Phosphatidic Acid Produced by Phospholipase D Promotes RNA Replication of a Plant RNA Virus

Eukaryotic positive-strand RNA [(+)RNA] viruses are intracellular obligate parasites replicate using the membrane-bound replicase complexes that contain multiple viral and host components. To replicate, (+)RNA viruses exploit host resources and modify host metabolism and membrane organization. Phospholipase D (PLD) is a phosphatidylcholine- and phosphatidylethanolamine-hydrolyzing enzyme that catalyzes the production of phosphatidic acid (PA), a lipid second messenger that modulates diverse intracellular signaling in various organisms. 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 environmental cues such as biotic and abiotic stresses in plants. However, the precise functions of PLD and PA remain unknown. Here, we report the roles of PLD and PA in genomic RNA replication of a plant (+)RNA virus, Red clover necrotic mosaic virus (RCNMV). We found that RCNMV RNA replication complexes formed in Nicotiana benthamiana contained PLDα and PLDβ. Gene-silencing and pharmacological inhibition approaches showed that PLDs and PLDs-derived PA are required for viral RNA replication. Consistent with this, exogenous application of PA enhanced viral RNA replication in plant cells and plant-derived cell-free extracts. We also found that a viral auxiliary replication protein bound to PA in vitro, and that the amount of PA increased in RCNMV-infected plant leaves. Together, our findings suggest that RCNMV hijacks host PA-producing enzymes to replicate.

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 dehydration, salt, and osmotic stress [20][21][22]. PA also accumulates in response to several microbe-associated molecular patterns (MAMPs) in plant cells and positively regulates salicylic acid (SA)-mediated defense signaling [23][24][25][26][27]. Moreover, effector proteins of bacterial and fungal pathogens, such as Cladosporium fulvum Avr4 and Pseudomonas syringae AvrRpm1 and AvrRpt2, trigger PA accumulation in their host cells, and multiple PLD isoforms contribute to AvrRpm1-triggered resistance in Arabidopsis thaliana [28][29][30]. PLDδ plays a positive role in MAMPs-triggered cell wall based immunity and nonhost resistance against Blumeria graminis f. sp. hordei [31]. Moreover, overexpression of rice diacylglycerol (DAG) kinase, which catalyzes the conversion of DAG to PA, enhances resistance against tobacco mosaic virus and Phytophthora parasitica infections in tobacco plants [32]. In accordance with this, direct application of PA to leaves has been shown to induce the expression of pathogenesis-related (PR) genes and cell death [28,33]. These findings indicate that PA is a positive regulator in plant defense against pathogens. In contrast, PLDβ1 acts like a negative regulator of the generation of reactive oxygen species (ROS), the expression of PR genes, and plant defenses against biotrophic pathogens in rice and Arabidopsis [34][35][36].
In this study, using two-step affinity purification and liquid chromatography-tandem mass spectrometry (LC/MS/MS) analysis, we identified Nicotiana benthamiana PLDα and PLDβ as interaction partners of RCNMV replication protein, p88 pol . Gene-silencing and pharmacological inhibition approaches show that PLDs-derived PA plays a positive role in viral RNA replication. Consistent with this role, direct application of PA to plant cells or plant-derived cellfree extracts enhanced RCNMV RNA replication and negative-strand RNA synthesis, respectively. We found that p27 auxiliary replication protein interacted with PA in vitro and that the accumulation of PA increased in RCNMV-infected plant leaves. Together, our findings suggest that RCNMV hijacks host PA-producing enzymes to achieve successful RNA replication.

