Host factors involved in RNA replication of Dianthovirus( Title Dissertation_全文 )

Author(s) Hyodo, Kiwamu

Citation 京都大学

Issue Date 2014-03-24

URL https://doi.org/10.14989/doctor.k18333

Right 許諾条件により本文は2015-03-24に公開

Type Thesis or Dissertation

Textversion ETD

Kyoto University

Host factors involved in RNA replication of Dianthovirus

Kiwamu Hyodo

2014

Contents

General Introduction 1

ChapterI 6 Identification of amino acids in auxiliary replicase protein p27 critical for its RNA-binding activity and the assembly of the replicase complex in Red clover necrotic mosaic virus

ChapterII 34 ADP Ribosylation Factor 1 Plays an Essential Role in the Replication of a Plant RNA Virus

ChapterIII 69 Functional analysis of phospholipase D and phosphatidic acid in a plant RNA virus replication

References 91

Summary 107

Acknowledgements 110

General Introduction

Viruses are obligate intracellular symbionts. Viral genomes consist of a single or double-stranded DNA or RNA encoding genetic information required for viral entry, replication and spread. Viruses are now thought to have been involved in evolution of organisms including animals and plants by symbiotic association (Roossinck 2005). Meanwhile, viruses have also been recognized as parasites that invade cells and hijack cellular machinery for their own purposes. In fact, viruses cause a large numbers of human diseases, including hepatic inflammation, immune deficiency syndrome, cancer, and so on. In addition, viruses are pathogenic agents that devastate cereals, vegetable and live stocks, which are essential to the viability of human-beings. As the world population continues to increase, more foods are needed for sustaining the population. To manage the food crisis, it is important to control viral infection in plants and live stocks. However, there is no established direct method to control viral diseases, in part because we do not fully understand how virus replicates in cells of plants and animals. Positive-strand RNA [(+)RNA] viruses are the most abundant in plant viruses, and include many economically important viruses in agriculture. Following entry into host cells, viral genomic RNAs are released from virions into the cytoplasm of the host cells and act as templates for translation to produce replication proteins using the host’s translation machinery (Dreher and Miller 2006; Simon and Miller 2013). The replication proteins recruit genomic RNA together with host proteins to form the viral replication complex (VRC). The VRC synthesizes a complementary negative-strand

RNA [(–)RNA] using the original genomic RNA as a template. The (–)RNA is then used as a template to synthesize many new (+)RNAs that undergo additional rounds of translation and replication, or move to adjacent cells, or are encapsidated into virions (Buck 1996; Nagy and Pogany 2012). Although (+)RNA viruses have a limited coding capacity and code for only several genes, RNA viruses perform many tasks (e.g., synthesis of viral proteins and RNAs,

1 regulation of RNA replication and gene expression, escape from antiviral responses, and cell-to-cell and long distance movement). To accomplish these tasks, viruses seem to have acquired the ability to use host-derived proteins, membranes, lipids, and metabolites, and to exploit or rewire cellular trafficking pathways during evolution. All characterized eukaryotic (+)RNA viruses assemble VRCs, which contain both viral and host proteins, on intracellular membranes (Ahlquist et al. 2003; den Boon et al. 2010; den Boon and Ahlquist 2010; Laliberté and Sanfaçon 2010; Miller and Krijnse-Locker 2008; Nagy and Pogany 2012). These membranes can be derived from various organelles such as the endoplasmic reticulum (ER), Golgi, mitochondria, chloroplasts, peroxisomes, vacuoles, as well as the plasma membranes. RNA replication mechanisms of positive-stranded RNA viruses have been extensively studied. However, many questions remain unanswered, for example: how do viral replication proteins recognize specifically viral genomic RNAs from a pool of host RNAs (mRNA, tRNA and rRNA), how do viruses establish VRCs, what host factors are involved in viral multiplication? To understand details of the replication mechanisms of positive-strand RNA viruses, I use Red clover necrotic mosaic virus (RCNMV) as a model virus to investigate RNA replication mechanisms. RCNMV has several interesting features not observed in other positive-strand RNA viruses (Okuno and Hiruki, 2013). RCNMV is a member of the family Tombusviridae and the genus Dianthovirus that includes Carnation ring spot virus (CRSV) as a type member, and Sweet clover necrotic mosaic virus (SCNMV) (Okuno and Hiruki, 2013). Dianthoviruses are taxonomically distinct from other viruses in Tombusviridae (e.g., TBSV) because of the bipartite nature of their single-stranded positive sense RNA genome. Virions of RCNMV are about 35 nm in size and composed of 180 copies of a 37 kDa coat protein (CP) (Okuno and Hiruki, 2013). The two RNA genomes RNA1 (3.9kb) and RNA2 (1.45kb) (Gould et al., 1981; Hiruki, 1987; Okuno et al., 1983), lack both cap structure at the 5! end and poly(A) tail at the 3! end (Lommel et al., 1988; Mizumoto et al., 2003). RNA1 and

2 RNA2 share fundamentally no homology, except for the first seven nucleotides at the 5! ends and two stem-loop structures at the 3! ends in both genomic RNAs. RNA1 encodes RNA replicase components, a 27-kDa protein (p27) and its N-terminally overlapping 88-kDa RNA-dependent RNA polymerase (RdRP) (p88pol). p88pol produced by -1 frameshifting (Kim and Lommel 1994, 1998). Both p27 and p88pol are required for replication of RNA1 and RNA2 in plants or protoplasts (Takeda et al., 2005; Okamoto et al., 2008; Mine et al., 2010b). p27 and p88pol form 480 kDa RNA replication complex in RCNMV-infected plants (Mine et al., 2010b). This 480 kDa complex retained RdRp activity in vitro (Mine et al., 2010b). RNA1 also encodes a 37-kDa coat protein (CP) that is expressed from subgenomic RNA (CP sgRNA) (Tatsuta et al., 2005; Zavriev et al., 1996). Transcription of CP sgRNA requires intermolecular interaction between RNA1 and RNA2 (Sit et al., 1998; Tatsuta et al., 2005). RNA1 alone can replicate in a single cell, but can not move to neighboring cells without 35-kDa movement protein (MP) encoded by RNA2 (Lommel et al., 1988; Xiong et al., 1993; Kaido et al., 2009). Mechanisms of translation and RNA replication differ between RNA1 and RNA2. Cap-independent translation of RNA1 is mainly mediated by the translation-enhancer element of dianthovirus RNA1 (3!TE-DR1) that resides between 3596 nt and 3732 nt in the 3!-untranslated region (UTR) (Mizumoto et al., 2003). On the other hand, RNA2 does not have such translational enhancer elements as 3!-TE-DR1. Cap-independent translation of RNA2 is coupled with RNA replication, and, therefore, cis-acting RNA elements required for RNA replication are also essential for translation of RNA2 (Mizumoto et al., 2006). Replication mechanisms are also different between RNA1 and RNA2. Replication proteins were required in cis for the replication of RNA1, whereas RNA2 can utilize replication proteins supplied in trans for its replication (Takeda et al., 2005; Okamoto et al., 2008). RCNMV RNA replication is also required for counter-defence against RNA interference (RNAi), which is induced by double-strand RNA (dsRNA) and is involved in antiviral defense (Takeda et al., 2005). Host factors involved in (+)RNA plant virus replication have been identified using

3 several approaches. Genome-wide screening of Saccharomyces cerevisiae led to the identification of up to 130 genes that affect the replication of Tomato bushy stunt virus (TBSV) (Jiang et al. 2006; Panavas et al. 2005) or Brome mosaic virus (BMV) (Gancarz et al. 2011; Kushner et al. 2003) and up to 30 genes that affect TBSV RNA recombination (Serviene et al. 2005, 2006). Proteome-wide overexpression screening in S. cerevisiae also identified numerous host proteins that affect TBSV replication (Shah Nawaz-ul-Rehman et al. 2012). Mass spectrometric analysis of cellular proteins coimmunopurified with viral replication proteins has identified the host proteins that are part of the VRC or that interact with the viral replication proteins in several viruses, including Tomato mosaic virus (ToMV) (Nishikiori et al. 2006), TBSV (Serva and Nagy 2006), Potyviruses (Dufresne et al. 2008; Hafrén et al. 2010), and RCNMV (Mine et al. 2010b). Mass spectrometric analysis has also been used to identify host proteins that interact with the viral RNA of RCNMV (Iwakawa et al. 2012), Bamboo mosaic virus (BaMV) (Huang et al. 2012; Lin et al. 2007; Prasanth et al. 2011), and ToMV (Fujisaki and Ishikawa 2008). Protein microarray analysis led to the identification of host proteins that interact directly with the replication proteins of TBSV or with the viral RNAs of TBSV and BMV (Li et al. 2008, 2009; Zhu et al. 2007). Microarray analysis of host gene expression during viral infection and subsequent functional analysis has also been used to identify host factors involved in virus replication (Chen et al. 2013a, b). However, the functions of most host factors that have been identified using such approaches remain unknown.
To reveal the RNA replication mechanisms, I investigated the functions of p27 replication protein. Using a viral translation/replication system containing cytoplasmic extracts prepared from tobacco BY-2 cells (BYL) (Komoda et al., 2004), which is a powerful tool to investigate individual determinants of the replication process of positive-strand RNA viruses, I revealed that p27 has multiple functions during RNA replication (Chapter I). Moreover, I identified the host proteins that interact with RCNMV replication proteins and revealed the importance of host membrane trafficking

4 pathway (Chapter II) and specific phospholipid (Chapter III) in viral RNA replication. Understanding the roles of host proteins as well as viral factors in viral replication provides information about the molecular pathways exploited by the virus and further targets that could be pursued in the development of antiviral strategies against existing and emerging plant virus diseases.

5 Chapter I Identification of amino acids in auxiliary replicase protein p27 critical for its RNA-binding activity and the assembly of the replicase complex in Red clover necrotic mosaic virus

Introduction Positive-strand RNA viruses are replicated by the viral replicase complex on intracellular membranes (Ahlquist, 2006). The replicase complex consists of viral replication templates, viral replicase proteins, such as RNA-dependent RNA polymerase (RdRP) and auxiliary proteins, and host-derived proteins (Ahlquist et al., 2003). In host cells, viral replicase proteins selectively recruit viral replication templates and other components of the replicase complex into intracellular membranes for the assembly of the viral replicase complex (Ahlquist et al., 2003; Nagy and Pogany, 2008). For example, the auxiliary replicase protein p33 of the tombusviruses (Tomato bushy stunt virus [TBSV] and Cucumber necrosis virus) recruits a replicon RNA and p92 RdRP into the peroxisomal membranes in yeast, a model host for the study of virus replication (Panavas et al., 2005). p33 binds directly to an internal replication element present in the replicon RNA (Pogany et al., 2005) and interacts with p92 via a protein"protein interaction (Rajendran and Nagy, 2004 and 2006). Both these interactions are required for the assembly of the Tombusvirus replicase complex (Panaviene et al., 2004 and 2005; Pogany et al., 2005; Rajendran and Nagy, 2006). In Brome mosaic virus (BMV), the 1a auxiliary protein recruits replication templates and 2a RdRP to the endoplasmic reticulum (ER) membrane, which is the site of replication in BMV (Liu et al., 2009; Schwartz et al., 2002 and 2004; Wang et al., 2005). The specific recognition of the replication templates by 1a depends on 1a-responsive elements, which are present at the 5! end of RNA2 and in the intergenic region of RNA3 (Chen et al., 2001; Schwartz et al., 2002; Sullivan and Ahlquist, 1999). 1a, 2a, and the viral nucleotide sequences of RNA3,

6 including the 1aresponsive element, are required for the assembly of the functional BMV replicase complex in yeast (Quadt et al., 1995). All these data imply that the viral auxiliary protein is a key player in the recognition of viral replication templates and the assembly of the viral replicase complex. Despite intensive studies of several viral auxiliary proteins, their functions in viral RNA replication are not fully understood. Red clover necrotic mosaic virus (RCNMV) is a positive-strand RNA plant virus and a member of the genus Dianthovirus in the family Tombusviridae. Dianthovirus is taxonomically distinct from the other viruses of Tombusviridae because of the bipartite nature of its genome. RCNMV genomic RNA1 and RNA2 have neither cap structures at their 5! ends nor poly (A) tails at their 3! ends (Lommel et al., 1988; Mizumoto et al., 2003). Instead, the 3! untranslated region (3! UTR) of RNA1 contains a cis-acting RNA element that is responsible for capindependent translation (Mizumoto et al., 2003; Sarawaneeyaruk et al., 2009). Although RNA2 has no such RNA element, efficient capindependent translation occurs after the replication of RNA2 (Mizumoto et al., 2006). RNA1 encodes two replicase component proteins, a 27-kDa auxiliary protein (p27) and its N-terminally overlapping "1 frameshifted product, an 88-kDa RdRP (p88) (Kim and Lommel, 1994 and 1998; Xiong et al., 1993b). RNA1 also encodes a coat protein, which is translated from the subgenomic RNA (sgRNA) (Xiong and Lommel, 1989). The transcription of sgRNA requires an intermolecular interaction between RNA1 and RNA2 (Sit et al., 1998; Tatsuta et al., 2005). RNA2 encodes a movement protein that is required for viral movement in plants (Kaido et al., 2009; Xiong et al., 1993a). p27, together with p88 and some host proteins, forms the 480-kDa replicase complex of RCNMV, which is a key player in the replication of both RNA1 and RNA2 (Mine et al., 2010a and b). p27 binds to the Y-shaped RNA element (YRE) located in the 3! UTR of RNA2 via a direct RNA"protein interaction and to RNA1 in a translation-coupled manner (Iwakawa et al., 2011). The former interaction is essential for the membrane association and subsequent negative-strand synthesis of RNA2 (An et

7 al., 2010; Iwakawa et al., 2011). The latter interaction, in combination with the translation-coupled interaction of p88 with RNA1, is important for the synthesis of negative-strand RNA1 (Iwakawa et al., 2011; Okamoto et al., 2008). Both p27 and p88 are localized to the ER membranes (Turner et al., 2004). These findings suggest that the auxiliary protein p27 plays important roles in the early replication processes of RCNMV, including the recruitment of the replication templates to the ER membranes, the formation of the 480-kDa replicase complex, and negative-strand RNA synthesis. However, the RNA-binding domain(s) of p27 have not been investigated and the functions of p27 in RCNMV infection are not fully understood. In this study, to characterize the functions of p27 and its role in the early replication processes of RCNMV, I first identified the regions and critical amino acid residues in p27 that are involved in its specific YRE-mediated interaction with RNA2, using an aptamer pull-down assay. I then analyzed the p27-mediated membrane association of RNA2 and the formation of the 480-kDa replicase complex using in vitro fractionation and blue-native polyacrylamide gel electrophoresis (BN-PAGE). I found that the recruitment of RNA2 to the membrane fraction depends on the binding of p27 to YRE. I also showed that the binding of p27 to RNA and to the replicase proteins, including itself, are insufficient but necessary to facilitate the formation of the functional 480-kDa replicase complex, which is essential for the negative-strand synthesis of RNA2. In this study, I also identified the amino acids whose mutations abolished the formation of the 480-kDa replicase complex, but did not affect the binding of p27 to the viral RNA or replicase proteins, suggesting an additional role for p27 in the assembly of the replicase complex. These results indicate that p27 plays multiple roles during the early replication processes of RCNMV.

8 Results

Mapping the region(s) in p27 involved in its specific binding to YRE To determine the domains in p27 responsible for its specific interaction with YRE of RNA2, Iconstructed a series of deletion mutants of C-terminally FLAG-tagged p27 (p27-FLAG; Fig. I-1A) and tested their binding activities to YRE using modified StreptoTag (STagT) affinity purification methods. STagT affinity purification has been used successfully to analyze RNA–protein interactions (Dangerfield et al., 2006), including the interaction between p27 and YRE (Iwakawa et al., 2011). In this assay, Iused a cell-free in vitro translation/replication system prepared from evacuolated tobacco BY-2 protoplasts (BYL; Komoda et al., 2004). The STagT-fused YRE was incubated in the supernatant fractions produced by the centrifugation at 21,000g of BYL (BYLS20) expressing wild-type or truncated p27-FLAG proteins, and the STagT-fused RNA–protein complexes were affinity purified with streptomycin-conjugated beads (Iwakawa et al., 2011). The purified RNA and copurified p27-FLAG proteins were analyzed with ethidium bromide staining and immunoblotting, using an anti-FLAG antibody, respectively. The STagT-fused YRE was purified efficiently in all incubations (Fig. I-1B, bottom panels). Wildtype p27-FLAG protein, but not RLuc-FLAG protein, was copurified with YRE in the affinity-purified fraction (Fig. I-1B, lanes 1 and 2, middle panels), confirming the specific interaction of p27 with YRE (Iwakawa et al., 2011). p27-FLAG#21–40, p27-FLAG#114–136, and p27-FLAG#137–156 were negligibly or not detected before affinity purification (Fig. I-1B, lanes 4, 11, and 12, top panels), suggesting the importance of these deleted amino acids for the stability of p27. p27-FLAG#41–60, p27-FLAG#81–100, p27-FLAG#157–176, and p27-FLAG#177– 196 were negligibly or not detected in the affinity-purified fractions (Fig. I-1B, lanes 5, 7, 13, and 14, middle panels). The remaining deletion derivatives of p27-FLAG were copurified with YRE as efficiently as was wild-type p27-FLAG (Fig. I-1B, lanes 3, 6, 8, and 15, middle panels). These results demonstrate that the N-terminal amino acids 21–

9 60 and 81–100 and the C-terminal amino acids 157–196 are required for the p27"YRE interaction.

Identification of the amino acid residues in p27 critical for the p27–YRE interaction To identify the amino acid residues within the N- and C-terminal regions of p27 that are critical for the p27"YRE interaction (Fig. I-1), I generated alanine-scanning mutants of p27-FLAG. The introduced mutations included single amino acid substitutions: lysine to alanine (K25A, K43A, and K86A); arginine to alanine (R42A and R164A); histidine to alanine (H53A, H179A, and H180A); glutamic acid to alanine (E178A); and a double amino acid substitution (HH179–180AA). Iselected these amino acids because positively charged amino acids are expected to be exposed on the protein surface and therefore to be involved in interactions with other macromolecules, such as RNA and proteins (Auweter et al., 2006; Draper, 1999; Panaviene et al., 2003; Rajendran and Nagy, 2003), and because these amino acids are conserved among the dianthoviruses (Fig. I-2A). The p27-FLAG mutants with these mutations were tested for the p27"YRE interaction using STagT affinity purification, as described above. p27-FLAG/R164A, p27-FLAG/H179A, p27-FLAG/H180A, and p27-FLAG/HH179–180AA were negligibly detected in the affinity-purified fractions (Fig. I-2B, lanes 10 and 12–14, middle panels), whereas the remaining mutants were copurified with YRE as efficiently as was wild-type p27 (Fig. I-2B, lanes 3–7, and 11, middle panels). These results indicate that the arginine residue at position 164 (R164) and the histidine residues at positions 179 and 180 (H179 and H180) are important for the p27"YRE interaction.

