway, similar to its viral homologous PDEs. This suggests an evolutionary process in which a cellular gene was acquired by viruses to antagonize the host innate immune system.

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