Results
To identify putative host proteins associated with the RCNMV replicase complex, we expressed six-His/FLAG-tagged p27 and p88 pol replication proteins (p27-HF and p88 pol -HF, respectively) together with an RNA2 replication template via agroinfiltration in N. benthamiana. Twodays after infiltration (dai), RCNMV replication proteins were purified via sequential affinity purification using nickel-agarose beads and FLAG-affinity resins as described in the Materials and Methods section. Note that p27-HF and p88 pol -HF replication proteins can support RNA2 replication in N. benthamiana plants, although their activities were low compared with those of non-tagged p27 and p88 pol (S1 Fig).
The purified fraction was subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis to separate the proteins that copurified with RCNMV replication proteins. A silverstained gel showed the presence of many protein bands that were absent in the control fraction prepared from Agrobacterium-infiltrated N. benthamiana leaves expressing non-tagged p27 and p88 pol together with RNA2 (Fig 1). LC/MS/MS analysis of the isolated proteins excised from gels led to the identification of many host proteins (Fig 1 and S1 Table). These proteins included PLDα and PLDβ, and several Arf1 effector proteins, such as coatomer subunits and clathrin heavy chain, in addition to previously identified host factors, HSP70 and HSP90 [16], and Arf1 [15]. It is known that the activities of yeast and mammalian PLDs are stimulated by Arf1 [37] and Arf1 is an essential host factor in RCNMV RNA replication [15]. Therefore, we investigated whether PLDα and PLDβ are also required for RCNMV RNA replication.
We isolated cDNAs encoding PLDα and PLDβ from N. benthamiana leaves. Deduced amino acid sequences and peptide sequences identified from LC/MS/MS analysis are presented in S2 Fig. To confirm the association of N. benthamiana (Nb)PLDα and NbPLDβ with RCNMV replication proteins, we performed a co-immunoprecipitation (co-IP) assay in N. benthamiana plants using green fluorescent protein (GFP)-fused NbPLDα or NbPLDβ as bait proteins. We co-expressed p88 pol -HF together with p27 and RNA2, because p88 pol can be detected by immunoblot in N. benthamiana only when viral RNA replication takes place [12]. Both p27 and p88 pol -HF were co-immunoprecipitated with GFP-NbPLDα or GFP-NbPLDβ, but not with GFP (Fig 2A), confirming the association of these PLDs with RCNMV replication proteins.
To investigate whether PLDs localize at VRCs, GFP-NbPLDα or GFP-NbPLDβ was co-expressed with mCherry-fused p27 (as a marker of VRC), p88 pol , and RNA2 via agroinfiltration Identification of proteins copurified with RCNMV replication proteins. The solubilized total fractions prepared from Agrobacterium-infiltrated leaves expressing p27 plus p88 plus RNA2 or p27-His-FLAG (p27-HF) plus p88-His-FLAG (p88-HF) plus RNA2 were subjected to affinity purification with Ni-NTA agarose beads, followed by anti-FLAG affinity resins. The affinity-purified fractions were subjected to SDS-PAGE and stained using MS-compatible silver staining. Protein bands were excised, subjected to in-gel digestion, and analyzed by tandem mass spectrometry. Some of the proteins that were identified in LC/MS/MS analysis are listed in S1 Table. doi:10.1371/journal.ppat.1004909.g001 Roles of Phosphatidic Acid in Virus RNA Replication PLOS Pathogens | DOI:10.1371/journal.ppat.1004909 May 28, 2015 in N. benthamiana. At 4 dai, the fluorescence of GFP-NbPLDα and GFP-NbPLDβ was partially merged with the fluorescence of p27-mCherry in large aggregate structures (Fig 2B panels I  and II), which are induced by p27 and are thought to be the site of RCNMV RNA replication [13,15,16]. The expression of GFP-NbPLDα or GFP-NbPLDβ alone did not induce such aggregated structures (Fig 2B panels III and IV), suggesting that both NbPLDα and NbPLDβ are recruited to viral replication sites during RCNMV replication.
To investigate whether p27 or p88 pol interact with NbPLDα and NbPLDβ, we performed a co-IP assay in cell lysates prepared from evacuolated tobacco BY-2 protoplasts (BYL) [38]. Cterminally HA-tagged viral replication protein (p27-HA or p88 pol -HA) was coexpressed with C-terminally FLAG-tagged PLD (NbPLDα-FLAG or NbPLDβ-FLAG) from capped transcripts in BYL. In BYL, p88 pol -HA was easily detected by immunoblot using anti-HA antibody in the absence of other viral components after 2 hour of in vitro translation ( Fig 2C). After in vitro translation, the extracts were subjected to immunoprecipitation using FLAG-affinity resin. Immunoblot analysis showed that p88 pol -HA, but not p27-HA, was copurified with both NbPLDα-FLAG and NbPLDβ-FLAG ( Fig 2C). p88 pol -HA was not copurified with C-terminally FLAG-tagged firefly luciferase (FLuc-FLAG) that was used as a negative control, excluding the possibility that p88 pol -HA binds to FLAG-affinity resin nonspecifically. In reciprocal co-IP experiments using HA antibody, we also found copurification of NbPLDα-FLAG with p88 pol -HA, but not with FLuc-HA or p27-HA (S3 Fig). However, as NbPLDβ-FLAG was co-purified not only with p88 pol -HA but also with FLuc-HA or p27-HA (S3 Fig), we could not confirm the specific interaction of p88 pol with NbPLDβ in BYL.