Binding activity of p27 to YRE is required for recruiting RNA2 to the membranes Ipreviously reported that the p27"YRE interaction is required for the membrane association of RNA2 (Iwakawa et al., 2011). To test the effects of the introduced mutations described above on the recruitment of RNA2 to the membranes, I performed

10 a fractionation assay using BYL. BYL expressing p27-FLAG or its derivatives in the presence of RNA2 were centrifuged at 21,000g to yield a membrane-containing pelleted fraction and a supernatant fraction. The proteins and RNAs extracted from the total, pelleted, and supernatant fractions were analyzed with immuno- and northern blotting, respectively. Sec61, an integral ER membrane protein, was detected in the pelleted fraction but not in the supernatant fraction (data not shown), confirming that the pelleted fraction contained the ER. All the p27-FLAG mutants accumulated to similar levels in the membrane fraction (Fig. I-3A). When the p27-FLAG proteins with mutations that I previously reported that the p27"YRE interaction is required for the membrane association of RNA2 (Iwakawa et al., 2011). To test the effects of the introduced mutations described above on the recruitment of RNA2 to the membranes, I performed a fractionation assay using BYL. BYL expressing p27-FLAG or its derivatives in the presence of RNA2 were centrifuged at 21,000g to yield a membrane containing pelleted fraction and a supernatant fraction. The proteins and RNAs extracted from the total, pelleted, and supernatant fractions were analyzed with immuno- and northern blotting, respectively. Sec61, an integral ER membrane protein, was detected in the pelleted fraction but not in the supernatant fraction (data not shown), confirming that the pelleted fraction contained the ER. All the p27-FLAG mutants accumulated to similar levels in the membrane fraction (Fig. I-3A). When the p27-FLAG proteins with mutations that compromised their interactions with YRE (R164A, H179A, H180A, and HH179–180AA; Fig. 3B) were tested, the levels of RNA2 accumulated in the membrane fractions were greatly reduced whereas the remaining p27-FLAG proteins with mutations that did not affect the p27"YRE interaction supported the recruitment of RNA2 to the membrane fraction at levels similar to that recruited by wild-type p27 (Fig. I-3B). The correlation between the levels of RNA2 accumulated in the membrane fraction and the binding activity of the p27-FLAG mutants to YRE (Figs. I-2B and I-3B) indicates that the binding of p27 to YRE plays an essential role in the recruitment of RNA2 to the membrane fraction and

11 confirms the importance of the p27"YRE interaction in RNA2 recruitment to the membranes (Iwakawa et al., 2011).

Roles of the binding activity of p27 to YRE in negative-strand RNA synthesis To investigate whether there is a functional relationship between the binding of p27 to YRE and negative-strand RNA synthesis, the p27-FLAG mutants were incubated with RNA2 together with RNA1-p88, which expresses p88 alone, in BYL. The total RNAs and proteins were extracted after incubation for 4 h. Northern blot analysis showed that the p27-FLAG proteins with mutations that compromised the p27"YRE interaction abolished or greatly reduced the accumulation of negative-strand RNA2 (Fig. I-4A, lanes 8 and 10–12). All p27 proteins accumulated to similar levels in BYL (Fig. I-4A). Similar results were obtained in BY-2 protoplasts, in which p88 was expressed from a plasmid that encodes an mRNA containing the open reading frame of p88 (Fig. I-4B). These results suggest that the binding of p27 to YRE is important for the negative-strand synthesis of RNA2. Interestingly, however, p27-FLAG/R42A, p27-FLAG/K86A, and p27-FLAG/E178A, which can interact with YRE, dramatically reduced the accumulated levels of negative-strand RNA2 in BYL (Fig. I-4A, lanes 4, 7, and 9), as well as the levels of positive-strand RNA2 in BY-2 protoplasts (Fig. I-4B, lanes 4, 7, and 9). These mutants were not copurified with YRE-M that possesses a substitution mutation in the loop sequence of SL8 in YRE (refer to R2-3!-84M-S in Iwakawa et al., 2011; data not shown). These results indicate that these mutations impair the function(s) of p27 required for the negative-strand synthesis of RNA2 other than its binding to YRE and the recruitment of RNA2 to the membranes. I refer hereafter to the p27-FLAG mutants that are compromised in the p27"YRE interaction (p27-FLAG/R164A, p27-FLAG/H179A, p27-FLAG/H180A, and p27-FLAG/HH179–180AA) as “class I mutants,” and the mutants that can interact with YRE but have little or no ability to support the negative-strand RNA synthesis of RNA2 (p27-FLAG/R42A, p27-FLAG/K86A, and p27-FLAG/E178A) as “class II mutants.”

12 The remaining p27-FLAG mutants, which are competent to support both the p27"YRE interaction and the negative-strand synthesis of RNA2 (p27-FLAG/K25A, p27-FLAG/K43A, and p27-FLAG/H53A), are referred to as “class III mutants.” The mutations present in these mutants will also be referred to as class I, class II, and class III mutations, respectively. To investigate whether the mutations described above also affect the replication of RNA1, I tested the p27-FLAG mutants for their ability to cause the accumulation of negative-strand RNA1 in BYL and to support the replication of RNA1 in BY-2 protoplasts. p27-FLAG and its derivatives were incubated with RNA1-p88 in BYL or were coexpressed with RNA1-p88 from plasmids in BY-2 protoplasts. I chose this combination because only RNA1, from which p88 is translated, is an efficient template for replication in the presence of p27 (Okamoto et al., 2008). The total RNAs and proteins were extracted from BYL or BY-2 protoplasts after incubation for 4 h or 24 h, respectively. All the p27-FLAG proteins accumulated efficiently in both BYL and BY-2 protoplasts (Fig. I-5). Northern blot analysis showed that the class I and class II mutations abolished or greatly reduced the levels of negative-strand RNA1 accumulated in both BYL (Fig. I-5A, lanes 4 and 7–12) and BY-2 protoplasts (Fig. 5B, lanes 4 and 7–12). These results indicate that the amino acids mutated in the class I and II mutants are important for the functions of p27 in the synthesis of negative-strand RNA1, as well as in the synthesis of RNA2. It should be noted that the levels of RNA1 accumulated by the class I mutants correlated with their binding activities to YRE of RNA2 (Fig. I-2B), even though RNA1 has no RNA element, like YRE, that interacts directly with replicase proteins supplied in trans (Iwakawa et al., 2011).

Effects of the introduced p27 mutations on the protein–protein interactions between RCNMV replicase proteins The C-terminal half of p27, encompassing amino acid residues 124–236 (p27C), is responsible for p27–p27 and p27–p88 interactions, and these interactions are required

13 for the assembly of the 480-kDa replicase complex, which is thought to be a key player in the replication of RCNMV RNA (Mine et al., 2010a and b). The class II mutation E178A and all class I mutations occur in the C-terminal region of p27. Therefore, it is possible that these mutations affect the p27–p27 and/or p27–p88 interactions, thus compromising the assembly of the 480-kDa replicase complex. To test the effects of these mutations on the p27–p27 and p27–p88 interactions, Iused a glutathione S-transferase (GST) pull-down assay. N-terminally histidine (His)- and C-terminally FLAG-tagged p27C mutants expressed in Escherichia coli were purified (Fig. I-6A) and tested for their interactions with N-terminally His- and GST-fused p27C (His/GST-p27C) and His/GST-p88#p27 (Mine et al., 2010a). His/GST-p88#p27 is a truncated form of p88 that lacks the region that overlaps p27 and is competent to interact with p27 and to support the negative-strand synthesis of RNA2 (Mine et al., 2010a). His/GST-p27C and His/GST-p88#p27 captured on glutathione-bound beads were incubated with purified His-p27C-FLAG mutants. After extensive washing, the bound proteins were analyzed by immunoblotting with an anti-FLAG antibody. The results showed that the substitution of H179 with alanine inhibited the interaction of p27 with His/GST-p27C but not its interaction with His/GST-p88#p27 (Fig. I-6B and C, lane 10). The substitution of H180 or HH179–180 with alanine(s) inhibited the interactions of p27 with both His/GST-p27C and His/GST-p88#p27 (Fig. I-6B and C, lanes 11 and 12). These results suggest that H179 and H180 are important not only for the p27"YRE interaction but also for the p27–p27 and/or p27–p88 interactions. Interestingly, however, the class I mutant p27-FLAG/R164A and the class II mutant p27-FLAG/E178A interacted with both p27 and p88#27 (Fig. I-6B and C, lanes 8 and 9). The ability of p27-FLAG/E178A to interact with p88#27 was much greater than the interaction of wild-type p27 (Fig. I-6C, lane 9).

14 Amino acids required for the formation of the 480-kDa replicase complex on RNA2 The successful interactions of the class I mutant p27-FLAG/R164A and the class II mutant p27-FLAG/E178A with both p27 and p88#27 prompted us to investigate whether these mutants are competent to form the 480-kDa replicase complex. To investigate the accumulation of the 480-kDa replicase complex, I incubated transcripts that expressed wild-type or mutant p27-FLAG, together with RNA1-p88 and RNA2 in BYL. After incubation for 4 h, the protein samples were subjected to BN-PAGE and analyzed by immunoblotting with an anti-FLAG antibody. The accumulated levels of the 480-kDa replicase complex were below the level of detection in BYL expressing the class II mutants (Fig. I-7, lanes 4, 7, and 9) or the class I mutant p27-FLAG/HH179–180AA (Fig. I-7, lane 12). It should be noted that the class II mutations R42A and K86A are in the region that is not essential for the p27–p27 or p27–p88 interaction (Mine et al., 2010a). The class I mutants with a single mutation accumulated reduced amounts of the 480-kDa replicase complex compared with that accumulated by wild-type p27 (Fig. I-7, lanes 8, 10, and 11), although the effect of the R164A mutation was milder than those of the other single mutations. These results indicate that the class II mutations and the HH179–180AA mutation had very deleterious effects on the capacity of p27 to form the 480-kDa replicase complex, and that the other single class I mutations also affected the capacity of p27 to form the replicase complex. The class III mutants allowed the accumulation of the 480-kDa replicase complex to levels similar to those accumulated by wild-type p27-FLAG (Fig. I-7, lanes 2, 3, 5 and 6).

15 Discussion

The present study demonstrated that the auxiliary replicase protein p27 of RCNMV consists of several domains responsible for its diverse functions that are required to play multiple roles during the early replication processes of RCNMV.

RNA-binding domains of p27 The deleterious effects of deletions in both the N- and C-terminal regions of p27 on the p27–YRE interaction (Fig. I-1B, lanes 5, 7, 13, and 14) suggest the involvement of multiple domains of p27 in its RNA-binding activity. However, alanine-scanning mutations introduced into the N-terminal region of p27-FLAG had no deleterious effects on the p27"YRE interaction (Fig. I-2B, lanes 3–7) whereas those introduced into the Cterminal region, such as R164A, H179A, H180A, and HH179–180AA, abolished or greatly reduced the p27"YRE interaction. This mutagenesis analysis suggests that the C-terminal region, encompassing at least amino acids 157–196, is a major determinant of the RNA-binding property of p27. This idea is supported by an in silico analysis of the p27 protein with the program RNABindR (Terribilini et al., 2006 and 2007), which predicted that the C-terminal region of p27 is a potential RNA-binding site, but that the N-terminal region is not (data not shown). The N-terminal region of p27 might be important in supporting the conformation of p27 required for its interaction with YRE. In our previous study, I demonstrated that C-terminal amino acid residues 124–236 of p27 are essential for both the p27–p27 and the p27–p88 interactions through direct contact, and that the p27–p88 interaction occurs between the C-terminal half of p27 and the non-overlapping unique region of p88 (Mine et al., 2010a). The deletion of C-terminal amino acids 197–236, which are essential for the p27–p27 and p27–p88 interactions (data not shown), had no deleterious effect on the RNA-binding activity of p27 (Fig. I-1B, lane 15), indicating that different functional domains of p27 are required

16 for its binding to RNA and protein. However, the class I mutations (R164A, H179A, H180A, and HH179–180AA) had deleterious effects on the RNA-binding activity of p27, and these mutations occur in the region required for protein binding. These results indicate that the main functional domains required for RNA and protein binding differ, but that these domains overlap in p27.

Roles of p27 in the replication of RNA2 The class I mutants with no or reduced activity to bind RNA were also compromised in their recruitment of RNA2 to the membrane fraction and in the negative-strand synthesis of RNA2 (Figs. I-2B, I-3, and I-4). The robust correlation between these functions of p27 strongly suggests that the RNA-binding activity of p27 plays an essential role in the recruitment of RNA2 to the membranes, which are the sites of RCNMV RNA replication. The class I mutations also compromised the formation of the 480-kDa replication complex, although the effect of the R164A mutation was very mild (Fig. I-7, lanes 8 and 10–11). This mild effect might be partly explained by the fact that the R164A mutation has no effect on the interactions between the replicase protein components (Fig. I-6B and C, lane 8). The class II mutants (R42A, K86A, and E178A) retained the ability to interact with both YRE and the replicase proteins (Figs. I-2B and I-6B and C), but lost the capacity to formthe 480-kDa replicase complex (Fig. I-7, lanes 4, 7, and 9). It is possible that the class II mutations affect the interactions between p27 and the host proteins, directly or indirectly, and the subsequent assembly of the 480-kDa replicase complex. Our previous study demonstrated that the RCNMV replicase complex contains many host proteins, including ubiquitin, heat shock protein 70, heat shock protein 90, and several ribosomal proteins (Mine et al., 2010b). It should be noted that the E178A mutation increased viral protein–protein interactions (Fig. I-6B and C) with retaining the ability for specific binding to YRE (Fig. I-2; data not shown). The undesirable interactions

17 between viral replicase proteins by themselves might affect the interactions between the viral proteins and the host proteins and disturb the assembly of the 480-kDa replicase complex. The mutated amino acid residues might be involved in the posttranslational modification of p27. Ubiquitination is a common posttranslational modification in eukaryotic cells (Pickart and Eddins, 2004) and is known to play a role during the infection processes of several viruses. For example, the ubiquitination of nonstructural protein 3D RdRP is required for the effective replication of Coxsackievirus B3 (Si et al., 2008). The auxiliary replicase protein p33 of TBSV is ubiquitinated, and this ubiquitination is important for the replication of the TBSV replicon RNA (Barajas and Nagy, 2010; Li et al., 2008). The posttranslational modifications of the RCNMV replicase proteins, including the ubiquitination of lysine at position 86, are yet to be analyzed. In contrast, the class II mutations might affect other unknown functions of p27, including as an RNA chaperone. The roles of virus-encoded RNA chaperones in replication have been reported in many RNA viruses (Züniga et al., 2009). It is noteworthy that the Tombusvirus auxiliary replicase protein p33 has RNA chaperone activity (Stork et al., 2011). Thus, p27 has distinct domains that interact with YRE, viral replicase proteins, and host proteins, and all these interactions are required for the replication of RNA2.

Roles of p27 in the replication of RNA1 In our previous study, I demonstrated that RNA1 lacks a replicase recruiter, such as YRE of RNA2, and that the binding of the replicase proteins to RNA1 is coupled to the translation process (Iwakawa et al., 2011). Therefore, the initial template recognition mechanisms of the replicase proteins differ for RNA1 and RNA2. I proposed a model for the recruitment of RNA1 into the replication process by p27 and p88, wherein p27 binds directly or indirectly to RNA1 during its translation, and recruits RNA1 to the ER membrane (Iwakawa et al., 2011). p88, which is produced by a"1 frameshifting event,

18 also binds to the 3!UTR of RNA1 in a translation-coupled manner (Iwakawa et al., 2011). Considering the correlation between the specific RNA-binding activity of p27 to YRE (Iwakawa et al., 2011; this study) and its ability to support the replication of RNA1 (Fig. I-5), it is plausible that p27 binds directly to RNA1 by recognizing RNA elements that become available only during the translation of RNA1 (Iwakawa et al., 2011). This direct interaction probably contributes not only to the recruitment of RNA1 to the ER membrane but also to the assembly of the 480-kDa replicase complex on RNA1, together with p88.

Materials and methods

Plasmid construction Plasmids with the prefixes “pUC” and “pRC” were used for in vitro transcription, and plasmids with the prefixes “pUB” and “pCold” or “pColdGST” were used for the protoplast experiments and the expression of recombinant proteins, respectively. pUCR1 and pRC2|G are full-length cDNA clones of RNA1 and RNA2, respectively, of the Australian RCNMV strain (Takeda et al., 2005; Xiong and Lommel, 1991). Previously described constructs that were used in this study include the following: pUCR1p88 (Takeda et al., 2005), pUCp27-FLAG (Mine et al., 2010a), pUCRLuc– FLAG (Iwakawa et al., 2011), pUCR2–3!-84-S (Iwakawa et al., 2011), pUCR2– 3!-84M-S (Iwakawa et al., 2011), pUBp27 (Takeda et al., 2005), pUBp88 (Takeda et al., 2005), pUBR1–p88 (Okamoto et al., 2008), pCold–p27C-FLAG (Mine et al., 2010a), pColdGST–p27C (Mine et al., 2010a), and pColdGST–p88#p27 (Mine et al., 2010a). All the constructs were verified by sequencing. The primers used in this study are listed in Table 1.

19 pUCp27!FLAG"1–20; pUCp27!FLAG"21–40; pUCp27!FLAG"41–60; pUCp27!FLAG"61–80; pUCp27!FLAG"81–100; pUCp27!FLAG"101–113; pUCp27!FLAG"114–136; pUCp27!FLAG"137–156; pUCp27!FLAG"157–176; pUCp27!FLAG"177–196; pUCp27!FLAG"197–236: DNA fragments were amplified by PCR from pUCp27-FLAG. The primer pairs used consisted of pUCSacI-F plus one each of the following: #1–20-R, #21–40-R, #41–60-R, #61–80-R, #81–100-R, #101–113-R, #114–136-R, #137–156-R, #157– 176-R, #177–196-R, or #197–236-R. Another primer, 3!UTR-R, was used together with one each of the following: #1–20-F, #21–40-F, #41–60-F, #61–80-F, #81–100-F, #101–113-F, #114–136-F, #137–156-F, #157–176-F, #177–196-F, or #197–236-F. Each recombinant PCR fragment was amplified with the primer pair pUCSacI-F and 3!UTR-R, digested with SacI and MluI, and inserted into the corresponding region of pUCp27-FLAG. pUCp27!FLAG/K25A; pUCp27!FLAG/R42A; pUCp27!FLAG/K43A; pUCp27!FLAG/H53A; pUCp27!FLAG/K86A; pUCp27!FLAG/R164A; pUCp27!FLAG/E178A; pUCp27!FLAG/H179A; pUCp27!FLAG/H180A; pUCp27!FLAG/HH179–180AA: DNA fragments were amplified by PCR from pUCp27-FLAG. The primer pairs used were pUCSacI-F plus one each of the following: K25A-R, R42A-R, K43A-R, H53A-R, K86A-R, R164A-R, E178A-R, H179A-R, H180A-R, or HH179–180AA-R. Another primer, 3!UTR-R, was used together with one each of the following: K25A-F, R42A-F, K43A-F, H53A-F, K86A-F, R164A-F, E178A-F, H179A-F, H180A-F, or HH179–180AA-F. Each recombinant PCR fragment was amplified with the primer pair pUCSacI-F and 3!UTR-R, digested with SacI and MluI, and inserted into the corresponding region of pUCp27-FLAG.