Both PLDα and PLDβ are required for RCNMV infection
To test whether NbPLDα and NbPLDβ are required for infection of host plants with RCNMV, endogenous transcript levels of NbPLDα or NbPLDβ were downregulated using Tobacco rattle virus (TRV)-based virus-induced gene silencing (VIGS) in N. benthamiana plants. A TRV vector harboring a partial fragment of NbPLDα (TRV:NbPLDα) or NbPLDβ (TRV:NbPLDβ) was 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. No morphological defects such as chlorotic and stunted phenotypes were observed at this stage (S4 Fig). Semiquantitative reverse-transcription-PCR (RT-PCR) or quantitative RT-PCR (RT-qPCR) analyses confirmed the specific reduction of NbPLDα or NbPLDβ mRNAs in plants infiltrated with TRV:NbPLDα or TRV:NbPLDβ, respectively (Figs 3 and S5). Northern blot analyses using ribonucleotide probes specifically recognizing RCNMV RNA1 or RNA2 showed that the accumulation of RCNMV RNAs was dramatically reduced in both NbPLDα-and NbPLDβ-knockdown plants compared with control plants (Fig 3).
It is known that, in PLDβ1 knockdown transgenic rice plants, the generation of ROS and the expression of defense-related genes are induced even in the absence of pathogen infection [35]. Therefore, it is possible that the poor viral infection in NbPLDβ-knokedown plants was due to activated defense responses. To address this possibility, we tested the effects of NbPLDαor NbPLDβ-knockdown by TRV-mediated VIGS on the expression of defense-related genes by RT-qPCR analysis. The defense-related genes analyzed here were SA-signaling marker genes (PR-1, PR-2, and PR-5) [39,40], jasmonic acid (JA)-signaling marker genes (LOX1, PR-4, and PDF1.2.) [39,41], ROS-detoxification enzymes (APX, GST, and SOD) [39], MAMP-triggered immunity marker genes (CYP71D20 and ACRE132) [42], and mitogen-activated protein kinases (WIPK and SIPK) [42]. The expression levels of these defense-related genes were not significantly increased in NbPLDα-or NbPLDβ-knockdown plants compared with those in TRV control plants (S5 Fig), excluding the possibility that the reduced viral infection in NbPLDα-or NbPLDβ-knockdown plants are due to activated defense responses in these plants. Some genes including PR-1, PR-2, and CYP71D20 genes were even repressed in NbPLDα-or NbPLDβknockdown plants (S5 Fig), consistent with the positive roles of PLDs and PLD-derived PA on plant defense signaling [24][25][26][27][28][29][30]. Altogether, these results suggest that both NbPLDα and NbPLDβ play a positive role in RCNMV infection.