20 pUBp27!FLAG; pUBp27!FLAG/R164A; pUBp27!FLAG/E178A; pUBp27!FLAG/H179A; pUBp27!FLAG/H180A; pUBp27!FLAG/HH179–180AA: DNA fragments were amplified by PCR from each of the following: pUCp27-FLAG, pUCp27-FLAG/R164A, pUCp27-FLAG/E178A, pUCp27-FLAG/H179A, pUCp27-FLAG/H180A, and pUCp27-FLAG/HH179–180AA. The primer pairs used were 22R plus FLAG–KpnI-R. Each amplified PCR fragment was digested with EcoRI and KpnI, and inserted into the corresponding region of pUBp27. pUBp27!FLAG/K25A; pUBp27!FLAG/R42A; pUBp27!FLAG/K43A; pUBp27!FLAG/H53A; pUBp27!FLAG/K86A: DNA fragments were amplified by PCR from pUBp27-FLAG. The primer pairs used were M4 plus one each of the following: K25A-R, R42A-R, K43A-R, H53A-R, or K86A-R. Another primer, FLAG–KpnI-R, was used together with one each of the following: K25A-F, R42A-F, K43A-F, H53A-F, or K86A-F. Each recombinant PCR fragment was amplified with the primer pair M4 and FLAG–KpnI-R, digested with XbaI and KpnI, and inserted into the corresponding region of pUBp27-FLAG. pColdp27C!FLAG/R164A; pColdp27C!FLAG/E178A; pColdp27C!FLAG/H179A; pColdp27C!FLAG/H180A; pColdp27C!FLAG/HH179–180AA: DNA fragments were amplified by PCR from each of the following: pUCp27-FLAG/R164A, pUCp27-FLAG/E178A, pUCp27-FLAG/H179A, pUCp27-FLAG/H180A, and pUCp27-FLAG/HH179–180AA. The primer pairs used were KpnI–p27C-F plus FLAG–KpnI-R. Each amplified PCR fragment was digested with KpnI, and inserted into the corresponding region of pColdp27C-FLAG.

RNA preparation

21 pUCR2–3!-84-S and pUCR2–3!-84M-S were digested with XbaI; pRC2 or its derivatives were digested with XmaI; pUCR1p88, pUCp27-FLAG, and their derivatives were digested with SmaI. The RNA transcripts were synthesized from these plasmids as described previously (Iwakawa et al., 2008).

Preparation of BYL and BYLS20 The preparation of BYL and BYLS20 was as described previously (Komoda et al., 2004; Iwakawa et al., 2007 and 2011).

STagT affinity purification STagT affinity purification was performed essentially as described previously (Iwakawa et al., 2011), with minor modifications. Briefly, capped p27-FLAG or its derivatives (2 µg) were added to 100 µL of BYLS20 mixture. After incubation at 17 °C for 120 min, 3.2 µg of probe RNA was added to the BYLS20 mixture, which was incubated on ice for 20 min, followed by incubation on ice for 40 min in the presence of 100 µg of heparin. The mixture was added to a 400 µL bed volume of streptomycin-coupled Sepharose beads that had been preequilibrated with column buffer (50 mM Tris–HCl [pH 7.5], 100 mM NaCl, 3 mM MgCl2). After incubation for 3 min, the beads were washed three times with 2 mL of column buffer. The bound RNA and proteins were eluted with 1.5 mL of column buffer containing 10 µM streptomycin, followed by acetone precipitation. All the purification steps were performed at 4 °C. These samples were subjected to 12.5% SDS–PAGE, followed by ethidium bromide staining, and immunoblotting with an anti-FLAG M2 monoclonal antibody.

In vitro fractionation assay In the fractionation assays, capped p27-FLAG or one of its derivatives (1 µg) was added to 50 µL of BYL translation/replication mixture in the presence of RNA2 (500 ng). After incubation at 17 °C for 120 min, the samples were centrifuged (21,000g) at

22 4 °C for 10 min to separate the supernatant and pelleted fractions. To remove possible contaminating soluble proteins, the pellet was washed with TR buffer (30 mM HEPES– KOH [pH 7.4], 100 mM KOAc, 2 mM Mg (OAc)2, 2 mM DTT) and then centrifuged at 4 °C for 10 min, after which the pellet was lysed in solubilization buffer (50 mM Tris– HCl [pH 8.0], 150 mM NaCl, 10 mM MgCl2, 1 mM DTT, 0.5% Triton X-100). Aliquots of each fraction were subjected to northern and immunoblotting analyses, as described previously (Iwakawa et al., 2011).

Replication assay The BYL translation/replication assay was performed as described previously (Iwakawa et al., 2007). Briefly, capped p27-FLAG or one of its derivatives (1 µg) was added to 30 µL of BYL translation/replication mixture in the presence of uncapped RNA2 (500 ng) and capped RNA1-p88 (300 ng). The BYL translation/replication mixture was incubated at 17 °C for 240 min. Aliquots of the reaction mixture were subjected to northern and immunoblotting analyses, as described previously (Iwakawa et al., 2007 and 2011). BN-PAGE analysis was performed as described previously (Mine et al., 2010a and b) with minor modifications. Briefly, the protein samples prepared from solubilized total BYL were subjected to BNPAGE, followed by immunoblotting using an anti-FLAG M2 monoclonal antibody. To assay viral replication in the protoplasts, approximately 5Å~105 tobacco BY2 protoplasts were resuspended in MMG buffer (0.4 M mannitol, 15 mM MgCl2, 5 mM MES [pH 5.7]) and inoculated with RNA2 (1 µg) and the following amounts of plasmids: 10 µg of pUBp27-FLAG or its derivatives and 5 µg of pUBp88. A twofold volume of polyethylene glycol (PEG) solution (40% PEG 4000 [Fluka], 0.4 M mannitol, 200 mMCa(NO3)2) was added, and the mixture was diluted with 2 mL of dilution buffer (0.4 M mannitol, 125 mM CaCl2, 5mM KCl, 5 mM glucose, 1.5 mM MES [pH 5.7]) and incubated on ice for 15 min. The transfected protoplasts were washed with 4

23 mL of dilution buffer. The protoplasts were incubated at 17 °C for 16 or 24 h in the dark. The total RNA and proteins were subjected to northern and immunoblotting analyses, respectively, as described previously (Iwakawa et al., 2011).

GST pull-down assay The expression and purification of the recombinant proteins and the GST pull-down assay were performed as described previously (Mine et al., 2010a).

24 Table I-1 List of primers and their sequences used for PCR to generate constructs.

Primer name Sequence

#1–20-F TTCGTACCAGTCATGTTCAATCCAGGCAAAATCCTGTCTGCAAT

#1–20-R GCCTGGATTGAACATGACTGGTACGAAAAGTAGAAATTTAAAAT

#21–40-F GTGTGGGTGAGTAAATTTCGCAAGTGGTTCTTTGGTCTCAACTTT

#21–40-R GAACCACTTGCGAAATTTACTCACCCACACCATTAATTTATCCA

#41–60-F GACTGTTGGAATCGCATTCCACTCATGCCACATTACACGGAGCA

#41–60-R GTGGCATGAGTGGAATGCGATTCCAACAGTCTATACCCAAGTTGC

#61–80-F GCGGTGGATGCCTTCGAAACCCCAGAATCCAAATTAGAAGACTG

#61–80-R GGATTCTGGGGTTTCGAAGGCATCCACCGCCCACATATGTGCAT

#81–100-F GACGACTTCTGCAGCTTCGATGAAGAGGTGTACAAGAAGGATGA

#81–100-R CACCTCTTCATCGAAGCTGCAGAAGTCGTCGACTACACGTTCCA

#101–113-F TCAGTAAATGAATTCATGAAACTCCAAAGGAGCGCTGCCAGGAA

#101–113-R CCTTTGGAGTTTCATGAATTCATTTACTGAGGTGTCCAGTTCCA

#114–136-F GATGAGGAAGGCGTGATTAAGGCTGTAGAAACGAGGATCAGAAA

#114–136-R TTCTACAGCCTTAATCACGCCTTCCTCATCCTTCTTGTACACCT

#137–156-F ATGATGCAAGCGGCCAAGGTAGACGAAGCTGCAGTTAGAGCAAC

#137–156-R AGCTTCGTCTACCTTGGCCGCTTGCATCATGCCTGGGCGCACAC

#157–176-F GGAGATGACATGGGAAACGAACACCACACCAATGCATTGGTGTA

#157–176-R GGTGTGGTGTTCGTTTCCCATGTCATCTCCAAAGATGGTATGGC

#177–196-F GGTGAGTTCAAGATCCAGAGGTCTATCGACAGCGTCAAGCTCGC

#177–196-R GTCGATAGACCTCTGGATCTTGAACTCACCACATATGTCACTGG

#197–236-F GACACCAGACGATTACAAGGACGACGATGA

#197–236-R CCTTGTAATCGTCTGGTGTCATGGCAAGGT

K25A-F GTAAATTCAATCCAGGCGCAATCCTGTCTGCAATCTGCAACTTG

K25A-R CAGATTGCAGACAGGATTGCGCCTGGATTGAATTTACTCACCCA

25 R42A-F ACTGTTGGAATCGCTTTGCCAAGTGGTTCTTTGGTCTCAACTTT

R42A-R AGACCAAAGAACCACTTGGCAAAGCGATTCCAACAGTCTATACC

K43A-F GTTGGAATCGCTTTCGCGCGTGGTTCTTTGGTCTCAACTTTGAT

K43A-R TTGAGACCAAAGAACCACGCGCGAAAGCGATTCCAACAGTCTAT

H53A-F GTCTCAACTTTGATGCAGCTATGTGGGCGGTGGATGCCTTCATT

H53A-R GCATCCACCGCCCACATAGCTGCATCAAAGTTGAGACCAAAGAA

K86A-F GCGAAACCCCAGAATCCGCATTAGAAGACTGTCTGGAACTGGAC

K86A-R TCCAGACAGTCTTCTAATGCGGATTCTGGGGTTTCGCTGCAGAA

R164A-F TAGACGAAGCTGCAGTTGCAGCAACTGCCAGTGACATATGTGGT

R164A-R ATGTCACTGGCAGTTGCTGCAACTGCAGCTTCGTCTACCTTTCC

E178A-F GTGAGTTCAAGATCAACGCACACCACACCAATGCATTGGTGTAT

E178A-R AATGCATTGGTGTGGTGTGCGTTGATCTTGAACTCACCACATAT

H179A-F AGTTCAAGATCAACGAAGCCCACACCAATGCATTGGTGTATGCC

H179A-R ACCAATGCATTGGTGTGGGCTTCGTTGATCTTGAACTCACCACA

H180A-F TCAAGATCAACGAACACGCCACCAATGCATTGGTGTATGCCGCA

H180A-R TACACCAATGCATTGGTGGCGTGTTCGTTGATCTTGAACTCACC

HH179–180AA-F TCAAGATCAACGAAGCCGCCACCAATGCATTGGTGTATGCCGCA

HH179–180AA-R ACCAATGCATTGGTGGCGGCTTCGTTGATCTTGAACTCACCACA pUCSacI-F AATTTCACACAGGAAACAGC

3’UTR-R GGGGTACCTAGCCGTTATAC

22R AGCAGATGGAACGTGTAG

FLAG–KpnI-R CGGGGTACCCTACTTGTCATCGTCGTCC

M4 GTTTTCCCAGTCACGAC

KpnI–p27C-F CGGGGTACCATGAAACTCCAAAGGAGCGCTGC

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Fig. I-1. Mapping the regions within p27 required for the p27–YRE interaction. (A) A schematic representation of the deleted derivatives of p27-FLAG. The deletions are represented as bent lines. All the mutants have a FLAG-tag sequence at their C-termini. (B) In vitro pull-down assay of FLAG-tagged p27 or its derivatives by Strepto-tagged YRE. (Top) Input of the indicated FLAG-tagged p27 proteins.

(Middle) FLAG-tagged p27 proteins copurified with Strepto-tagged RNA. Immunoblots were probed with an anti-FLAG antibody. (Bottom) Purified Strepto-tagged YRE. The gel was stained with ethidium bromide.

27 .

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Fig. I-2. Residues R164, H179, and H180 in p27 are important for its binding to YRE. (A) Sequence alignment of the regions within p27 involved in its binding to YRE in RCNMV and other dianthoviruses.

Asterisks indicate the conserved amino acids selected for mutagenesis in RCNMV p27. The following abbreviations are used: SCNMV, Sweet clover necrotic mosaic virus; CRSV, Carnation ring spot virus.

(B) In vitro pull-down assays of FLAG-tagged p27 and its alanine-scanning mutant derivatives by YRE.

Other details are as described in the legend to Fig. I-1.

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Fig. I-3. Effects of the mutations introduced into p27 on the recruitment of viral genomic RNA2 to the membrane fraction. (A) Immunoblot analysis of the accumulation of p27-FLAG and its derivatives in the pelleted fraction from BYL. (B) Northern blot analysis of the distribution of RNA2 in the total (T),

(21,000g centrifuged) supernatant (S), and pelleted (P) fractions from BYL expressing RNA2 with p27–

FLAG or its derivatives. The percentage of pellet-associated RNA2 (P/[S+P]) is graphed. The signal intensities were quantified with the Image Gauge program (Fuji Photo Film, Japan). The averages of three independent experiments are shown. Error bars indicate the standard deviations.

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Fig. I-4. Effects of the mutations introduced into p27 on the negative-strand synthesis and replication of

RNA2. (A) Accumulation of negative-strand RNA2 in BYL incubated with RNA2 and RNA1-p88, together with p27-FLAG or its derivatives. The total RNAs and proteins were extracted after incubation for 4 h and used for northern and immunoblotting analyses, respectively. The details are as described in

Materials and methods. The ribosomal RNA (rRNA) is shown below the northern blots as the loading control. (B) Full-length RNA2 was cotransfected with plasmid expressing p27-FLAG that carried a specific mutation and plasmid expressing p88 in BY2 protoplasts (5$~105 cells per experiment). The total

RNAs and proteins were extracted 16 h after infection and used for northern and immunoblotting analyses, respectively. The relative accumulation of negative- (A) and positive-strand (B) RNA2 with wild-type p27-FLAG and its derivatives is shown (wild-type p27-FLAG was set at 100%). The signal intensities were quantified with the Image Gauge program. The averages of three independent experiments are shown. Error bars indicate the standard deviations.

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Fig. I-5. Effects of the mutations introduced into p27 on the negative-strand synthesis and replication of

RNA1. (A) The accumulation of negative-strand RNA1-p88 in BYL incubated with RNA1-p88, together with p27-FLAG or its derivatives. (B) p27-FLAG and its derivatives were coexpressed with RNA1-p88 from plasmids in the protoplasts. The total RNAs and proteins were extracted 24 h after infection and used for northern and immunoblotting analyses, respectively. For other details, refer to the legend to Fig.

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RCNMV replicase proteins. (A) SDS–PAGE analysis of purified His–p27C–FLAG or its derivatives expressed in E. coli. The purified proteins (500 ng) were visualized with Coomassie Brilliant Blue (CBB) staining (left panel), and analyzed by immunoblotting with anti-FLAG antibody (right panel). (B and C)

In vitro interactions between His–p27C–FLAG or its derivatives and His/GST–p27C or His/GST– p88#p27. His–p27C–FLAG or its derivatives were incubated with glutathione-resin-bound His/GST– p27C (B) or His/GST–p88#p27 (C). After they were washed, the pulled-down complexes were subjected to SDS–PAGE and blotted onto membrane. The separated proteins on the membrane were visualized with

Ponceau S staining (upper panel), and analyzed by immunoblotting with anti-FLAG antibody (lower panel).

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Fig. I-7. Effects of the mutations introduced into p27 on the formation of the 480-kDa RCNMV replicase complex. Accumulation of the 480-kDa complex in BYL incubated with transcripts that expressed the p27-FLAG mutants and p88 in the presence of RNA2. The total proteins were extracted 4 h after incubation and subjected to SDS–PAGE and BN-PAGE analyses, followed by immunoblotting with an anti-FLAG antibody.

33 Chapter II ADP Ribosylation Factor 1 Plays an Essential Role in the Replication of a Plant RNA Virus

Introduction Eukaryotic positive-strand RNA [(+)RNA] viruses replicate their genomes using membrane-bound replicase complexes, which contain multiple viral and host components. A growing number of host proteins that affect viral RNA replication have been identified using genome-wide and proteomics analyses in several animal and plant viruses (Berger et al., 2009; Cherry et al., 2005 and 2006; Gancarz et al., 2011; Jiang et al., 2006; Kushner et al., 2003; Li et al., 2009; Panavas et al., 2005; Reiss et al., 2011; Serva and Nagy. 2006; Shah Nawaz-ul-Rehman et al., 2012; Tai et al., 2009; Zhu et al., 2007). These host proteins are involved in translation, template selection, and the assembly of the viral replication complex (VRC) on intracellular membranes, which serve as the site of viral RNA replication (Nagy and Pogany. 2012). However, the functions of host proteins remain largely unknown. The replication compartments of (+)RNA viruses are derived from various cellular organelle membranes, such as the endoplasmic reticulum (ER), mitochondria, chloroplasts, peroxisomes, and the Golgi apparatus (den Boon and Ahlquist. 2010; den Boon et al., 2010; Laliberté and Sanfaçon. 2010). The formation of viral replication compartments generally involves the emergence of spherules, vesicles, and multivesicular bodies associated with various organelles (den Boon and Ahlquist. 2010; Laliberté and Sanfaçon. 2010). Although viral proteins play an essential role in the formation of replication compartments containing VRCs, host factors also regulate this process (den Boon and Ahlquist. 2010; Miller and Krijnse-Locker. 2008; Nagy and Pogany. 2012). Tomato bushy stunt virus (TBSV) coopts the proteins of the endosomal sorting complexes that are required for transport (ESCRT) to assemble the replicase

34 complex properly on the peroxisome membrane via an interaction with the auxiliary replication protein p33 (Barajas et al., 2009; Barajas and Nagy. 2010). ESCRT proteins play a major role in the sorting of ubiquitin-modified cargo proteins from the endosomal membrane to the internal vesicles of multivesicular bodies (Reyes et al., 2011). Brome mosaic virus (BMV) replication protein 1a interacts with the reticulon homology proteins (Rhps), which play an important role in the formation of the VRC, probably by regulating membrane curvature (Diaz et al., 2010). Coxsackievirus B3 (CVB3) 3A protein recruits 4-kinase III% (PI4KIII%) to the viral replication site to facilitate the formation of the phosphatidylinositol-4-phosphate-enriched compartment, which has a high affinity for the 3D RNAdependent RNA polymerase (RdRP) (Hsu et al., 2010). Another PI4KIII, PI4KIII& is required for hepatitis C virus (HCV) replication (Berger et al., 2009; Reiss et al., 2011; Tai et al., 2009; Tai and Salloum. 2011). Red clover necrotic mosaic virus (RCNMV) is a (+)RNA plant virus that is a member of the genus Dianthovirus in the family Tombusviridae. The genome of RCNMV consists of RNA1 and RNA2. RNA1 encodes the p27 auxiliary replication protein, the p88 RdRP, and the coat protein (Xiong et al., 1993a; Xiong and Lommel. 1989). RNA2 encodes the movement protein (MP) that is required for viral cell-to-cell movement (Kaido et al., 2009; Xiong et al., 1993b). p27, p88, and host proteins form the 480-kDa replicase complex, which is a key player in the replication of RCNMV RNA (Mine et al., 2010b). p27 and p88 colocalize at the ER membrane (Kusumanegara et al., 2012; Turner et al., 2004). RCNMV infection or the expression of p27 is sufficient for inducing a large aggregate structure and the proliferation of the ER membrane (Kusumanegara et al., 2012; Mine et al., 2012; Turner et al., 2004). I previously found that an affinity-purified RdRP fraction of RCNMV-infected leaves contains several host proteins, including heat shock protein 70 (HSP70), HSP90, and ribosomal proteins (Mine et al., 2010b). To identify further the host factors involved in RCNMV replication, I reevaluated the same affinity-purified RdRP fraction using a MASCOT search and identified ADP ribosylation factor 1 (Arf1). Arf1 is a

35 highly conserved, ubiquitous, small GTPase and is implicated in the formation of coat protein complex I (COPI) vesicles on Golgi membranes (D’Souza-Schorey and Chavrier. 2006; Memon. 2004). Brefeldin A (BFA) is a well-known fungal metabolite that inhibits the early secretory pathway (Nebenführ et al., 2002). BFA inhibits the activation of Arf small GTPases by targeting BFA-sensitive guanine nucleotide-exchange factors (GEFs) (Geldner et al., 2003; Teh and Moore. 2007) via the locking of the abortive Arf-GDPGEF complex, thereby blocking guanine nucleotide release (Mossessova et al., 2003). Arf1 and its BFA-sensitive GEFs are required for the replication of several vertebrate (+)RNA viruses, such as poliovirus, CVB3, mouse hepatitis coronavirus (MHV), and HCV (Belov et al., 2007, 2008, and 2010; Cuconati et al., 1998; Goueslain et al., 2010; Hsu et al., 2010; Lanke et al., 2009; Matto et al., 2011; Verheije et al., 2008). However, the direct involvement of Arf1 in RNA replication of plant (+)RNA viruses has not been reported (Verchot. 2011). In this study, using in vitro pulldown and bimolecular fluorescence complementation (BiFC) analyses, I show that Arf1 interacts with the RCNMV replication protein p27 within the virusinduced large punctate structures of the ER membrane. I found that BFA treatment reduced the accumulation of the 480-kDa viral replicase complex and RCNMV RNA and decreased p27-induced ER proliferation in RCNMV-infected tobacco BY-2 protoplasts. Similarly, expression of dominant negative Arf1 mutants compromised RCNMV RNA replication in protoplasts. Interestingly, expression of the dominant negative mutant of Sar1, which is a key regulator of the biogenesis of the COPII vesicles at ER exit sites (ERES), also reduced the accumulation of RCNMV RNA. These results suggest that the RNA replication of RCNMV depends on the host membrane traffic machinery.