PA promotes viral RNA replication
To test the possible contribution of PLD-derived PA to viral RNA replication, we exploited the transphosphatidylation activity of PLDs, which uses primary alcohols as substrates to form an artificial phosphatidyl alcohol. The preferential formation of this compound impairs PA production [43]. Thus, we tested the effect of n-butanol that inhibits PA production by PLDs on RCNMV RNA replication. N. benthamiana protoplasts were inoculated with RCNMV RNA1 and RNA2 and incubated with n-butanol or tert-butanol, an alcohol with no inhibitory effect on PA production, and viral RNA accumulation was determined by northern blot analysis. Increasing n-butanol concentration caused a progressive reduction of viral RNA accumulation ( Fig 4A). By contrast, viral RNA accumulation was only moderately reduced in protoplasts treated with tert-butanol compared with the water control. Note that n-butanol did not affect the accumulation of rRNA (Fig 4A). The inhibitory effect of n-butanol on RCNMV replication was also observed in tobacco BY-2 protoplasts (S6 Fig). We also tested the effect of n-butanol and tert-butanol on defense-related gene expressions in N. benthamiana protoplasts. n-butanol did not increase the expression of defense-related genes (S7 Fig). This result corresponds well with the finding that n-butanol has no effects on the basal transcription of the PR-1 gene in Arabidopsis seedlings [27]. In contrast, tert-butanol treatment caused the induction of CYP71D20, ACRE132, and WIPK genes (S7 Fig). This may explain weak negative effect of tert-butanol on RCNMV replication (Figs 4A and S6). Altogether, these results suggest that PLD-derived PA plays a positive role in viral RNA replication.
To verify further the importance of PA in viral RNA replication, commercially available soy-derived PA was supplied to RCNMV-inoculated N. benthamiana protoplasts and viral Knockdown of NbPLDα and NbPLDβ mRNAs levels via gene silencing inhibits the accumulation of RCNMV RNAs in N. benthamiana. The tobacco rattle virus (TRV) vector harboring a partial fragment of N. benthamiana NbPLDα and NbPLDβ mRNAs (TRV:NbPLDα and TRV:NbPLDβ, respectively) 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 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 RNA was analyzed by northern blotting. Ethidium bromide-stained ribosomal RNAs (rRNAs) are shown below the northern blots, as loading controls. The numbers below the images represent the relative accumulation levels (means ± standard error) of viral RNAs (RNA1 and RNA2, respectively) using the Image Gauge program (Fuji Film), which were calculated based on three independent experiments. NbPLDα and NbPLDβ 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. sgRNA, subgenomic RNA; SR1f, a small RNA fragment that derived from non-coding region of RNA1 [47].
RNA accumulation was determined by northern blot analysis. Exogenously added PA enhanced the accumulation of RNA1 in a dose-dependent manner (up to 6-fold increase by 5 μM 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 hours 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. The numbers below the images represent the relative accumulation levels (means ± standard error) of viral RNAs (RNA1 and RNA2, respectively) using the Image Gauge program, which were calculated based on three independent experiments. In (C) and (D), total protein was analyzed by Immunoblot using anti-HA antibody. Coomassie Brilliant Blue (CBB) staining served as a loading control. sgRNA, subgenomic RNA; SR1f, a small RNA fragment that derived from non-coding region of RNA1 [47].
doi:10.1371/journal.ppat.1004909.g004 PA), whereas the effect of PA on the accumulation of RNA2 was negligible ( Fig 4B). Neither exogenously supplied PC nor PE significantly affected the accumulation of viral RNA in N. benthamiana protoplasts (S8 Fig). These results suggest that PA plays an important role in RCNMV RNA replication, and that the requirement for PA may differ between RNA1 and RNA2.
Replication of RNA2 depends entirely on RNA1 simply because RNA2 uses replication proteins supplied from RNA1. The negative impact of n-butanol on the accumulation of RNA2 ( Fig 4A) may be the indirect action of this primary alcohol through inhibition of RNA1 replication. Therefore, it is possible that RNA2 replication does not require any PLD-derived PA. To investigate this possibility, we inoculated N. benthamiana protoplasts with RNA2 and plasmids expressing p27 and p88 pol , as suppliers of the replication proteins. The accumulation of RNA2 was decreased by n-butanol in a dose-dependent manner ( Fig 4C). Note that in this experiment, the accumulation of p27 replication protein was not significantly changed ( Fig 4C).
These results indicate that PLD-derived PA was also required for the replication of RNA2 as in the case of RNA1. However, the replication of RNA2 was not enhanced by exogenously supplied PA (Fig 4D), suggesting that the threshold of PA requirement for RNA2 replication is lower than that for RNA1. The differential requirement for PA in the replication of RNA1 and RNA2 is discussed later.
Next, we investigated the effect of n-butanol on RNA replication of Brome mosaic virus (BMV), another plant (+)RNA virus, which is unrelated to RCNMV. Increasing n-butanol concentration caused progressive reduction in the accumulation of BMV RNA ( Fig 5A). Co-IP experiments showed interactions between BMV replication proteins and NbPLDβ-FLAG (Fig 5B  and 5C). These results suggest that PLD-derived PA is also important for BMV RNA replication. However, exogenously added PA did not affect the accumulation of BMV RNA ( Fig 5D) as similarly seen for RCNMV RNA2.