36 RESULTS

Arf1 interacts directly with p27. I reevaluated the data obtained by LC-MS/MS analyses of the affinity-purified RdRP membrane fraction (Mine et al., 2010b) and found that Arf1 was present in the fraction that coimmunoprecipitated with FLAG-tagged p27 (Table II-2). This result led us to perform a GST pulldown assay to determine whether Arf1 interacts with p27. Bacterially expressed and purified p27 with an N-terminal 6$His tag and C-terminal FLAG tag (His-p27-FLAG) (Mine et al., 2010a) was incubated with N-terminally GST-fused Arf1 (GST-Arf1) or GST captured on glutathione-bound beads. Immunoblot analyses using an anti-FLAG antibody showed that His-p27-FLAG was pulled down by GST-Arf1 but not by GST (Fig. II-1A), indicating that p27 binds to Arf1 in vitro. A parallel GST pulldown assay using purified N-terminally 6$His-tagged and C-terminally FLAG-tagged Arf1 (His-Arf1-FLAG) (Fig. II-1B) and glutathione resin-bound N-terminally GST-fused p27 or its truncated proteins (Fig. II-1C) confirmed a p27-Arf1 interaction in vitro (Fig. II-1D). Moreover, His-Arf1-FLAG was pulled down more efficiently by the C-terminal half of p27 than by the N-terminal half of p27, suggesting that Arf1 interacted preferentially with the C-terminal region of p27 (Fig. II-1D). To further test the interaction between Arf1 and p27 in vivo, I performed a BiFC experiment in N. benthamiana epidermal cells. Arf1 that was fused to the N-terminal half of yellow fluorescent protein (YFP) at the C terminus (Arf1-nYFP) and p27 that was fused to the C-terminal half of YFP at the C terminus (p27-cYFP) were expressed together with mCherry containing an ER targeting signal (ER-mCherry) and the TBSV silencing suppressor p19 in N. benthamiana via agroinfiltration. At 3 dpi, fluorescence was observed using confocal laser scanning microscopy (CLSM). YFP fluorescence was reconstituted in the presence of Arf1-nYFP and p27-cYFP (Fig. II-1E, top left

37 panel). YFP fluorescence merged well with ER-mCherry fluorescence in the large punctate structures (Fig. II-1E, top panels), a characteristic feature of morphological ER changes induced by p27 or RCNMV infection (Kusumanegara et al., 2012; Mine et al., 2012; Turner et al., 2004). Negligible or no YFP fluorescence was detected in control experiments (Fig. II-1E, middle and bottom panels). These results indicate that p27 interacts directly with Arf1 at the p27-induced modified ER membrane.

Arf1 is recruited from the Golgi apparatus to the RCNMV replication site by p27. In plant cells, Arf1 is localized at the Golgi apparatus (Cui et al., 2010; Lee et al., 2002; Stefano et al., 2006; Takeuchi et al., 2002). However, the Arf1-p27 interaction occurred at the ER in N. benthamiana epidermal cells (Fig. II-1E). This result prompted us to test the possibility that p27 recruits Arf1 from the Golgi apparatus to the ER membrane, which is a putative site of RCNMV replication (Turner et al., 2004). First, I examined whether p27 colocalizes with Arf1. For this, a plasmid expressing C-terminally GFP-fused Arf1 (Arf1-GFP) was cotransfected with a plasmid expressing C-terminally mCherry-fused p27 (p27-mCherry) or a control plasmid into tobacco BY-2 protoplasts, and fluorescence was observed via CLSM after 16 h of incubation. In the absence of p27-mCherry, Arf1-GFP fluorescence was observed as small punctate structures throughout the cell, a typical fluorescence pattern of Golgi compartment-localized proteins (Fig. II-2A) (Boevink et al., 1998; Nebenführ et al., 1999). Interestingly, however, in the presence of p27-mCherry, the small punctate fluorescence pattern of Arf1-GFP was dramatically reduced, and Arf1-GFP fluorescence was observed as large aggregate structures located around the nucleus (Fig. II-2B, left panel). Moreover, these aggregated structures colocalized with p27-mCherry (Fig. II-2B, middle and right panels). Similar results were obtained for Erd2-GFP (data not shown), a well-established cis-Golgi marker (Saint-Jore et al., 2002). Expression of C-terminally mCherry-fused p88 (p88-mCherry) also affected the distribution pattern of Arf1-GFP, but large aggregated structures of Arf1-GFP were not observed (Fig. II-2C).

38 p27-mCherry and p88-mCherry efficiently supported the replication of RNA2 (data not shown). It should be noted that both mCherry-fused p27 and p88 colocalized with the ER marker protein in BY-2 protoplasts (see Fig. II-5; also data not shown).

Arf1 plays an important role inRCNMVRNAreplication. To investigate whether Arf1 is required for infection of host plants with RCNMV, I downregulated Arf1 using Tobacco rattle virus (TRV)-based virus-induced gene silencing in N. benthamiana plants. A TRV vector harboring a partial fragment of NbArf1 (TRV:NbArf1) was expressed via Agrobacterium-mediated expression. An empty TRV vector (TRV:00) was expressed as a control. Newly developed leaves were inoculated withRCNMVRNA1 and RNA2 at 7 dpi. Two days after inoculation, three inoculated leaves from three different plants were harvested and mixed, and total RNA was extracted. Semiquantitative RT-PCR analyses confirmed the specific reduction of NbArf1 mRNA in plants infiltrated with TRV:NbArf1 (Fig. II-3A). Northern blot analyses showed that the accumulation of RCNMV RNA was reduced 10-fold in NbArf1-silenced plants compared with control plants (Fig. II-3A), suggesting that Arf1 plays a positive role in RCNMV infection. It has been well documented that BFA inhibits the secretion of proteins from the Golgi apparatus back to the ER (Nebenführ et al., 2002) via inhibition of the nucleotide-exchange reaction of Arf proteins (Mossessova et al., 2003). To investigate whether Arf1 plays a role in RCNMV RNA replication, I tested the effect of BFA on the accumulation of RCNMV RNA replication in a single cell. Tobacco BY-2 protoplasts were inoculated withRCNMVRNA1 and RNA2 and incubated for 16 h in the presence or absence of BFA. Northern blot analyses showed that the accumulation of viral RNAs was reduced 5- to 10-fold in the presence of BFA (Fig. II-3B). BFA did not affect the accumulation of rRNA (Fig. 3B, bottom panel). Moreover, BFA had a negligible effect on the accumulation of BMV RNA in BY-2 protoplasts (Fig. II-3C). These results indicate that the inhibitory effect of BFA on RCNMV RNA replication

39 was not due to its nonspecific toxicity and that the inhibition of Arf function by BFA specifically affects RCNMV RNA replication. Next, to investigate whether the inhibitory effect of BFA is due to the inhibition of Arf1 GEF activity, I tested the effects of constitutive-negative (Arf1 with a T31N mutation [Arf1-T31N]) and constitutive-active (Arf1-Q71L) Arf1 mutants on RCNMV RNA replication. Arf1-T31N is a GDP-restricted Arf1 mutant that impairs the normal COPI function and inhibits the formation of ERES (Lee et al., 2002; Stefano et al., 2006; Takeuchi et al., 2002; Xu and Scheres. 2005). Arf1-Q71L is a GTP-restricted Arf1 mutant that inhibits the vacuolar sorting of sporamin in tobacco BY-2 cells (Takeuchi et al., 2002). Tobacco BY-2 protoplasts were transfected with plasmids expressing wild-type Arf1, Arf1-T31N, or Arf1-Q71L together with plasmids expressing RCNMV RNA1 and RNA2. Northern blot analyses showed that the expression of Arf1 reduced the accumulation of viral RNAs by about 25% (Fig. II-3D, first and second lanes), whereas the expression of Arf1-T31N or Arf1-Q71L reduced the accumulation of viral RNAs by more than 80% (Fig. II-3D, third and fourth lanes). These results suggest that the constitutive-active mutant of Arf1 is insufficient to support the replication of RCNMV RNA but that GDP/GTP cycling of Arf1 is required for viral RNA replication. Taken totogether, these results strongly suggest that Arf1 plays an essential role in RCNMV RNA replication.

Arf1 is required for the assembly of the viral replicase complex. The replication of (+)RNA viruses proceeds through many steps, including translation of viral replication proteins, formation of the VRC on appropriate intracellular membranes, and viral RNA synthesis (Nagy and Pogany. 2012). To determine which step(s) of RCNMV RNA replication requires Arf1 function, I first tested whether BFA affects the translation of RCNMV RNA1 using an uncapped reporter RNA containing the firefly luciferase (Luc) gene flanking the 5= and 3= untranslated region (UTR) of the RCNMV RNA1 (R1-Luc-R1) and a capped nonviral reporter Luc RNA with a 3'poly(A) tail (Luc-pA60) (Fig. II-4A) (Iwakawa et al., 2012;

40 Sarawaneeyaruk et al., 2009). Tobacco BY-2 protoplasts were inoculated with these reporter RNAs and incubated for 6 h in the presence or absence of BFA. Subsequently, luciferase activity was measured. BFA did not affect the translational activity of these RNAs (Fig. II-4A), suggesting that BFA affects neither 3=cap-independent translation element (CITE)-mediated translation of RCNMV RNA nor canonical cap-dependent translation. Next, I investigated whether BFA affects the formation of the 480-kDa viral RNA replicase complex (Mine et al., 2010b). I expressed C-terminally hemagglutinin (HA)-tagged p27 (p27-HA) and p88 (p88-HA) together with RNA2 in BY-2 protoplasts and analyzed the accumulation of the viral replicase complexes by SDS- and BNPAGE using anti-HA antibody. The accumulation of the 480-kDa replicase complex was significantly decreased by BFA treatment (Fig. II-4B, upper panel), whereas the accumulation of p27-HA and p88-HA was not affected by BFA (Fig. II-4B, lower panel), suggesting that Arf1 is required for the formation of the RCNMV replicase complex. Because the formation of the 480-kDa complex requires p27-p27/p88 interactions (Hyodo et al., 2011; Mine et al., 2010a), I examined the effect of BFA on the interactions between viral replication proteins. For this, BY-2 protoplasts were transfected with plasmids expressing p27-HA and p88-HA and RNA2 together with either p27 or C-terminally FLAG-tagged p27 (p27-FLAG). After 16 h of incubation in the presence or absence of BFA, cell extracts were subjected to FLAG affinity resin, and eluates were analyzed by immunoblotting using an anti-HA antibody. BFA had no effect on the copurification of either p27-HA or p88-HA by p27-FLAG (Fig. II-4C, lanes 5 and 6). p27-HA and p88-HA were not immunoprecipitated with the anti-FLAG antibody (Fig. II-4C, lane 4), ruling out the possibility of nonspecific copurification. These results showed that BFA does not prevent the p27-p27/p88 interactions.

Arf function is required for p27-mediated ER modification.

41 Because the membrane association of p27 is critical for the assembly of the 480-kDa replicase complex (Kusumanegara et al., 2012) and because Arf1 is involved in COPI vesicle formation and membrane curvature (Pucadyil and Schmid. 2009), I investigated whether Arf1 function is required to keep the ER membrane association of p27. For this, BY-2 protoplasts were transfected with a plasmid expressing GFP containing an ER-targeting sequence (ER-GFP) together with or without a plasmid expressing p27-mCherry. Protoplasts were incubated for 14 h, followed by an additional incubation of 2 h with BFA or DMSO. As reported previously (Kusumanegara et al., 2012; Turner et al., 2004), p27-mCherry colocalized with ER-GFP in p27-induced large aggregate structures in the DMSO control (Fig. II-5B, upper panels). In contrast, BFA treatment canceled the colocalization of p27-mCherry with ER-GFP in protoplasts treated with BFA (Fig. II-5B, lower panels). The fluorescence of both ER-GFP and p27-mCherry dispersed in the BFA-treated protoplasts compared with that observed in control protoplasts (Fig. II-5B, data not shown). Interestingly, in the absence of p27-mCherry, BFA treatment did not affect the distribution patterns of ER-GFP (Fig. II-5A). Moreover, BFA had no effect on the distribution patterns of either p27-HA or p88-HA, as assessed using a fractionation assay (Fig. II-5C). These observations suggest that Arf1 is required for p27-mediated ER modification.

Disturbance of COPII vesicle formation inhibits the replication of RCNMV RNA. It is known that BFA treatment or expression of Arf1-T31N not only impairs the normal COPI function but also inhibits COPII vesicle formation (Stefano et al., 2006). COPII vesicles bud from ERES and are thought to mediate anterograde traffic out of the ER (daSilva et al., 2004; Marti et al., 2010; Yang et al., 2005). Sar1 is a small GTPase that is an essential cytosolic component of the COPII complex that accumulates at ERES (Hanton et al., 2008). To examine whether the COPII vesicle transport system is involved in RCNMV RNA replication, I used a dominant negative mutant of Sar1 (Sar1-H74L). Sar1-H74L is the GTP-restricted mutant that exerts a known dominant

42 negative effect on COPII vesicle transport. This mutant was shown previously to trap vesicles in a coated configuration so that they are unable to fuse with the target membrane (daSilva et al., 2004; Takeuchi et al., 2000). Tobacco BY-2 protoplasts were transfected with plasmids expressing either wild-type Sar1 or Sar1-H74L together with plasmids expressing RCNMV RNA1 and RNA2, and the accumulation of RNA1 and RNA2 was determined at 24 hpi. Northern blot analyses showed that the expression of Sar1 did not significantly affect the accumulation of viral RNAs (Fig. II-6A, first and second lanes). In contrast, the expression of the Sar1-H74L mutant dramatically reduced the accumulations of viral RNAs (Fig. II-6A, third lane). The fluorescence of C-terminally GFP-fused Sar1 (Sar1-GFP) was observed as small punctate structures dispersed in cells (Fig. II-6B). Interestingly, coexpression of p27-mCherry redistributed Sar1-GFP to perinuclear aggregated structures in which these proteins were well colocalized (Fig. II-6C).

RCNMV replication proteins colocalize with newly synthesized viral RNAs. To determine whether newly synthesized RCNMVRNAs colocalized with viral replication proteins, I performed double immunofluorescence staining of RCNMV-infected tobacco BY-2 protoplasts. BY-2 protoplasts were inoculated with RNA1 and RNA2. After 14 h of incubation, protoplasts were incubated with actinomycin D for 1 h to block host DNA-dependent RNA polymerases. Protoplasts were then incubated with BrUTP for an additional 3 h, fixed, and processed for double immunofluorescence labeling using an anti-p27 antiserum and an antibody that recognizes bromouridine-containing RNA. To assess the background fluorescence, mock-inoculated protoplasts were subjected to the same immunofluorescence labeling conditions, and fluorescent signals were adjusted to set the background threshold level. No significant background was detected in uninfected protoplasts (Fig. II-7). Immunofluorescence labeling showed that bromouridine-labeled RNA (red) colocalized

43 with p27 (green) (Fig. II-7), indicating that RCNMV replication proteins colocalize with newly synthesized viral RNA.

DISCUSSION

In this study, I found that p27 interacts with Arf1 and relocalizes this protein to the aggregate structures of ER membranes, where they colocalize (Fig. II-1 and II-2), suggesting that p27 recruits Arf1 to the viral replication sites. Membrane remodeling, including the formation of the aggregate structures and membrane proliferation, has been observed in plant cells infected with RCNMV or expressing p27 alone (Kusumanegara et al., 2012; Mine et al., 2012; Turner et al., 2004). Arf1 seems to be required for the formation of virus-induced ER membrane proliferation. Pharmacological inhibition of Arf function by BFA canceled the p27-induced formation of aggregate structures on ER membranes (Fig. II-5B). However, BFA treatment did not affect the membrane association of p27 (Fig. II-5C) or the localization of ER-GFP, an ER marker in the absence of p27 (Fig. II-5A). Moreover, the N-terminal half of p27 interacted poorly with Arf1 (Fig. II-1D) but retained the ability to efficiently localize at the ER membrane (Kusumanegara et al., 2012). This truncated p27 mutant does not induce large aggregate structures on ER membranes although it induces smaller punctate structures (Kusumanegara et al., 2012). All these observations strongly support the importance of functional Arf1 in membrane remodeling, such as the induction of a large aggregate structure that is likely the site of vital RNA replication (Fig. II-7).

In animal and yeast cells, it is known that membrane-bound Arf1 can recruit a diverse array of effectors, including COPI, clathrin, cytoskeletal regulators, and lipid-modifying enzymes, such as phospholipase D and PI4KIII% (Donaldson and

44 Jackson. 2011). Arf1 also plays a crucial role in the replication of several vertebrate (+)RNA viruses. Arf1 colocalizes with the enteroviral replication machinery (Belov et al., 2007; Hsu et al., 2010) and binds to and hydrolyzes GTP during infection, suggesting the utilization of Arf1 effectors by the virus (Altan-Bonnet and Balla. 2012). The enterovirus replication protein 3A recruits PI4KIII%, one of the Arf1 effectors, over many other Arf1 effector proteins to membranes (Hsu et al., 2010). Although there is no information regarding the function of Arf1 effector proteins as lipid-modifying enzymes in plant cells (Memon. 2004), plant Arf1 effectors might contribute to the establishment of VRCs of RCNMV via a mechanism similar to that reported in vertebrate viruses (Hsu et al., 2010). Interestingly, I noted that treatment of RCNMV-infected BY-2 protoplasts with 1-butanol, which inhibits the production of phosphatidic acid catalyzed by phospholipase D (Brown et al., 1993), inhibited the accumulation of viral RNAs (K. Hyodo and T. Okuno, unpublished data). Arf1 appears to manage not only coat recruitment but also curvature generation in membranes (Pucadyil and Schmid. 2009). After membrane binding, Arf1 can remodel membranes into highly curved tubules in vitro (Beck et al., 2008; Krauss et al., 2008). The membrane deformation activity of Arf1 per se may contribute to the formation of RCNMV VRCs. In the case of BMV, Rhps are recruited to viral RNA replication site by 1a protein to induce the membrane curvature (Diaz et al., 2010).