PA stimulates VRC activity in vitro
PA acts as a second messenger in signal transduction during multiple biotic and abiotic stress responses and plays multiple roles including that for transcriptional reprogramming [20][21][22]. To investigate whether PA has a direct role in viral RNA replication, we took advantage of BYL, a nucleus-depleted (therefore, the effects of PA on transcriptional reprogramming are negligible) in vitro translation/replication system that has been used successfully to recapitulate the negative-strand RNA synthesis of RCNMV [12,[44][45][46][47][48][49]. Addition of PA into BYL stimulated the accumulation of newly synthesized negative-strand RNA (Fig 6A), and moderately enhanced the accumulation of the 480-kDa replicase complex (Fig 6B), suggesting that PA stimulates the activity and/or assembly of the viral replicase complex in a direct manner. The stimulation effects of PA on the accumulation of the 480-kDa replicase complex was less obvious than that on the accumulation of newly-synthesized viral RNA at the highest concentration of PA used in this experiment (50 μM). This result may also support a direct role for PA in the enhancement of RdRP activity rather than the formation of the replicase complex.

p27 replication protein binds to PA in vitro
Because PA directly stimulated the viral negative-strand RNA synthesis in BYL, we hypothesized that p27, the multifunctional RCNMV replication protein, has an affinity for PA. To investigate this possibility, we conducted a lipid overlay assay using bacterially expressed, purified C-terminally FLAG-tagged p27 (p27-FLAG) [49]. Purified p27-FLAG protein was incubated with phospholipid-spotted nitrocellulose membranes, and the interaction between p27-FLAG and phospholipids was detected using anti-FLAG antibody. p27 gave strong PA binding signals on the blot (Fig 7A and 7B). p27 also exhibited weak binding to phosphatidylinositol-4-phosphate (PI4P), phosphatidylinositol (3,5)-bisphosphate, phosphatidylinositol (4,5)-bisphosphate, and phosphatidylinositol (3,4,5)-trisphosphate, but negligible binding to other lipids, including PC and PE (Fig 7A and 7B). These results were consistent with the findings that neither exogenously supplied PC nor PE promoted RCNMV replication in N. benthamiana protoplasts (S8 Fig). N-or C-terminal halves of p27 fragments did not give PA binding signals (Fig 7C and 7D), suggesting that overall protein conformation may be important for p27-PA binding. Appropriate combinations of capped transcripts were added to BYL. After in vitro translation at 25°C for 2 h, the extract was solubilized and mixed with a 10 μl bed volume of anti-FLAG M2 antibody agarose and incubated further at 4°C for 1 hours. After washing, proteins copurified with FLAG-tagged proteins were analyzed by immunoblotting with appropriate antibodies. (D) The effect of an exogenously supplied PA on the replication of BMV. N. benthamiana protoplasts were inoculated with BMV RNA1, RNA2, and RNA3 transcribed in vitro. The inoculated protoplasts were incubated at 20°C for 18 hours in the presence of progressively increasing concentrations of PA. Total RNA was analyzed by northern blotting using ribonucleotide probes that specifically recognize the 3'-UTR of BMV RNAs. Ethidium bromide-stained rRNA was used as a loading control and is shown below the northern blots. The numbers below the images represent the relative accumulation levels (means ± standard error) of viral RNAs (RNA1 + RNA2 + RNA3 + RNA4) using the Image Gauge program, which were calculated based on three independent experiments.

RCNMV promotes PA accumulation in planta
Next we investigated whether RCNMV infection affects the amount of PA in plant leaves. N. benthamiana leaves were inoculated with RCNMV via agroinfiltration. At 2 dai, lipids were extracted and the amount of PA was analyzed by thin layer chromatography (TLC). Compared with control plant leaves infiltrated with Agrobacterium harboring an empty vector, the signal intensity of a lipid spot that showed a migration similar to that of soy-PA was increased in RCNMV-infected plant leaves (about 3-fold higher) (Fig 8A and 8B). To identify the lipid species of the spot, the same samples were again subjected to TLC, and the spot was scratched out from Coomassie Brilliant Blue R-250 stained TLC plates and subjected to LC/MS analysis. The lipid was identified as PA (Fig 8C-8E). We concluded that RCNMV infection upregulated PA accumulation in plants. It is known that expressions of PLD genes were induced by pathogen infection [50]. Therefore, the enhanced accumulation of PA in RCNMV-infected plants could