BFA treatment or expression of dominant negative mutants of Arf1 in plant cells not only inhibits the COPI pathway but also compromises COPII vesicle trafficking (Stefano et al., 2006). Interestingly, I found that the accumulation of RCNMV RNA was inhibited by the expression of the Sar1 H74L mutant (Fig. II-6A), which inhibits COPII vesicle formation in plant cells (daSilva et al., 2004; Takeuchi et al., 2000). Sar1 was relocalized with p27 in p27-induced large aggregate structures of ER membranes (Fig. II-6B and C). These results suggest that the COPII vesicle trafficking system is required for the formation of the RNA replication compartment in RCNMV. The COPII

45 assembly system also plays an important role in MHV and poliovirus replication (Oostra et al., 2007; Trahey et al., 2012). Poliovirus induces COPII vesicles in the early replication phase, and viral RNA localizes at or near the ERES (Hsu et al., 2010; Trahey et al., 2012).

The p27-induced remodeling of the early secretory pathway is reminiscent of the membrane remodeling caused by potato virus X (PVX) and turnip mosaic virus (TuMV) infection (Grangeon et al., 2012; Tilsner et al., 2012). The triple gene block 1 (TGB1) of PVX organizes the X body, a virally induced inclusion structure that contains host actin, the ER, and the Golgi apparatus, at the perinuclear region (Tilsner et al., 2012). However, TGB1 is not required for PVX replication. Therefore, the X body seems to couple RNA replication and cell-to-cell movement processes (Tilsner et al., 2012). TuMV infection leads to the amalgamation of the host ER, Golgi apparatus, components of the COPII coatomer, and chloroplasts into a perinuclear globular structure that also contains viral proteins (Grangeon et al., 2012). However, the formation of this globular structure is independent of the early secretory pathway (Grangeon et al., 2012). In contrast to TuMV, p27-induced membrane remodeling relied on the functional secretory pathway because BFA treatment disrupted this membrane remodeling (Fig. II-5). Rearrangement of host endomembrane systems has also been reported in several vertebrate (+)RNA viruses (Miller and Krijnse-Locker. 2008). Remodeling of the early secretory pathway may be a common phenomenon during (+)RNA virus infection although the molecular mechanisms underlying its formation may be different in each virus.

BFA inhibits Arf function through BFA-sensitive GEFs, which assist the GDP-GTP exchange reaction of Arf proteins (Geldner et al., 2003; Teh and Moore. 2007). Poliovirus uses Arf1 and GBF1, which is a BFA-sensitive Arf-GEF (Belov et al., 2008). The poliovirus 3A protein interacts with GBF1 (Teterina et al., 2011). However, it is

46 well known that the replication of several viruses in the Picornaviridae is insensitive to BFA (Irurzun et al., 1992; Sasaki et al., 2012). Among plant RNA viruses, TuMV, melon necrotic spot virus (MNSV), and BMV were insensitive to BFA during RNA replication (Genovés et al., 2010; Grangeon et al., 2012) (Fig. II-3C), whereas grapevine fanleaf virus (Ritzenthaler et al., 2002) and RCNMV (Fig. II-3) were sensitive to BFA during RNA replication. The BFA-insensitive viruses may require BFA-insensitive GEFs or other host factors involved in the secretory pathway. Recent proteomics analyses suggested that a large number of host proteins involved in protein transport affect TBSV replication (Shah Nawaz-ul-Rehman et al., 2012). Interestingly, downregulation of the COPI coatomer subunit decreases TBSV replication but increases recombination in Saccharomyces cerevisiae (Jiang et al., 2006; Serviene et al., 2006). (+)RNA viruses may evolve to hijack a diverse set of component proteins in the host secretory pathway to accomplish efficient viral infection.

The involvements of the early secretory pathway in the cell-to-cell movement of several plant (+)RNA viruses has been reported (Harries et al., 2010). Disruption of the early secretory pathway by BFA treatment or expression of a dominant negative mutant of Arf1 in N. benthamiana epidermal cells leads to the inhibition of cell-to-cell movement of TuMV and MNSV, with little effect on viral RNA replication (Genovés et al., 2010; Grangeon et al., 2012). Although inhibition of the early secretory pathway by BFA compromised the RNA replication of RCNMV, it is possible that the early secretory pathway also affects the cell-to-cell movement of RCNMV. Our previous study showed that RCNMV MP localizes to the cell wall and later to the ER (Kaido et al., 2009). Interestingly, the replication of RCNMV RNA1 is coupled to the ER localization of MP (Kaido et al., 2009). p27-induced endomembrane remodeling might contribute not only to VRC formation but also to the recruitment of MP to the VRC and subsequent cell-to-cell movementof RCNMV, which is required for efficient viral spread.

47 MATERIALS AND METHODS

Gene cloning and plasmid construction. pUCR1 (Takeda et al., 2005) and pRC2_G (Xiong and Lommel. 1991) are full-length cDNA clones of RNA1 and RNA2 of an RCNMV Australian strain, respectively. pB1TP3, pB2TP5, and pB3TP8 are full-length cDNA clones of RNA1, RNA2, and RNA3 of the BMV M1 strain, respectively (Janda et al., 1987) (generous gift from Paul Ahlquist). The constructs described previously used in this study include pBICp27-HA:cYFP (where HA is hemagglutinin and cYFP is a C-terminal fragment of the yellow fluorescent protein) (Mine et al., 2012), pBICHA:cYFP (Mine et al., 2012), pBICmyc:nYFP (where nYFP is an N-terminal fragment of YFP) (Mine et al., 2012), pBICp19 (Takeda et al., 2005), pBICER:mCherry (Kaido et al., 2011), pUBp27-FLAG (Hyodo et al., 2011), pUBp27-HA (Mine et al., 2012), pUBp88-HA (Mine et al., 2012), pCOLDp27 (Mine et al., 2010a), pCOLDp27N (Mine et al., 2010a), pCOLDp27C (Mine et al., 2010a), pCOLDGSTp27 (where GST is glutathione S-transferase) (Mine et al., 2010a), pR1-Luc-R1 (where Luc is luciferase) (Sarawaneeyaruk et al., 2009), pLUCpA60 (Mizumoto et al., 2003), and pSP64-RLUC (Mizumoto et al., 2003). pUC118 was purchased from TaKaRa Bio Inc. (Shiga, Japan). Escherichia coli DH5_ was used for the construction of all plasmids. All plasmids constructed in this study were verified by sequencing. The primers used in this study are listed in Table II-1. RNA extraction from Nicotiana benthamiana leaves, tobacco (Nicotiana tabacum) BY-2 cells, or Arabidopsis thaliana (Col-0) leaves was performed using an RNeasy Plant Mini Kit (Qiagen, Hilden, Germany). Reverse transcription-PCR (RT-PCR) was catalyzed by Superscript III reverse transcriptase (Invitrogen) using oligo(dT) (Mine et al., 2012).

48 pTVNbArf1. A 399-bp Arf1 cDNA fragment (Coemans et al., 2008) was amplified from cDNA derived from N. benthamiana (NbArf1) RNA using primers 1 and 2. The generated PCR product was then cloned in the antisense orientation into the SmaI site of pTV00 (Ratcliff et al., 2001). pBYLNtArf1. The coding sequence of Arf1 was amplified from cDNA derived from N. tabacum BY-2 (NtArf1) RNA using primers 3 and 4. The amplified DNA was digested with AscI and inserted into pBYL2 (Mine et al., 2010b) that had been cut with the same restriction enzyme. pColdGST-NtArf1. The sequence of NtArf1 was amplified from pBYLNtArf1 using primers 5 and 6. The amplifiedDNAwas digested with KpnI and inserted into pColdGST (Mine et al., 2010a) that had been cut with the same restriction enzyme. pColdNtArf1-FLAG. PCR fragments from pBYLNtArf1 were amplified using primers 7 and 8 and primers 9 and 10. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 7 and 10. The amplified fragment was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35 (Takeda et al., 2005) to generate pUBNtArf1-FLAG. PCR fragments from pUBNtArf1-FLAG were amplified using primers 5 and 10. The amplified DNA was digested with KpnI and inserted into the corresponding region of pCOLDI (TaKaRa Bio Inc.). pBICNtArf1-myc:nYFP. The sequence of NtArf1 was amplified from pBYLNtArf1 using primers 7 and 12. The sequence of the myc-tagged N-terminal half of YFP was amplified from pBICmyc:nYFP (Mine et al., 2012) using primers 13 and 14. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 7 and 14. The amplified fragment was digested with BamHI and inserted into the corresponding region of pBICP35 (Takeda et al., 2005).

49 pBYLAtArf1. The coding sequence of Arf1 was amplified from cDNA derived from A. thaliana (AtArf1) RNA using primers 15 and 16. The amplified DNA was digested with AscI and inserted into pBYL2 that had been cut with the same restriction enzyme. pUBAtArf1. The sequence of AtArf1 was amplified from pBYLAtArf1 using primers 17 and 18. The amplified DNA was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35. pUBAtArf1-GFP. The sequence of AtArf1 was amplified from pUBAtArf1 using primers 19 and 20. The sequence of a green fluorescent protein (GFP) gene was amplified from pUBsGFP (where sGFP is synthetic GFP) (Kaido et al., 2011) using primers 10 and 21. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 10 and 19. The amplifiedDNAwas digested with StuI and KpnI and inserted into the corresponding region of pUBP35. pUBAtArf1-T31N. PCR fragments from pBYLAtArf1 were amplified using primers 17 and 22 and primers 18 and 23. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 17 and 18. The amplified fragment was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35. pUBAtArf1-Q71L. PCR fragments from pBYLAtArf1 were amplified using primers 17 and 24 and primers 18 and 25. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 17 and 18. The amplified fragment was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35. pBYLAtSar1. The coding sequence of Sar1 was amplified from cDNA derived from A. thaliana (AtSar1) RNA using primers 26 and 27. The amplified DNA was digested with AscI and inserted into pBYL2 that had been cut with the same restriction enzyme. pUBAtSar1. The sequence of AtSar1 was amplified from pBYLAtSar1 using primers 28 and 29. The amplified DNA was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35.

50 pUBAtSar1-GFP. The sequence of AtSar1 was amplified from pUBAtSar1 using primers 19 and 30. The sequence of GFP was amplified from pUBsGFP using primers 10 and 31. Then, a recombinantPCRproduct was amplified from the mixture of these fragments using primers 10 and 19. The amplified DNA was digested with StuI and KpnI, and inserted into the corresponding region of pUBP35. pUBAtSar1-H74L. PCR fragments from pBYLAtSar1 were amplified using primers 28 and 32 and primers 29 and 33. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 28 and 29. The amplified fragment was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35. pUBAtErd2-GFP. The sequence of C-terminally GFP-fused AtErd2 was amplified from pMT121-AtErd2-sGFP (Takeuchi et al., 2000) (generous gift from Akihiko Nakono) using primers 10 and 34. The amplified DNA was digested with StuI and KpnI, and inserted into the corresponding region of pUBP35. pUBp27-mCherry. A PCR fragment from pUBp27 (Takeda et al., 2005) was amplified using primers 19 and 35, and a PCR fragment from pBICER-mCherry (Kaido et al., 2011) was amplified using primers 10 and 36. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 10 and 19. The amplified fragment was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35. pUBp88-mCherry. A PCR fragment from pUBp88 (Takeda et al., 2005) was amplified using primers 37 and 38, and a PCR fragment from pUBp27-mCherry was amplified using primers 10 and 39. Then, a recombinant PCR product was amplified from the mixture of these fragments using primers 10 and 37. The amplified fragment was digested with XhoI and KpnI and inserted into the corresponding region of pUBp88. pUBER-GFP. A PCR fragment from pUCmGFP5-ER (Carette et al., 2000) (generous gift from Joan Wellink) was amplified using primers 40 and 41, and a PCR fragment from pUBsGFP was amplified using primers 42 and 43. Then, a recombinant PCR

51 product was amplified from the mixture of these fragments using primers 40 and 43. The amplified fragment was digested with BamHI and KpnI and inserted into the corresponding region of pUBP35.

Identification of Arf1. Protein identification based on in-gel digestion and liquid chromatography-tandem mass spectrometry (LC-MS/MS) was carried out as described previously (Taniguchi et al., 2010). Data obtained using a hybrid Fourier transform-ion cyclotron resonance (FT-ICR) mass spectrometer (LTQ-FT; Thermo, San Jose, CA) were processed using the Mascot Distiller software (version 2.3.2; Matrix Science, United Kingdom), and the peak lists obtained were used to search the NCBI nonredundant protein database (NCBInr 20120416) using the Mascot search engine (version 2.3; Matrix Science, United Kingdom). The taxonomy was restricted to Viridiplantae (green plants). The mass tolerances were set to ± 2 ppm for MS and ± 0.25 Da for MS/MS analyses.

Antibodies. Rabbit anti-p27 antiserum (Takeda et al., 2005) was used at 1:100 for immunofluorescence labeling. Mouse monoclonal antibodies were used at the following dilutions: for immunoblotting, anti-FLAG M2 antibody (F3165; Sigma-Aldrich) at 1:10,000 and, for immunofluorescence labeling, anti-bromodeoxyuridine (BrdU) antibody (B2531; Sigma-Aldrich) at 1:100. A rat anti-HA antibody (1867423; Roche) was used at 1:2,000 for immunoblotting. The secondary antibodies were goat anti-rabbit conjugated to Alexa Fluor 488 (A-11008; Molecular Probes) at 1:100 and goat anti-mouse conjugated to Alexa Fluor 594 (A-21125; Molecular Probes) at 1:100 for immunofluorescence labeling. For immunoblotting, a horseradish peroxidase-conjugated anti-mouse IgG antibody (074-1806; KPL) and alkaline phosphatase-conjugated anti-rat IgG antibody (sc-2021; Santa Cruz Biotechnology Inc.) were used to visualize the antigen-antibodycomplexes.

52

Expression and purification of recombinant proteins. The expression and purification of the recombinant N-terminally 6 $ His- and C-terminally FLAG-tagged p27 protein were performed as described previously (Mine et al., 2010a). For expression of NtArf1, E. coli strain BL21 (DE3) transformed with pColdNtArf1-FLAG was grown overnight at 37°C in Luria Broth (LB) medium containing ampicillin (50 µg/ml). The overnight culture (2 ml) was added to 100 ml of LB medium containing ampicillin (50 µg/ml). After the culture was incubated for 1 h at 37°C and subsequently for 30 min at 15°C, protein expression was induced by the addition of 0.3 mM isopropyl-%-D-thiogalactopyranoside (IPTG), followed by incubation at 15°C for 24 h. The induced cells were harvested by centrifugation at 5,000 ! g for 5 min, resuspended in 1 ml of His buffer (100 mM HEPES, pH 7.5, 300 mM NaCl) supplemented with 30 mM imidazole, and sonicated on ice. Subsequently, Triton X-100 was added at a final concentration of 0.5% and centrifuged at 21,000 ! g at 4°C for 10 min. The supernatant was added to the 50-µl bed volume of equilibrated Ninitrilotriacetic acid (NTA)-agarose (Qiagen, Hilden, Germany) and incubated at 4°C for 1 h with gentle rotation. The resin was washed three times with 1 ml of His buffer supplemented with 30 mM imidazole and eluted with TBS buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 2 mM dithiothreitol [DTT]) containing 15% glycerol and 250 mM imidazole. The concentration of purified protein was measured using a Coomassie Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA). The purified protein was subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and visualized with Coomassie brilliant blue R-250 to check its purity.

GST pulldown assay. E. coli BL21 (DE3) transformed with plasmids containing the prefix pColdGST was grown overnight at 37°C in LB medium containing ampicillin (50 µg/ml). The overnight cultures of the transformed E. coli were diluted 1:50 in LB medium

53 containing ampicillin (50 µg/ml). After the cultures were incubated at 37°C for 1.5 h and subsequently at 15°C for 30 min, protein expression was induced by addition of 0.3 mM IPTG. The cells were cultured for 2 h. The induced cells were harvested by centrifugation at 5,000 ! g for 5 min. Cells collected from 5 ml of medium were resuspended in 500 µl of TBS buffer and sonicated on ice to disrupt the cells. After sonication, Triton X-100 was added at a final concentration of 0.5% and centrifuged at 21,000 ! g at 4°C for 10 min. The supernatant was added to a 12.5-µl bed volume of equilibrated glutathione-Sepharose 4B (GE Healthcare, Little Chalfont, Buckinghamshire, United Kingdom) and incubated at 4°C for 1 h with gentle rotation. The resin was washed three times with 1 ml of TBS buffer supplemented with 0.5% Triton X-100. After the washing step, the resin was incubated at 4°C for 2 h in 200 µl of TBS buffer containing 1 µg of His-p27-FLAG or His-NtArf1-FLAG. After incubation, the resin was washed three times with 500 µl of TBS buffer. The bound proteins were eluted by the addition of Laemmli sample buffer (Laemmli. 1970), followed by incubation at 95°C for 3 min. Protein samples were subjected to immunoblotting using an anti-FLAG antibody.

RNA preparation. RCNMV RNA1 and RNA2 were transcribed from SmaI-linearized pUCR1 and pRC2|G, respectively, using T7 RNA polymerase (TaKaRa Bio, Inc.). BMV RNAs were transcribed from EcoRI-linearized pB plasmids using T7 RNA polymerase and capped with a ScriptCapm7G capping system (Epicentre Biotechnology). R1-Luc-R1 was synthesized from a SmaI-linearized plasmid. Luc mRNA and R-Luc mRNA were transcribed from EcoRI-linearized pLUCpA60 and pSP64-RLUC, respectively, and capped using the ScriptCapm7G capping system. All transcripts were purified with a Sephadex G-50 fine column (Amersham Pharmacia Biotech). RNA concentration was determined spectrophotometrically, and its integrity was verified by agarose gel electrophoresis.

54

Agrobacterium infiltration. The plasmids containing the prefixes pBIC and pTV were introduced by electroporation into Agrobacterium tumefaciens GV3101 (pMP90) and A. tumefaciens GV3101 (pSoup), respectively. Agrobacterium suspensions were mixed at final optical density at 600 nm (OD600) of 0.2 each for BiFC experiments and an OD600 of 0.5 each for gene silencing experiments. The mixture was infiltrated into N. benthamiana leaves as described previously (Mine et al., 2012).

BiFC experiment. Appropriate combinations of YFP fragment-fused or fluorescent protein-fused proteins were expressed in N. benthamiana leaves by Agrobacterium infiltration. The fluorescence of YFP and mCherry was visualized at 3 days postinfiltration (dpi).

Confocal microscopy. Tobacco BY-2 protoplasts were inoculated with plasmids expressing fluorescent protein-fused proteins (10 µg each) and incubated at 17°C for 16 h as described previously (Hyodo et al., 2011). The fluorescence of GFP and mCherry was visualized using an Olympus FluoView FV500 confocal microscope (Olympus Optical Co., Tokyo, Japan) as described previously (Kaido et al., 2009). To test the effect of BFA, the inoculated protoplasts were incubated with dimethyl sulfoxide (DMSO; Sigma-Aldrich) or 10 µg/ml BFA (Wako, Osaka, Japan) for an additional 2 h before observation. All images shown are from a 1-µm single optical section and were processed using Adobe Photoshop CS3 software.