Discussion
A growing number of studies have suggested that PLD and PLD-derived PA play vital roles in environmental responses in plants [19][20][21][22]50]. The properties of PA (i.e., normally present in small amounts, and rapidly and transiently accumulates in response to various environmental cues) seem to be suitable for its function in biotic and abiotic responses in which plants need to rapidly accommodate their surrounding environments. Indeed, PA has been shown to accumulate in response to several MAMPs and pathogen effector proteins, or SA that is a key hormone involved in plant resistance against biotrophic pathogens [23][24][25][26][27][28][29]. PA induces PR gene expression and cell death [28,33,34], and has been proposed to act as an important component in resistance to biotrophic pathogens such as tobacco mosaic virus and Phytophthora parasitica. Plants have diverse numbers of PLD isoforms and they appear to have distinct but somewhat overlapping functions in cellular responses [50]. In Arabidopsis, it is proposed that multiple PLD isoforms cooperatively contribute to AvrRpm1-triggered resistance [30]. In rice, PLDβ1 acts like a negative regulator of defense signaling because PLDβ1-knockdown rice plants exhibit constitutive ROS production, expression of PR genes, and enhanced resistance against pathogens [35]. In this study, we showed that PLD and PA are essential for and play a key role in RCNMV replication. Poor infection of RCNMV to NbPLDα-or NbPLDβ-knockdown plants was not due to constitutively activated defense responses, indicating that both NbPLDα and NbPLDβ act as essential host factors in RCNMV replication. The findings reveal novel aspects of PLD and PA in their roles during biotic stress responses in plants.
The replication of Tomato bushy stunt virus is enhanced by deletion of the PAH1 gene encoding a PA phosphatase, which converts PA into DAG in yeast [51]. Moreover, ectopic expression of Arabidopsis PA phosphatase, Pah2 in N. benthamiana results in the inhibition of tombusviruses and RCNMV infection [51]. These findings suggest that PA is positively involved in the life cycles of these viruses. However, whether viruses manipulate PA production for viral replication has been unknown. In the current study, we showed that RCNMV replication proteins interacted with PA-producing enzymes, NbPLDα and NbPLDβ. RCNMV infection induced a high accumulation of PA in plant tissues, suggesting that RCNMV alters cellular lipid metabolism to establish a suitable environment for viral replication. It is known that transcription of PLD genes is induced by pathogen infection [50]. RCNMV infection increased the accumulation of NbPLDα and NbPLDβ transcripts (S9 Fig). Currently, whether the enhancement of PA accumulation observed in RCNMV-infected plants is due to the upregulation of PLD gene expression remains unknown. PLDs may be activated through direct or indirect interaction with RCNMV replication proteins. Although we failed to show the interaction between p88 pol and NbPLDα or NbPLDβ in vivo because p88 pol accumulates below detection limits in the absence of viral RNA replication in N. benthamiana [12], p88 pol interacted with PLDs, at least with NbPLDα in a co-IP assay in BYL. The interaction of p88 pol with PLDs seems to make sense for viral replication strategy because it brings them to the sites of replication. This strategy could increase PA only at VRCs and not at other cellular membranes where PA might affect cellular metabolism or activate the SA-mediated defense responses that is detrimental for successful viral infection. This is critical for the viral life cycle because PA interacts with p27 auxiliary replication protein (Fig 7) and enhances viral replication through upregulating the activity and/or assembly of the 480-kDa replicase complex (Fig 6). To our knowledge, this is the first demonstration of a functional role for PA in (+)RNA virus replication. It is likely that PA is involved in RNA replication of many (+)RNA viruses. Indeed, the replication proteins, 1a and 2a pol of BMV, a virus unrelated to RCNMV, also interacted with NbPLDβ, and BMV RNA replication was sensitive to n-butanol, which is an inhibitor of PLDs-derived PA production ( Fig 5). However, exogenously added PA did not affect the accumulation of BMV RNA (Fig 5), implying that the threshold of PA requirement for BMV RNA replication seems to be lower than that for RCNMV. The differential PA requirement may be explained by very weak affinity of BMV replication proteins with NbPLDα, which is more active in PA production than NbPLDβ (S10 Fig). Moreover, Dengue virus induces the accumulation of several lipids in infected mosquito cells, including PA [52]. It has been reported that Coxsackievirus B3 and mouse hepatitis coronavirus replication is insensitive to n-butanol [53,54]. However, because PA can also be formed through the combined action of phospholipase C and DAG kinase [55], it is unknown whether replication of these viruses depends on PA or not.
How does PA affect viral RNA replication? 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 [55]. We found that exogenous PA enhanced the accumulation of newly synthesized viral RNA and the formation of 480-kDa replicase complexes in BYL in vitro translation/replication systems, and that p27 had affinity for PA in vitro (Figs 6 and 7). Therefore, PA could promote viral replicase activity and/or assembly directly. The 480-kDa replicase complexes contain p27 oligomer. Therefore, it is possible that PA-binding to p27 in the replicase complexes assists the conformational change of p27 that is suitable for RNA synthesis. Alternatively, PA could serve as an assembly platform for host PA-binding proteins. PA binds to various proteins, including transcription factors, kinases, phosphatases, enzymes involved in central metabolism, and proteins involved in vesicular trafficking and cytoskeletal rearrangements [20,21]. Several known cellular PA-binding proteins were also identified in our LC/MS/MS analysis (S1 Table). These include NADPH oxidase [56], glyceraldehyde-3-phosphate dehydrogenase (GAPDH) [57,58], and SNF1-related kinase [58]. RCNMV-induced accumulation of high PA levels may facilitate the recruitment of these PAbinding proteins to viral replication sites. One of the candidate proteins is GAPDH-A. Although GAPDH-A is not required for RCNMV replication, it recruits RCNMV movement protein to viral replication sites and plays an important role in virus cell-to-cell movement [59]. Therefore, PA may also function in bridging viral replication and cell-to-cell movement. RCNMV induces large ER aggregates in infected cells, which are thought to be viral RNA replication factories [13][14][15][16]. PA may play a direct role in the formation of RNA replication factories. In support of this hypothesis, deletion of pah1 in yeast causes the expanded ER membranes and leads to enhanced TBSV replication on these membranes, although TBSV replicates normally at peroxisomes [51]. In addition, it is also possible that PA might affect unknown cellular factors that are involved in viral RNA replication. Further studies are needed to elucidate the molecular functions of PA in viral RNA replication.
Our results suggest that RNA1 and RNA2 have differential PA requirements in RNA replication. How does this difference contribute to RCNMV infection? RNA1 requires a larger amount of PA for maximum efficiency of its replication than that required by RNA2. This may reflect differences in the translation and replication mechanisms between RNA1 and RNA2. Translation of MP from RNA2 couples with RNA replication: only progeny RNA2 generated de novo through the RNA replication pathway could function as mRNA [60]. Poor enhancement of RNA2 replication by PA may be beneficial for switching from replication to translation. By contrast, because RNA1 has a cap-independent translation enhancer that is an effective recruiter of translation factors [61], newly synthesized RNA1 serves as a template for further translation of the replication proteins that would activate PA production in infected cells. Moreover, RNA1 also serves as a template for transcription of subgenomic RNA, which encodes CP. Therefore, PA-mediated enhancement of RNA1 replication may be suitable for the production of CP subgenomic RNA during the late stage of viral infection. High PA requirement for maximum RNA1 replication may explain the use of both NbPLDα and NbPLDβ as essential host factors.
A growing number of studies have suggested that multiple lipid species affect virus replication and that (+)RNA viruses employ a multifaceted strategy to rewire host machinery involved in lipid transport and synthesis [62]. HCV infection stimulates the production of cellular PC, PE, and PI4P and HCV co-opts PI4KIIIα for replication [63,64]. In addition, HCV infection stimulates the accumulation of cellular sphingomyelin, which binds to and activates HCV NS5B polymerase [65,66]. Enterovirus utilizes PI4KIIIβ for RNA replication and viral RdRP 3D pol binds to PI4P [67]. Enteroviruses also upregulate cellular uptake of fatty acids, which are channeled toward highly upregulated PC synthesis in infected cells [68]. The elevated PC has been proposed to serve as a building block for the formations of the viral replication factory [68,69]. Recently, it has been shown that PE and PC stimulate TBSV RdRP activity in vitro [70]. RCNMV p27 showed affinity, not only for PA but also for other phospholipids such as PI4P in vitro. However, our LC/MS/MS analysis failed to detect any PI4P-producing enzymes, such as PI4K. Combined lipidomics, proteomics, and transcriptome analysis will be helpful for a comprehensive understanding of lipid species involved in viral RNA replication.
pBYLAtPAH1-HA. A PCR fragment from cDNA derived from A. thaliana was amplified using primers 5'-CTGGCGCGCCATGAGTTTGGTTGGAAGAGTTGGGAGTTTGATTT-3' and 5'-GCTGGCGCGCCTCAAGCGTAATCTGGAACATCGTATGGGTATTCAACCTC TTCTATTGGCAGTTTCC-3'. The amplified fragment was digested with AscI and inserted into the corresponding region of pBYL2.