Silencing of Arf1 in N. benthamiana plants. Appropriate combinations of silencing vectors were expressed via Agrobacterium infiltration in 3- to 4-week-old N. benthamiana plants as described previously (Ratcliff

55 et al., 2001). At 7 days postinfiltration (dpi), the leaves located above the infiltrated leaves were inoculated with in vitro transcribed RNA1 and RNA2 (1 µg each). At 2 days after inoculation, three inoculated leaves from three different plants infected with same inoculum were pooled, and total RNA was extracted using PureLink (Invitrogen), treated with RQ1 RNase-free DNase (Promega, Madison, WI), purified by phenol-chloroform and chloroform extractions, and precipitated with ethanol. Viral RNAs were detected by Northern blotting, as described previously (Mine et al., 2012). The mRNA levels of NbArf1 were examined by RT-PCR using primers 44 and 45. As a control to show the equal amounts of cDNA templates in each reaction mixture, the ribulose 1,5-biphosphate carboxylase small subunit gene (RbcS), a gene that is constitutively expressed, was amplified by RT-PCR using primers 46 and 47.

Replication assay. Tobacco BY-2 protoplasts were inoculated with RCNMVRNA1 (1.5 µg) and RNA2 (0.5 µg) and incubated with 10 µg/ml BFA at 17°C for 16 h. For the BMV replication assay, tobacco BY-2 protoplasts inoculated with BMV RNA1, RNA2, and RNA3 (1.5 µg each) were incubated with 10 µg/ml BFA at 17°C or 22°C for 20 h. For transient expression of Arf1, tobacco BY-2 protoplasts inoculated with plasmids expressing RNA1 and RNA2 (1 µg each), together with plasmids expressing Arf1 or its GTP- or GDP-fixed mutants (18 µg each), were incubated at 17°C for 24 h. Total RNA was extracted and subjected to Northern blotting, as described previously (Hyodo et al., 2011). Each experiment was repeated at least three times using different batches of protoplasts.

Luciferase assay. Tobacco BY-2 protoplasts inoculated with reporter RNAs were incubated with 10 µg/ml BFA at 17°C for 6 h. Luciferase assays were performed using a Dual-Luciferase Assay System (Promega, Madison, WI), as described previously (Mizumoto et al.,

56 2003). Each experiment was repeated at least three times using different batches of protoplasts.

BN-PAGE analyses. Plasmids expressing p27-HA and p88-HA were transfected to BY-2 protoplasts in the presence of RNA2. After 16 h of incubation at 17°C, protoplasts were harvested and resuspended in TR buffer (Komoda et al., 2004) supplemented with 1% Triton X-100. Subsequently, protoplasts were disrupted freezing and thawing and centrifuged at 21,000 ! g at 4°C for 10 min to remove cell debris and unbroken cells. The supernatants were subjected to Blue native (BN)- and SDS-PAGE followed by immunoblotting using an anti-HA antibody.

Coimmunopurification. Plasmids expressing p27-HA and p88-HA were transfected with either p27 or p27-FLAG into BY-2 protoplasts in the presence of RNA2. After 16 h of incubation at 17°C, protoplasts were harvested and resuspended in TR buffer supplemented with 1% Triton X-100. Subsequently, protoplasts were disrupted by freezing and thawing and centrifuged at 21,000 ! g at 4°C for 10 min to remove cell debris and unbroken cells. The supernatants were subjected to 12.5 µl of bed volume of ANTI-FLAG M2-Agarose Affinity Gel (Sigma-Aldrich) and incubated for 2 h with gentle mixing at 4°C. The resin was washed two times with 200 µl of TR buffer supplemented with 0.5% Triton X-100. The bound proteins were eluted by addition of Laemmli sample buffer, followed by incubation at 95°C for 3 min. Protein samples were subjected to SDS-PAGE, followed by immunoblotting using the appropriate antibodies.

Fractionation assay. Plasmids expressing p27-HA and p88-HA were transfected into BY-2 protoplasts in the presence of RNA2. After 16 h of incubation at 17°C, protoplasts were harvested and

57 resuspended in 500 µl of TR buffer. Subsequently, protoplasts were disrupted by freezing and thawing and centrifuged at 4,000 ! g at 4°C for 10 min. The cell extracts obtained were further centrifuged at 21,000 ! g at 4°C for 10 min to separate the supernatant and pellet fractions. The pellet fraction was resuspended in 50 µl of TR buffer supplemented with 0.5% Triton X-100. Aliquots of each fraction were subjected to immunoblotting using an anti-HA antibody.

In vivo RNA labeling. Tobacco BY-2 protoplasts (approximately 3 ! 105 cells) were inoculated with RNA1 (1.5 µg) and RNA2 (0.5 µg), as described previously (Hyodo et al., 2011). After 14 h, actinomycin D (Sigma-Aldrich) (10 µg/ml) was added, and protoplasts were incubated for 1 h to block RNA transcription from cellular DNA-dependent RNA polymerases. Subsequently, 2mM 5-bromouridine 5'-triphosphate (BrUTP; Sigma-Aldrich) was added, and protoplasts were incubated for an additional 3 h. Protoplasts were placed on a cover slide pretreated with poly-L-lysine. The reaction was stopped by the addition of PHEM buffer (60 mM PIPES [piperazine-N,N'-bis(2-ethanesulfonic acid)], 25 mM HEPES, 2 mM MgCl2, 5 mM EGTA, pH 6.9) containing 3% formaldehyde and 2.5% DMSO. After 15 min of incubation, the slide was washed three times with PHEM buffer and incubated in ice-cold methanol for 10 min. The samples were incubated with the primary antibodies for 1 h and washed three times with PHEM buffer. Then, they were incubated with Alexa Fluor 488- and 594-conjugated secondary antibodies for 1 h and washed three times with PHEM buffer.

58 Table II-1. List of primers used in this study.

Sequence

#1 CTGTGCTGCTTGTTTTTGCT

#2 CTTCGTTTACAAATTTATG

#3 GGCGCGCCATGGGGCTGTCTTTCGGCAAACTTTTCAGTCG

#4 GGCGCGCCCTATGCCTTGTTTGAAATATTGTTCGAAAGCC

#5 CGGGGTACCATGGGGCTGTCTTTCGGCAAACTTTTCAGTCGCC

#6 CGGGGTACCCTATGCCTTGTTTGAAATATTGTTCGAAAGCC

#7 AGAGGCCTACGGGGATCCAAGGAGATATAACAATGGGGCTGTCT

#8 CTACTTGTCATCGTCGTCCTTGTAATCTGCCTTGTTTGAAATAT

#9 GATTACAAGGACGACGATGACAAGTAGTAAGAATTCCGGGTACC

#10 CAGGAAACAGCTATGACCATG

#11 GCTCGGTACCATGGGGCTGTCTTTCGGCAAACTTTTCAGTCGCC

#12 AGCTTCTGCTCCATGCCCCCTGCCTTGTTTGAAATATTGTTCGA

#13 TTTCAAACAAGGCAGGGGGCATGGAGCAGAAGCTGATCAGCGAG

#14 CGGGGATCCCCGTAGGCCTCTAGGCCATGATATAGACGTTGTGG

#15 GGCGCGCCATGGGGTTGTCATTCGGAAAGTTGTTCAGC

#16 GGCGCGCCCTATGCCTTGCTTGCGATGTTGTTGGAGAGCC

#17 GGGGATCCGATGGGGTTGTCATTCGGAAAGTTGTTCAGCAGGC

#18 GGGGTACCCGGAATTCTTACTATGCCTTGCTTGCGATGTTGTTG

#19 AAGGGATGACGCACAATCCC

#20 TGATGATGATGATGATGCATTGCCTTGCTTGCGATGTTGTTGGA

#21 ACAACATCGCAAGCAAGGCAATGCATCATCATCATCATCATGTG

#22 AGCTTGTAGAGGATAGTGTTCTTACCAGCAGCATCGAGACCAAC

#23 GTCTCGATGCTGCTGGTAAGAACACTATCCTCTACAAGCTCAAA

#24 AATGGACGGATCTTGTCTAGACCCCCAACATCCCACACGGTGAA

#25 CCGTGTGGGATGTTGGGGGTCTAGACAAGATCCGTCCATTGTGG

#26 GGCGCGCCATGTTCTTGTTCGATTGGTTCTACGG

59 #27 GGCGCGCCTTAGTTGATGTACTGAGAGAGCCATTTGAATCC

#28 GGGGATCCGATGTTCTTGTTCGATTGGTTCTACGGAATCTTAGC

#29 GGGGTACCCGGAATTCTTATTAGTTGATGTACTGAGAGAGCCAT

#30 TGATGATGATGATGATGCATGTTGATGTACTGAGAGAGCCATTT

#31 GGCTCTCTCAGTACATCAACATGCATCATCATCATCATCATGTG

#32 ACCCTACGAGCAATCTGAAGACCACCCAAATCAAAAGCCTTGAA

#33 AGGCTTTTGATTTGGGTGGTCTTCAGATTGCTCGTAGGGTTTGG

#34 AGAGGCCTACGGGGGGATCCGATGAATATCTTTAGATTTGCTGG

#35 CCCTTGCTCACCATGCCCCCAAAATCCTCAAGGGATTTGAACCC

#36 CCCTTGAGGATTTTGGGGGCATGGTGAGCAAGGGCGAGGAGGAT

#37 CCTGTCGATGTACTCGAGAAGGTGGCGTTT

#38 TCCTCGCCCTTGCTCACCATGCCCCCTCGGGCTTTGATTAGATC

#39 TAATCAAAGCCCGAGGGGGCATGGTGAGCAAGGGCGAGGAGGAT

#40 CGGGATCCAAGGAGATATAACAATG

#41 CTCCTCGCCCTTGCTCACGAATTCGGCCGAGGATAA

#42 TTATCCTCGGCCGAATTCGTGAGCAAGGGCGAGGAG

#43 GGGGTACCTTAAAGCTCATCATGCTTGTACAGCTCGTCCAT

#44 AATGACAGAGACCGTGTTGTTGA

#45 ACAGCATCCCGAAGCTCATC

#46 CCTCTGCAGTTGCCACC

#47 CCTGTGGGTATGCCTTCTTC

60 Table II-2. The host proteins co-purificated with p27-FLAG.

name

gi | 1703374 ADP-ribosylation factor 1

gi | 10798648 purative DNAJ protein

gi | 175363751 elongation factor 1 gamma-like protein

gi | 120661 Glyceraldehyde-3-phosphate dehydrogenase A

gi | 120665 Glyceraldehyde-3-phosphate dehydrogenase B

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Fig. II-1. p27 interacts with Arf1 in vitro and in vivo. (A) Glutathione resin-bound GST-fused Arf1 (GST-Arf1) was incubated with the purified recombinant His-p27-FLAG. After a washing step, pulled down complexes were subjected to SDS-PAGE and analyzed by Western blotting (Wb) using an anti-FLAG antibody. After detection, the proteins separated on the membrane were visualized using Ponceau S staining. (B) SDS-PAGE analyses of purified His-Arf1-FLAG expressed in E. coli. The purified protein (1 µg) was visualized using Coomassie brilliant blue (CBB) staining (top panel) and analyzed by immunoblotting using an anti-FLAG antibody (bottom panel). (C) Schematic representation of the deleted derivatives of

GST-p27. The deletions are represented as dotted lines. The locations of known functional domains in wild-type p27 are depicted. aa, amino acids. (D)

Glutathione resin-bound GST-p27 or its derivatives were incubated with purified recombinant His-Arf1-FLAG. After a washing step, pulled down complexes were subjected to SDS-PAGE and analyzed by immunoblotting using an anti-FLAG antibody. After detection, the proteins separated on the membrane were visualized using Ponceau S staining. (E) Bimolecular fluorescence complementation analyses of the interactions between p27 and

Arf1. p27 fused to the C-terminal half of YFP, at the C terminus (p27-cYFP), was expressed together with Arf1 fused to the other half of YFP, at theCterminus (Arf1-nYFP), in the presence of ER-mCherry and of an RNA-silencing suppressor of the TBSV, p19, in N. benthamiana leaves via

Agrobacterium infiltration. The fluorescence of the reconstructed YFP and ER-mCherry in all agroinfiltrated leaves was visualized using confocal microscopy 3 days after agroinfiltration. The panels on the right show the merging of mCherry and YFP (yellow). Scale bar, 20 µm.

62

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Fig. II-2. Colocalization of Arf1 with RCNMV replicase proteins. A plasmid expressing C-terminally

GFP-fused Arf1 (Arf1-GFP) was cotransfected with plasmids expressing C-terminally mCherry-fused p27 (p27-mCherry) or with an empty vector into tobacco BY-2 protoplasts. Fluorescence was observed after 16 h of incubation. (A) In the absence of p27-mCherry, Arf1-GFP fluorescence was observed as small punctate structures throughout the cell. (B) Arf1-GFP fluorescence was observed as large aggregate structures in the presence of p27-mCherry. (C) Arf1-GFP fluorescence in the presence of p88-mCherry.

Fluorescence was visualized by confocal microscopy. The merging of green and red fluorescence is shown in yellow. Scale bar, 5 µm.

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Fig. II-3. (A) Knockdown of Arf1 mRNA levels via gene silencing inhibits the accumulation of RCNMV RNAs in N. benthamiana plants. The tobacco rattle virus (TRV) vector harboring a partial fragment of N. benthamiana Arf1 (TRV:NbArf1) was expressed in N. benthamiana via

Agrobacterium infiltration. The empty TRV vector (TRV:00) was used as a control. The newly developed leaves were inoculated withRCNMVRNA1 and RNA2 7 days after agroinfiltration. Total RNA was extracted from the mixture of three independent inoculated leaves 2 days after inoculation.

Accumulation of RCNMV RNAs was analyzed by Northern blotting. Ethidium bromide-stained ribosomal RNAs (rRNAs) are shown below the

Northern blots, as loading controls. Arf1 mRNA levels were assessed by RT-PCR using primers that allow the amplification of the region of Arf1 that is not present in TRV:NbArf1. RT-PCR results for the RbcS gene demonstrated that equal amounts of total RNA were used for the RT and showed equivalent efficiency of the RT reaction in the samples. (B) An inhibitor of Arf1 impairs RCNMV RNA replication in a single cell. Tobacco BY-2 protoplasts were inoculated with in vitro transcribed RNA1 and RNA2. The inoculated protoplasts were incubated at 17°C for 16 h in the presence of

10 µg/ml BFA. (C) An inhibitor of Arf1 does not impair BMV RNA replication in a single cell. Tobacco BY-2 protoplasts were inoculated with in vitro transcribed BMV RNAs. The inoculated protoplasts were incubated at 17°C for 20 h in the presence of 10 µg/ml BFA. (D) Dominant negative mutants of Arf1 inhibit RCNMV replication. Tobacco BY-2 protoplasts were transfected with plasmids expressing either wild-type Arf1, Arf1-T31N, or Arf1-Q71L together with pUBRC1 and pUBRC2, which encode RNA1 and RNA2, respectively, under the control of the cauliflower mosaic virus

35S promoter. The inoculated protoplasts were incubated at 17°C for 24 h. Total RNA was analyzed by Northern blotting. Ethidium bromide-stained rRNAs were used as loading controls and are shown below the Northern blots. The accumulated levels of RCNMV or BMV RNAs from three separate experiments were quantified using the Image Gauge program and were plotted in the graphs. The error bars indicate standard deviations.

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Fig. II-4. (A) Effects of BFA on RCNMV translation. Tobacco BY-2 protoplasts were inoculated with R1-Luc-R1 and cap-Luc-pA60 in the presence of 10 µg/ml BFA. The luciferase activities of R1-Luc-R1 and cap-Luc-pA60 in the absence of inhibitor were defined as 100%. The error bars indicate standard deviations. (B) Effects of BFA on the accumulation of the 480-kDa replicase complex. RCNMV RNA2 was cotransfected with plasmids expressing p27-HA and p88-HA in BY-2 protoplasts in the presence or absence of 10 µg/ml BFA. After 16 h of incubation, protoplasts were harvested and disrupted by freezing and thawing. Total proteins were extracted from the cell extracts obtained and subjected to BN- and SDS-PAGE analyses, followed by immunoblotting using an anti HA-antibody. Ponceau S staining served as a loading control. (C) BY-2 protoplasts were inoculated with plasmids expressing either p27 or p27-FLAG together with RNA2 and plasmids expressing HA-tagged viral replication proteins in the presence or absence of 10 µg/ml BFA. After 16 h of incubation, protoplasts were harvested and disrupted by freezing and thawing. Subsequently, the cell extracts were subjected to immunoprecipitation using an anti-FLAG antibody. Five percent of the total fraction (lanes 1 to 3) and the eluted fraction (lanes 4 to 6) were subjected to SDS-PAGE and analyzed by immunoblotting (IB) using anti-HA and anti-FLAG antibodies. *, Nonspecific IgG heavy chain.

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Fig. II-6. (A) Dominant negative mutants of Sar1 inhibitRCNMVreplication. Tobacco BY-2 protoplasts were transfected with plasmids expressing either wild-type Sar1 or Sar1-H74L together with pUBRC1 and pUBRC2. The inoculated protoplasts were incubated at 17°C for 24 h. Total RNA was analyzed by Northern blotting. Ethidium bromide-stained rRNAs were used as loading controls and are shown below the Northern blots. The accumulation levels of RCNMV RNAs from three separate experiments were quantified using the Image Gauge program and were plotted in the graphs. The error bars indicate standard deviations. (B and C) Colocalization of Sar1 with p27. A plasmid expressing C-terminally GFP-fused Sar1 (Sar1-GFP) was cotransfected with either the empty vector or a plasmid expressing C-terminally mCherry-fused p27 (p27-mCherry) into tobacco BY-2 protoplasts. Fluorescence was observed after 16 h of incubation. (B) Localization of Sar1-GFP in the absence of p27-mCherry. (C) Localization of Sar1-GFP in the presence of p27-mCherry. Fluorescence was visualized by confocal microscopy. The merging of green and red fluorescence is shown in yellow. Scale bar, 5 µm.

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Fig. II-7. Colocalization of incorporated BrUTP with RCNMV replication proteins. Representative images from mock-inoculated (upper row) or RCNMVinfected (lower row) BY-2 protoplasts incubated for 14 h postinoculation, incubated for an additional 1 h with actinomycin D, labeled with BrUTP, fixed, and processed for double-labeled immunofluorescence using antibodies that recognize p27 (green) and incorporated BrUTP (red). Scale bar, 10 µm.

68 Chapter III Functional analysis of phospholipase D and phosphatidic acid in a plant RNA virus replication

Introduction

Positive-strand RNA [(+)RNA] viruses have limited coding capacity, however, these viruses efficiently replicate in infected cells owing to their ability to utilize host-derived proteins, membranes, lipids, and metabolites. All characterized eukaryotic (+)RNA viruses replicate their genomes using the viral replication complexes (VRCs), which contain multiple viral and host components on intracellular membranes (Miller and Krijnse-Locker, 2008; Nagy and Pogany, 2012; den Boon and Ahlquist, 2010). A growing number of host proteins that affect viral RNA replication have been identified by genome-wide and proteomics analysis in several animal and plant viruses (Cherry et al., 2005 and 2006; Berger et al., 2009; Tai et al., 2009; Reiss et al., 2011; Panavas et al., 2005; Jiang et al., 2006; Li et al., 2009; Nawaz-ul-Rehman et al., 2012; Gancarz et al., 2011; Kushner et al., 2003). These host proteins are likely to play diverse roles during (+)RNA virus replication, including roles as transporters of intracellular localization of viral replication proteins and template RNAs; as protein chaperones that facilitate folding of viral proteins and assist the proper assembly of VRCs; as mediators of membrane remodeling; and as enzymes involved in lipid metabolism to modulate proper lipid microenvironments (Hyodo and Okuno, 2014 in press; Mine and Okuno, 2012; Nagy and Pogany, 2012). Red clover necrotic mosaic virus (RCNMV) is a (+)RNA plant virus and a member of the genus Dianthovirus in the family Tombusviridae. The genome of RCNMV consists of RNA1 and RNA2. RNA1 encodes p27 auxiliary replication protein, p88pol RNA-dependent RNA polymerase (RdRp), and the coat protein (Xiong and Lommel,

69 1989, Xiong et al., 1993b). RNA2 encodes the movement protein that is required for viral cell-to-cell movement (Kaido et al., 2009; Xiong et al., 1993a). p27, p88pol, and host proteins form the 480-kDa replicase complex, which is a key player in the viral RNA replication (Mine et al., 2010a). p27 and p88pol colocalize at the ER membrane (Turner et al., 2004, Kusumanegara et al., 2012), where RCNMV replication takes place (Hyodo et al., 2013). The RNA replication of RCNMV depends on host heat shock proteins (HSPs), including HSP70 and HSP90 (Mine et al., 2012), and ADP-ribosylation factor 1 (Arf1) (Hyodo et al., 2013). Arf1 is recruited from Golgi to RCNMV replication sites via p27. Pharmacological inhibition of Arf1 inhibits 480-kDa replicase complex formation and disrupts the p27-induced ER membrane alternation. It is known that mammalian and yeast Arf1 recruits and/or stimulates numerous numbers of its effector proteins, including coatomer, phosphatidylinositol 4 kinase III % (PI4KIII%), and phospholipase D (PLD) (reviewed in Donaldson and Jackson, 2011). These findings raise intriguing question of whether these Arf1 effector proteins also play a role in RCNMV RNA replication. Arf1 can directly activate mammalian PLD1 and PLD2, and PA generation resulting from Arf1-mediated PLD activation has been proposed to be associated with vesicle formation (reviewed in Jang et al., 2012). Moreover, Arf1 have been shown to interact with PA (Manifava et al., 2001). PLD hydrolyses structural phospholipids such as phosphatidylcholine (PC) and phosphatidylethanoamine (PE) to form phosphatidic acid (PA) and remaining headgroup. The 12 different PLD isoforms encoded in the Arabidopsis thaliana genome are classified into six groups (&, %, ', (, ), and *) based on sequence similarity and in vitro activity (Li et al., 2009). PLD*1 and *2 have N-terminal phox homology (PX) and pleckstrin homology (PH) domains and share high sequence similarities to two PX/PH-PLDs in mammals. The remaining PLDs contain the Ca2+-dependent phospholipid-binding C2 domain and are unique to plants. PA is normally present in small amounts (less than 1% of total phospholipids), but rapidly and transiently accumulates in lipid bilayers in response to different abiotic stresses such as abscisic

70 acid (ABA), dehydration, and salt and osmotic stress (Li et al., 2009; Testerink and Munnik, 2011; Hong et al., 2010). PA has been also shown to accumulate in response to several microbe-associated molecular patterns (MAMPs) in plant cells (van der Luit et al., 2000; Kirik and Mudgett, 2009; Yamaguchi et al., 2005; Suzuki et al., 2007). Moreover, specific bacterial and fungal pathogen effector proteins, such as Cladosporium fulvum Avr4 and Pseudomonas syringae AvrRpm1 and AvrRpt2 trigger PA response in their host cell (de Jong et al., 2004; Andersson et al., 2006). Analysis of plants deficient in or overexpressing specific PLDs in plants have provided evidence for different PLDs in specific physiological responses (Li et al., 2009). PLD%1 has been found as a negative regulator of salicylic acid (SA)-dependent resistance to Pseudomonas syringae, but a positive regulator of the jasmonic acid (JA)-dependent pathway and resistance to Botrytis cinerea (Zhao et al., 2013). PLD(, but not other PLDs, plays a positive role in nonhost resistance against Blumeria graminis f. sp. hordei (Pinosa et al., 2013). Overexpression of rice diacylgrycerol kinase, which catalyzes diacylgrycerol (DAG) to PA conversion, enhances resistance to tobacco mosaic virus and Phytophthola parasitica infection in tobacco (Zhang et al., 2008). In accordance with this, direct application of PA to leaves has been shown to induce pathogen-related gene expression and cell death (Park et al., 2004; Andersson et al., 2006). These findings indicate that PA is an important mediator within plant-pathogens interactions. In this study, by using two-step affinity purification and liquid chromatography-tandem mass spectrometry (LC/MS/MS) analysis, I identified Nicotiana benthamiana PLD& and PLD% as interaction partners of RCNMV replication proteins. Gene-silencing and pharmacological inhibition approaches showed that PLDs-derived PA played a positive role in viral RNA replication. In consistent with this, direct application of PA to virus-infected plant cells or plant-derived cell-flee systems enhanced the viral RNA replication. I found that the effects of exogenously applied PA on viral RNA replication were different between RCNMV genomic RNAs. I also showed that p27 is a PA-binding protein. RCNMV-infected plant leaves showed high

71 accumulation of PA, leading to the possibility that RCNMV hijacks host PA signaling pathway for successful RNA replication.

Results and Discussions

To identify putative host proteins associated with the RCNMV replicase complex, I expressed six-His/FLAG-tagged p27 and p88pol replication proteins together with RNA2 replication template via agroinfiltration in N. benthamiana plant. Two-days after infiltration (dai), I purified RCNMV replication proteins via two-step affinity purification as described in Materials and Methods. It is note that six-His/FLAG-tagged RCNMV replication proteins can support RNA2 replication in N. benthamiana plants (data not shown). The two-step affinity purified fraction was subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) to separate the proteins copurified with RCNMV replication proteins. Silver-staining analysis revealed that the presence of numerous numbers of proteins that were absent in the control fraction prepared from similar preparation from Agrobacterium-infiltrated N. benthamiana leaves expressing non-tagged p27 and p88 pol together with RNA2 (Fig. III-1). LC/MS/MS analysis of isolated proteins excised from gels led to the identification of numerous numbers of host proteins. These included previously identified host factors, HSP70 and HSP90 (Mine et al., 2012), and Arf1 (Hyodo et al., 2013). Sar1 was also identified in this analysis. Sar1 is a small GTPase implicated in the biogenesis of COPII vesicles at the ER exit sites (Hanton et al., 2008). Inhibition of COPII biogenesis by overexpression of Sar1 dominant-negative mutant (Sar1 H74L) severely inhibited RCNMV RNA replication (Hyodo et al., 2013). Moreover, colocalization of Sar1 and p27 at ER was observed (Hyodo et al., 2013). These observations, together with the possible interaction between Sar1 and RCNMV replication proteins, strongly suggest that Sar1 is positively utilized during viral RNA replication. Interestingly, several

72 known Arf1 effector proteins, namely coatomer subunits, clathrin heavy chain, and PLD& and PLD%, were also identified in this LC/MS/MS analysis. This suggested that RCNMV hijacks Arf1 pathway for successful viral proliferation. Other factors identified by LC/MS/MS analysis were listed in Table III-1. It is known that the activities of yeast and mammalian PLDs are stimulated by Arf1 (Brown et al., 1993) and Arf1 is an essential host factor in RCNMV RNA replication (Hyodo et al., 2013). These previous findings tempted me to investigate whether PLD& and PLD% are also required for RCNMV RNA replication. I tested whether PLD& and PLD% are required for infection of host plants with RCNMV. I downregulated PLD& or PLD% using Tobacco rattle virus (TRV)-based virus-induced gene silencing in N. benthamiana plants. A TRV vector harboring a partial fragment of NbPLD& (TRVNbPLD&) or NbPLD% (TRVNbPLD%) were expressed via Agrobacterium-mediated expression. An empty TRV vector (TRV:00) was expressed as a control. Newly developed leaves were inoculated with RCNMV RNA1 and RNA2 at 18 dai. Two days after RCNMV inoculation, three inoculated leaves from three different plants were harvested and mixed, and total RNA was extracted. Semiquantitative RT-PCR analyses confirmed the specific reduction of NbPLD& or NbPLD% mRNAs in plants infiltrated with TRV:NbPLD& or TRV:NbPLD%, respectively (Fig. III-2). Northern blot analyses showed that the accumulation of RCNMV RNA was dramatically reduced in NbPLD-silenced plants compared with control plants (Fig. III-2), suggesting that both PLD& and PLD% play a positive role in RCNMV infection.

To test the possible contribution of PLD-derived PA to viral RNA replication, I exploited the transphosphatidylation activity of PLD, which uses primary alcohols as substrates to form an artificial phosphatidyl alcohol. The preferential formation of this compound impairs PA production (Munnik et al., 1995). Thus, I tested the effect of n-butanol that inhibits PA formation by PLD on the RNA replication of RCNMV in

73 single plant cells. N. benthamiana protoplasts were inoculated with RCNMV RNA1 and RNA2 and incubated for 18 h in the presence of n-butanol or tert-butanol, an alcohol with no inhibitory effect on PA formation, and viral RNA accumulations were investigated by Northern blot analysis using ribonucleotide probes specifically recognizing RCNMV RNA1 and RNA2, respectively. Increasing n-butanol concentrations caused progressively reduction of the accumulations of viral RNAs (Fig. III-3A). In contrast, viral RNA accumulations for N. benthamiana protoplasts treated with tert-butanol was not significantly different compared with that of the water control (Fig. III-3A). Note that n-butanol did not affect the accumulation of rRNA (Fig. III-3A).

To further verify the importance of PA in viral RNA replication, PA was exogenously supplied to RCNMV-infected N. benthamiana protoplasts and viral RNA accumulations were examined by Northern blot analysis. Surprisingly, exogenously added PA enhanced the accumulation of RNA1 in a dose-dependent manner (up to 6-fold increase in 5 µM PA), whereas the effect of PA on the accumulation of RNA2 was negligible (Fig. III-3B). These results not only suggest that PLD-produced PA plays an important role in RCNMV RNA replication but also raise the possibility that requirements of PA for viral RNA replication differ between bipartite viral genomic RNAs.

Replication of RNA2 depends entirely on RNA1 because RNA2 does not encode any replication proteins. Therefore, it is possible that the reason of the negative effect of n-butanol on the accumulation of RNA2 is its negative impact on RNA1 replication. To investigate whether replication of RNA2 requires PLD-derived PA, I inoculated RNA2 together with plasmids expressing C-terminally HA-tagged p27 and p88pol into N. benthamiana protoplasts. Inoculated protoplasts were incubated for 18 h in the presence of n-butanol or control tert-butanol. Immunoblot analysis showed that n-butanol did not affect the accumulation of p27 (Fig. III-3C). However, the accumulation of RNA2 was

74 decreased by n-butanol in a dose-dependent manner (Fig. III-3C). These results indicated that PLD-derived PA is also required for the replication of RNA2 as in the case of RNA1. However, exogenously added PA did not affect the accumulation of RNA2 (Fig. III-3D). Together, these results strongly suggest that requirements of PA for RNA replication differ between bipartite viral genomic RNAs.

PA acts as a second messenger in signal transduction during multiple biotic and abiotic stress responses and plays multiple roles (Li et al., 2009; Testerink and Munnik, 2011; Hong et al., 2010). So, it is possible that the positive effect of PA on the replication of RCNMV may be due to its indirect contribution. To investigate whether PA has a direct role in viral RNA replication, I used BYL, an in vitro translation/replication system (Komada et al., 2004). BYL has been used successfully to recapitulate the negative-strand synthesis of RCNMV (An et al., 2010; Hyodo et al., 2011; Iwakawa et al., 2007, 2008, 2011; Mine et al., 2010a and b). Addition of n-butanol to BYL did not affect the accumulation of negative-strand RNAs (data not shown), implying that PLD activities are not essential for viral negative-strand RNA synthesis in this in vitro system. However, addition of PA into BYL stimulated the accumulation of negative-strand RNA1 (Fig. III-4), indicating that PA has positive roles in negative-strand RNA synthesis of RCNMV in a direct manner.

As PA directly stimulated the viral negative-strand synthesis in BYL in vitro system, I hypothesized that p27, the multifunctional RCNMV replication protein, has the affinity for PA. To investigate this possibility, I conducted a lipid overlay assay using bacterially expressed, purified C-terminally FLAG-tagged p27 (p27-FLAG). Purified p27-FLAG protein (Fig. III-5A) was incubated with phospholipids-spotted nitrocellurose membranes and p27-FLAG–phospholipids interaction was detected using anti-FLAG antibody. Lipid overlay assays revealed that p27 gave strong PA binding signals on the blot (Fig. III-5B). p27 exhibited weak binding to

75 phosphatidylinositol-4-phosphate [PtdIns(4)P], but negligible binding to other lipids, including PC, PE, PS, PtdIns, lysoPA, lysoPC, sphingosine-1-phosphate, and DAG (Fig. III-5B and C). These results indicate that p27 binds to PA. PA has been reported to bind to various proteins, including transcription factors, kinases, phosphatases, enzymes involved in central metabolism, and proteins involved in vesicular trafficking and cytoskeletal rearrangements (Li et al., 2009; Testerink and Munnik, 2011). It has been proposed that PA binding to proteins modulates the catalytic activity of target proteins, tethers proteins to the membranes, and promotes the formation and/or stability of protein complexes (Guo and Wang, 2012). Because p27 localizes at the ER membranes (Turner et al., 2004, Kusumanegara et al., 2012), PA may tether p27 to the ER membranes. Alternatively, PA-p27 interaction modulates the formation and/or activity of the RCNMV replicase complex. Defining the PA-binding domain of p27 replication protein will be helpful for understanding the roles of PA in viral RNA replication. Moreover, several PA-binding proteins also bind to other phospholipid and mediate signals to other phospholipid pathway (Li et al., 2009). Therefore, it is interesting to examine the impact of p27–PtdIns4P interaction on viral RNA replication in future study. It is noteworthy that PI4KIII% plays a positive role in Coxsackievirus B3 replication and that viral RdRP 3Dpol binds to PtdIns4P in vitro (Hsu et al., 2010).

I next measured PA accumulation during RCNMV infection. N. benthamiana leaves were infected with RCNMV via agroinfiltration. At 2 dai, lipids were extracted and PA accumulation was analysed by thin-layer chromatography. Comparing with empty vector-infiltrated control plant leaves, PA accumulation was about 4.3-fold higher in RCNMV-infected plant leaves (Fig. III-6A). However, accumulations of PLD& or PLD% transcripts were not significantly affected by RCNMV infection (Fig. III-6B). From these results, it may be possible that RCNMV replication proteins interact with and recruit PLD& and PLD% to VRC and stimulate the PLD activity for viral RNA replication. Alternatively, RCNMV-induced high PA accumulation is due to recognition

76 of viral components during plant defense responses. It is known that several MAMPs and bacterial or fungal proteins stimulate PA accumulations in plants and PA is an important second messenger in biotic stress responses (Li et al., 2009; Testerink and Munnik, 2011). Further studies are needed to reveal the impact of RCNMV replication proteins on PLDs localizations and activities.

Finally, I investigated the effects of PA on other viral RNA replication. To this end, I inoculated Brome mosaic virus (BMV), unrelated to RCNMV, into N. benthamiana protoplasts and examined the effects of n-butanol and PA on BMV replication by Northern blot analysis. Increasing n-butanol concentrations caused progressive reduction in the accumulation of BMV RNAs (Fig. III-7A), whereas exogenously added PA did not affect the accumulation of BMV RNAs (Fig. III-7B). These results suggested that PLD-derived PA is also important for BMV RNA replication.

In this study, I showed that PLD& and PLD% of N. benthamiana play an important role in RCNMV replication. RCNMV infection induced a high accumulation of PA in plant tissues, suggesting that RCNMV activates cellular PLD activities. Interestingly, yeast and mammalian Arf1 interacts with PA and directly activates PLDs (Brown et al., 1993; Manifava et al., 2001), and RCNMV recruits Arf1 from Golgi to the viral replication sites (Hyodo et al., 2013). These findings imply that RCNMV recruits Arf1 and its effector proteins, such as PLD, and efficiently uses these host proteins for viral RNA replication. However, it is unknown that plant specific C2-domain-containing PLDs are also Arf1 effector proteins. Further studies are needed to unveil the link between Arf1 and PLD in plants.

Several known cellular PA-binding proteins were also identified in this LC/MS/MS analysis (Table III-1). These include NADPH oxidase (Zhang et al., 2009), Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Kim et al., 2013; McLoughlin

77 et al., 2013), and SNF1-related kinase (SnRK) (McLoughlin et al., 2013). RCNMV-induced high PA accumulation may recruit these PA-binding proteins and affect the physiological changes of host plants.

Materials and Methods

Gene cloning and plasmid construction. pUCR1 (Takeda et al., 2005) and pRC2_G (Xiong and Lommel, 1991) are full-length cDNA clones of RNA1 and RNA2 of an RCNMV Australian strain, respectively. pB1TP3, pB2TP5, and pB3TP8 are full-length cDNA clones of RNA1, RNA2, and RNA3 of the BMV M1 strain, respectively (Janda et al., 1987) (generous gift from Paul Ahlquist). The constructs described previously used in this study include pBICp27 (Takeda et al., 2005), pBICp88 (Takeda et al., 2005), pBICR2 (Takeda et al., 2005), pBICR1R2 (Takeda et al., 2005), and pCOLDIp27-FLAG (Mine et al., 2010). pUC118 was purchased from TaKaRa Bio Inc. (Shiga, Japan). Escherichia coli DH5& was used for the construction of all plasmids. All plasmids constructed in this study were verified by sequencing. RNA extraction from Nicotiana benthamiana leave was performed using an RNeasy Plant Mini Kit (Qiagen, Hilden, Germany). Reverse transcription-PCR (RT-PCR) was catalyzed by Superscript III reverse transcriptase (Invitrogen) using oligo(dT) (Mine et al., 2012). pBICp27-His-FLAG. A PCR fragment from pBICp27 was amplified using primers 5'-CGGGGATCCATGGGTTTTATAAATCTTTCG-3' and 5'-CGGGGATCCCTACTTGTCATCGTCGTCCTTGTAATCATGATGATGATGATG ATGAAAATCCTCAAGGGATTTG-3'. The amplified fragment was digested with BamHI and inserted into the corresponding region of pBICP35. pBICp88-His-FLAG. A PCR fragment from pBICp88 was amplified using primers 5'- CCGGGTACCATGGGTTTTATAAATCTTTCG-3' and 5'-

78 CGGGGTACCTTACTTGTCATCGTCGTCCTTGTAATCATGATGATGATGATGA TGTCGGGCTTTGATTAGATCTTTG-3'. The amplified fragment was digested with KpnI and inserted into the corresponding region of pBICP35. pTVNbPLD#. A PLD& cDNA fragment was amplified from cDNA derived from N. benthamiana (NbPLD&) RNA using primers 5'-ATCCCCCGGGAGTTCCTTGTAGCTCTGTGACATCCCC-3' and 5'-GCAGCCCGGGTCAGCCAACATAAACCAGAGATCGATGGACGG-3'. The generated PCR product was then cloned in the antisense orientation into the XmaI site of pTV00 (Ratcliff et al., 2001). pTVNbPLD$. A PLD% cDNA fragment was amplified from cDNA derived from N. benthamiana (NbPLD%) RNA using primers 5'-CCCCCGGGTCGACTTTTCTGACTGAGTTCCTGAGGTGTAGCAGG-3' and 5'-AGCCCGGGTTGATACCAATGGAGATTGCTCTAAAAATTGCC-3'. The generated PCR product was then cloned in the antisense orientation into the XmaI site of pTV00.

Tandem affinity purification. Four week-old N. benthamiana plants were agroinfiltrated as described previously (Hyodo et al., 2013). At 2 dpi, total proteins were extracted from 6 g of leaves in 10 ml of Buffer A (50 mM HEPES, 150 mM NaCl, 0.1 % 2-mercaptoethanol, 0.5 % Triton X-100, 5 % Glycerol, pH 7.5) containing 30 mM imidazole and protease inhibitor cocktail (Roche). Aliquots were centrifuged at 21,000 g, 4
10 min to remove cell debris and supernatants were added to 400 µl of Ni-NTA beads (Quiagen) and incubated for 1 h at 4
with gentle mixing. Beads were washed 3 times with 1 ml of buffer A containing 100 mM imidazole. Bound proteins were eluted with 1 ml of buffer A containing 500 mM imidazole. Eluates were added to 50 µl of FLAG-beads (Sigma) and incubated for overnight at 4
with gentle mixing. Beads were washed 3 times with 1 ml of buffer A. Bound proteins were eluted with 300 µl of buffer A containing 150

79 µg/ml 3 $ FLAG peptides (Sigma). Eluates were concentrated by Acetone precipitation and dissolved in 1 $ NuPAGE sample Buffer (Invitrogen). Purified proteins were separated by SDS-PAGE (NuPAGE 3-12 % Bis-Tris gel: Invitrogen) and visualized by Silver staining (Wako, Japan). Proteins in excised gel pieces were subjected to digestion with typsin, LC-MS/MS analysis, and MASCOT searching as described previously (Mine et al., 2010b).

RNA preparation. RCNMV RNA1 and RNA2 were transcribed from SmaI-linearized pUCR1 and pRC2|G, respectively, using T7 RNA polymerase (TaKaRa Bio, Inc.). BMV RNAs were transcribed from EcoRI-linearized pB plasmids using T7 RNA polymerase and capped with a ScriptCapm7G capping system (Epicentre Biotechnology). All transcripts were purified with a Sephadex G-50 fine column (Amersham Pharmacia Biotech). RNA concentration was determined spectrophotometrically, and its integrity was verified by agarose gel electrophoresis.

Silencing of PLD! and PLDß in N. benthamiana plants. Appropriate combinations of silencing vectors were expressed via Agrobacterium infiltration in 3- to 4-week-old N. benthamiana plants as described previously (56). At 18 days postinfiltration (dpi), the leaves located above the infiltrated leaves were inoculated with in vitro transcribed RNA1 and RNA2 (500 ng each). At 2 days after inoculation, three inoculated leaves from three different plants infected with same inoculum were pooled, and total RNA was extracted using PureLink (Invitrogen), treated with RQ1 RNase-free DNase (Promega, Madison, WI), purified by phenol-chloroform and chloroform extractions, and precipitated with ethanol. Viral RNAs were detected by Northern blotting, as described previously (32). The mRNA levels of NbPLD" and NbPLDß were examined by RT-PCR using primer pairs 5'-TATCAAGGTAGAGGAGATAGGTGC-3' and

80 5'-TACATCATCTCCATCGTTCTCCTC-3', and 5'-GAAGGCTTCAAAGCGCCATG-3' and 5'-CTTAGGCAAGGGACATCAGC-3', respectively. As a control to show the equal amounts of cDNA templates in each reaction mixture, the ribulose 1,5-biphosphate carboxylase small subunit gene (RbcS), a gene that is constitutively expressed, was amplified by RT-PCR as described previously (Mine et al., 2012).

Protoplasts experiments. N. benthamiana protoplasts were inoculated with RCNMV RNA1 (1.5 µg) and RNA2 (0.5 µg) and incubated with various concentrations of n-butanol (Sigma-Aldrich), tert-butanol (Sigma-Aldrich), or phosphatidic acid (PA) (Soy-derived; Avanti Polar Lipid) at 20°C for 18 h. PA was dissolved in DMSO. Total RNA was extracted and subjected to Northern blotting, as described previously (Hyodo et al., 2013). Each experiment was repeated at least three times using different batches of protoplasts.

BYL experiments. The preparation of BYL was as described previously (Komoda et al., 2004; Iwakawa et al., 2007). The BYL translation/replication assay was performed essentially as described previously (Iwakawa et al., 2007). Briefly, 300 ng of RNA1 and 100 ng of RNA2 were added to 20 µL of BYL translation/replication mixture in the presence of various concentrations of PA. The mixture was incubated at 17 °C for 240 min. Aliquots of the reaction mixture were subjected to northern and immunoblotting analyses, as described previously (Iwakawa et al., 2007 and 2011).

Protein purification and lipid overlay assay. Protein expression in E. coli BL21 (DE3) and subsequent purification were done as described previously (Hyodo et al., 2013). Protein concentrations were measured by Bladfold assay. Protein purity was checked by SDS-PAGE and CBB staining.

81 Lipid overlay assay was carried out as recommended by manufacture protocol. Briefly, the membrane (PIP StripsTM or Membrane Lipid ArraysTM; Echelon Bioscience Inc.) was incubated in 3 % fatty acid free BSA in a mixture of phosphate-buffered saline and 0.1 % Tween 20 (PBST) for 1 h at room temperature (RT) and then incubated in same solution containing 500 ng of purified recombinant protein for 1 h at RT. After washing three times with PBST, the membrane was incubated with a mouse anti-FLAG antibody (1:10000; Sigma-Aldrich) for 1 h at RT, followed by three washes with PBST. An anti-mouse IgG conjugated with horseradish peroxidase (KPL; 1:10000) was used as secondary antibody. Binding of proteins to phospholipids was visualized by incubating with chemiluminescent substrate.

Lipid extraction and TLC. Four week-old N. benthamiana plants were infected with RCNMV via agroinfiltration. At 2 dpi, 0.33 g of infiltlated leaves were ground in liquid nitrogen and extracted in 900 µl of water. Total lipids were extracted by adding 3 ml CHCl3/CH3OH (2:1, v/v) to each samples. Samples were vortexed and centrifuged at 1690 $ g, 4
, 10 min. The organic phase was recovered and dried under nitrogen gas stream. Lipids were dissolved in 100 µl CHCl3/CH3OH (2:1, v/v). Then 5 µl of samples were analysed on thin-layer chromatography (TLC) plate (Merck, Gemany). The chromatography was performed using CHCl3/CH3OH/formic acid/asetic acid (12:6:0.6:0.4, v/v). Plates were air dried, soaked into 10 % CuSO4, and charred at 180  for 10 min to visualize lipids. qRT-PCR analysis Four week-old N. benthamiana plants were infected with RCNMV via agroinfiltration. Total RNA was extracted as described above. cDNA was synthesized by reverse transcriptase (TAKARA) and accumulations of transcripts were assessed by using primer as described above. UBQ3 was used as control.

82 Table III-1. List of host factors copurified with RCNMV replication proteins.

protein name

ADP ribosylation factor 1
cullin-associated NEDD8-dissociated protein 1-like

coatomer subunit alpha E3 ubiquitin protein ligase upl2

coatomer subunit beta E3 ubiquitin-protein ligase KEG-like

coatomer subunit gamma Ubiquitin carboxyl-terminal hydrolase

GNOM-like 1 protein spindle disassembly related protein CDC48

brefeldin A inhibited guanine nucleotide exchange factor1 SNF1 protein

phospholipase D alpha SNF1-related kinase

phospholipase D beta Tm-1 protein

clathrin heavy chain 2 calcium-dependent protein kinase 2

Sar1 protein phosphatase 2A

RabB N-alpha-acetyltransferase 16, NatA auxiliary subunit-like

RabE N-terminal acetyltransferase complex ARD1 subunit-like

HMG-CoA synthase Germin like protein

Heat shock protein 70 NADPH oxidase

Heat shock protein 90 Glyceraldehyde-3-phosphate dehydrogenase C

luminal-binding protein Glyceraldehyde-3-phosphate dehydrogenase A

T-complex protein 1 subunit epsilon-like

T-complex protein 1 subunit theta

T-complex protein 1 subunit zeta

T-complex protein 1, eta subunit

83 !"#$!%%$&'("!"#)*$!%%)*$&'("

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Fig. III-1. Identification of proteins copurified with RCNMV replication proteins. The solubilized fractions prepared from Agrobacterium-infiltrated leaves expressing p27 plus p88 plus RNA2 or p27-His-FLAG plus p88-His-FLAG plus RNA2 were subjected to affinity purification with Ni-NTA agarose beads, followed by anti-FLAG antibody-conjugated agarose beads. The affinity-purified fractions were subjected to SDS-PAGE and stained using MS-compatible silver staining. Protein bands of interests were excised, subjected to in-gel digestion, and analyzed by tandem mass spectrometry.

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Fig. III-2. Knockdown of PLD& and PLD% mRNAs levels via gene silencing inhibits the accumulation of RCNMV RNAs in N. benthamiana plants. The tobacco rattle virus (TRV) vector harboring a partial fragment of N. benthamiana PLD& and PLD% (TRV:NbPLD& and TRV:NbPLD%, respectively) were expressed in N. benthamiana via Agrobacterium infiltration. The empty TRV vector (TRV:00) was used as a control. The newly developed leaves were inoculated with RCNMV RNA1 and RNA2 at 18 days after agroinfiltration. Total RNA was extracted from the mixture of three independent inoculated leaves 2 days after virus inoculation. Accumulation of RCNMV RNAs was analyzed by Northern blotting. Ethidium bromide-stained ribosomal RNAs (rRNAs) are shown below the Northern blots, as loading controls. PLD& and PLD% mRNA levels were assessed by RT-PCR using primers that allow the amplification of the region of coding regions that are not present in TRV vectors. RT-PCR results for the RbcS gene demonstrated that equal amounts of total RNA were used for the RT and showed equivalent efficiency of the RT reaction in the samples.

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;(( ;(( Fig. III-3. (A) An inhibitor of PLDs impairs RCNMV RNA replication in a single cell. N. benthamiana protoplasts were inoculated with in vitro transcribed RNA1 and RNA2. The inoculated protoplasts were incubated at 20°C for 18 h in the presence of n-butanol.

(B) An exogenously supplied phosphatidic acid (PA) enhances RCNMV RNA replication. N. benthamiana protoplasts were inoculated with in vitro transcribed RNA1 and RNA2. The inoculated protoplasts were incubated at 20°C for 18 h in the presence of progressively increasing concentrations of PA. (C) An inhibitor of PLDs impairs replication of RNA2 in a single cell. N. benthamiana protoplasts were inoculated with in vitro transcribed RNA2 together with plasmids expressing p27-HA and p88-HA.

The inoculated protoplasts were incubated at 20°C for 18 h in the presence of n-butanol. (D) The effect of an exogenously supplied

PA on the replication of RNA2. N. benthamiana protoplasts were inoculated with in vitro transcribed RNA2 together with plasmids expressing p27-HA and p88-HA. The inoculated protoplasts were incubated at 20°C for 18 h in the presence of progressively increasing concentrations of PA. Total RNA was analyzed by Northern blotting using ribonucleotide probes that recognize specifically RCNMV RNA1 and RNA2, respectively. Ethidium bromide-stained rRNAs were used as loading controls and are shown below the Northern blots. For (C) and (D), total proteins were analysed by Immunoblotting using anti-HA antibody.

Coomassie brilliant blue (CBB) staining served as a loading control.

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Fig. III-4. PA enhances accumulations of negative-strand RNA1 in BYL. BYL was incubated for 4 h at 17°C with in vitro-synthesized RNA1 (300 ng) and RNA2 (100 ng) in the presence of various concentrations of PA. Accumulations of negative-strand RNA1 were detected by Northern blotting. Accumulations of p27 were analyzed by Western blotting with anti-p27 antisera. EtBr-stained rRNAs and Coomassie brilliant blue (CBB)-stained cellular proteins are shown as loading controls.

87 D M 2@L'4 8%&).9$#)./:;:%&$#%&'(/ ;=F: !"#$%&$#%&'()*)+,'+)* J5: !"#$%&$#%&$+&$-)./ 0(*1.#23<540= 5K: 0(*1.#23<640= 3K: 0(*1.# A)#:%=B:C!DE =J: 0(*1.#2340 0(*1.#25<640=

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Fig. III-5. p27 interacts with PA. (A) Purified p27-FLAG expressed in E. coli BL21 (DE3) was used for SDS-PAGE, followed by Coomassie brilliant blue (CBB) staining. (B and C) Lipid overlay analysis of various lipids with p27. Purified protein (500 ng) was used, followed by immunoblotting with anti-FLAG antibody.

88 # 9

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Fig. III-6. RCNMV stimulates the accumulation of PA in a plant tissue. (A) The extracted lipids from Agrobacterium-infiltrated leaves expressing empty vector or RCNMV RNA1 plus RNA2 were subjected to thin-layer chromatography and phospholipids were visualized by CuSO4 staining. (B) Expression of PLD& and PLD% in N. benthamiana plants infiltrated with Agrobacterium harboring empty vector (used as a control) or RCNMV RNA1 plus RNA2 for 2 days. The relative expression level of PLD& and PLD% was analyzed by quantitative PCR. Expression levels were normalized with respect to the UBQ3 gene. Data bars represent the mean (± SD) of three repeats.

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Fig. III-7. (A) An inhibitor of PLDs impairs BMV RNA replication in a single cell. N. benthamiana protoplasts were inoculated with in vitro transcribed BMV RNA1, RNA2, and RNA3. The inoculated protoplasts were incubated at 20°C for 18 h in the presence of n-butanol. (B) The effect of an exogenously supplied PA on the replication of BMV. N. benthamiana protoplasts were inoculated with in vitro transcribed BMV RNA1, RNA2, and RNA3. The inoculated protoplasts were incubated at 20°C for 18 h in the presence of progressively increasing concentrations of PA. Total RNA was analyzed by Northern blotting using ribonucleotide probes that recognize specifically 3`UTR of BMV RNAs. Ethidium bromide-stained rRNAs were used as loading controls and are shown below the Northern blots.

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106 Summary

Chapter I

The specific recognition of genomic RNAs by viral replicase proteins is a key regulatory step during the early replication process in positive-strand RNA viruses. In this study, I characterized the RNA-binding activity of the auxiliary replicase protein p27 of Red clover necrotic mosaic virus (RCNMV), which has a bipartite genome consisting of RNA1 and RNA2. Aptamer pull-down assays identified the amino acid residues of p27 involved in its specific interaction with RNA2. The RNA-binding activity of p27 correlated with its activity in recruiting RNA2 to membranes. I also identified the amino acids required for the formation of the 480-kDa replicase complex, a key player of RCNMV RNA replication. These amino acids are not involved in the functions of p27 that bind viral RNA or replicase proteins, suggesting an additional role for p27 in the assembly of the replicase complex. Our results demonstrate that p27 has multiple functions in RCNMV replication.

Chapter II

Eukaryotic positive-strand RNA viruses replicate using the membrane-bound replicase complexes, which contain multiple viral and host components. Virus infection induces the remodeling of intracellular membranes. Virus-induced membrane structures are thought to increase the local concentration of the components that are required for replication and provide a scaffold for tethering the replicase complexes. However, the mechanisms underlying virus-induced membrane remodeling are poorly understood. RNA replication of red clover necrotic mosaic virus (RCNMV), a positive-strand RNA plant virus, is associated with the endoplasmic reticulum (ER) membranes, and ER morphology is perturbed in RCNMV-infected cells. Here, I identified ADP ribosylation

107 factor 1 (Arf1) in the affinity-purified RCNMV RNA-dependent RNA polymerase fraction. Arf1 is a highly conserved, ubiquitous, small GTPase that is implicated in the formation of the coat protein complex I (COPI) vesicles on Golgi membranes. Using in vitro pulldown and bimolecular fluorescence complementation analyses, I showed that Arf1 interacted with the viral p27 replication protein within the virus-induced large punctate structures of the ER membrane. I found that inhibition of the nucleotide exchange activity of Arf1 using the inhibitor brefeldin A (BFA) disrupted the assembly of the viral replicase complex and p27-mediated ER remodeling. I also showed that BFA treatment and the expression of dominant negative Arf1 mutants compromised RCNMV RNA replication in protoplasts. Interestingly, the expression of a dominant negative mutant of Sar1, a key regulator of the biogenesis of COPII vesicles at ER exit sites, also compromised RCNMV RNA replication. These results suggest that the replication of RCNMV depends on the host membrane traffic machinery.

Chapter III

Eukaryotic positive-strand RNA viruses replicate using the membrane-bound replicase complexes, which contain multiple viral and host components. These cellular membranes are thought to facilitate the building of viral factories, promote a high concentration of membrane-bound viral proteins, and provide protection against cellular nucleases and proteases. The membrane lipids and proteins may serve as scaffolds for targeting the viral replication proteins or for the assembly of the viral replicase complex. However, understanding the roles of various lipids and lipid biosynthesis enzymes and pathways in (+)RNA virus replication is limited. In this study, by using two-step affinity purification and liquid chromatography-tandem mass spectrometry analysis, I identified two Nicotiana benthamiana phospholipase D (PLD), namely PLD& and PLD%, as interaction partners of RCNMV replication proteins. Gene-silencing and pharmacological inhibition approaches showed that PLDs-derived PA played a positive role in viral RNA replication. In consistent with this, direct application of PA to virus-infected plant cells or plant-derived cell-flee systems enhanced the viral RNA

108 replication. I found that the effects of exogenously applied PA on viral RNA replication were different between RCNMV genomic RNAs. I also showed that p27 is a PA-binding protein. RCNMV-infected plant leaves showed high accumulation of PA, leading to the possibility that RCNMV hijacks host PA signaling pathway for successful RNA replication.

109 Acknowledgments

I would like to express my deepest appreciation to my advisor, Professor Dr. Tetsuro Okuno for his guidance to the fields of molecular plant virology and plant pathology, helpful suggestion, considerable encouragement and invaluable discussion. Also, I would like to express my gratitude to Professor Dr. Masayuki Sakuma (Kyoto University) and Associate Professor Dr. Takashi Yoshida (Kyoto University) for reviewing this thesis. I would like to express my sincere appreciation to Associate Professor Dr. Kazuyuki Mise for instruction, helpful suggestion and valuable discussion throughout of the course of this study. I would like to extend my appreciation to Dr. Masanori Kaido and Associate Professor Dr. Yoshitaka Takano for their advice, support and holding a hot discussion on this work. I am deeply grateful to Professor Dr. Hisaaki Taniguchi (The University of Tokushima) and Dr. Takako Taniguchi (The University of Tokushima) for their technical help on mass spectrometry, and to Professor Dr. Tatsuya Sugawara (Kyoto University) and Mr. Yuuki Manabe for their technical help on TLC analysis. I greatly appreciate Dr. Hiro-oki Iwakawa (Tokyo University) and Dr. Akira Mine (Max-Planck-Institute) for their valuable cooperation and helpful suggestions. I heartily thank Ms. Keiko Hashimoto for her cooperation and cheerful supports. I express my sincere thanks to all the previous and present members of the laboratory of Plant Pathology, Kyoto University, for their cheerful support. Finally, I would like to thank my friends and family for their continued support of my academic endeavors.

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