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). Capped mRNAs were transcribed from NotI-linearized pBYL plasmids using T7 or SP6 RNA polymerase (TaKaRa Bio, Inc) 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.

Tandem affinity purification
Four-week-old N. benthamiana plants were agroinfiltrated as described previously [15]. At 2 days postinfiltration (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 a cocktail of protease inhibitors (Roche). Following the removal of cell debris by filtering the mixture through cheesecloth, the extract was centrifuged at 21,000g at 4°C for 10 min and the supernatant was mixed with Ni-NTA beads (400 μl) (Qiagen, Hilden, Germany) and incubated for 1 h at 4°C with gentle mixing. The beads were washed three times with 1 ml of buffer A containing 100 mM imidazole. The bound proteins were eluted with 1 ml of buffer A containing 500 mM imidazole. The eluted proteins were mixed with 50 μl of ANTI-FLAG M2-Agarose Affinity Gel (Sigma-Aldrich) and incubated for overnight at 4°C with gentle mixing. The beads were washed 3 times with 1 ml of buffer A. The bound proteins were eluted with 300 μl of buffer A containing 150 μg/ml 3 × FLAG peptides (Sigma-Aldrich). The eluted proteins were concentrated by acetone precipitation and dissolved in 1 × NuPAGE sample buffer (Invitrogen). The purified proteins were separated by sodium dodecyl sulfate (SDS)-PAGE (NuPAGE 3%-12% bis-Tris gel: Invitrogen) and visualized by silver staining (Wako, Osaka, Japan). Proteins in excised gel pieces were subjected to digestion with trypsin, LC-MS/MS analysis, and MASCOT searching as described previously [12].

Silencing of PLDα and PLDß in N. benthamiana
Appropriate combinations of silencing vectors were expressed via Agrobacterium infiltration in 3-to 4-week-old N. benthamiana plants as described previously [16]. At 18 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 the same inoculum were pooled, and total RNA was extracted using RNA extraction reagent (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 [16]. The mRNA levels of NbPLDα and NbPLDß were examined by RT-PCR using primer pairs 5 0 -TATCAAGGTAG AGGAGATAGGTGC-3 0 and 5 0 -TACATCATCTCCATCGTTCTCCTC-3 0 , and 5 0 -GAAGG CTTCAAAGCGCCATG-3 0 and 5 0 -CTTAGGCAAGGGACATCAGC-3 0 , 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 [16].

BYL experiments
The preparation of BYL was as described previously [38,46]. The BYL translation/replication assay was performed essentially as described previously [46]. Briefly, 300 ng of RNA1 was added to 30 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 [45,46,48].

Coimmunopurification assay
Four-week-old N. benthamiana plants were agroinfiltrated as described previously [15]. At 4 days postinfiltration (dpi), total proteins were extracted from 0.33 g of leaves in 1 ml of buffer A containing a cocktail of protease inhibitors (Roche). Following the removal of cell debris by centrifugation at 21,000g at 4°C for 10 min, the supernatant was mixed with GFP-Trap agarose beads (10 μl) (ChromoTek) and incubated for 1 h at 4°C with gentle mixing. The beads were washed 3 times with 1 ml of buffer A. The bound proteins were eluted by addition of 1 × SDS gel loading buffer, followed by incubation for 3 min at 95°C. Protein samples were subjected to SDS-PAGE, followed by immunoblotting with appropriate antibodies.
FLAG-or HA-tagged proteins were expressed in BYL by adding an in vitro transcript. After incubation at 25°C for 120 min, a 10-μl bed volume of anti-HA Affinity Matrix (Roche) or ANTI-FLAG M2-Agarose Affinity Gel (Sigma-Aldrich) was added to the BYL and further incubated for 60 min with gentle mixing. The resin was washed three times with 200 μl of TR buffer [38] supplemented with 150 mM NaCl and 0.5% Triton X-100. The bound proteins were eluted by addition of 1 × SDS gel loading buffer, followed by incubation for 3 min at 95°C. Protein samples were subjected to SDS-PAGE, followed by immunoblotting with appropriate antibodies.

Subcellular localization assays
Appropriate combinations of fluorescent protein-fused proteins were expressed in N. benthamiana leaves by Agrobacterium infiltration. Fluorescence of GFP and mCherry was visualized with confocal microscopy at 4 dai as described previously [59].

Protein purification and lipid overlay assay
Protein expression in E. coli BL21 (DE3) and subsequent purification were done as described previously [15]. The concentration of purified protein was measured using a Coomassie Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA). The purified protein was subjected to SDS-PAGE and visualized with Coomassie brilliant blue R-250 to check its purity.
Lipid overlay assays were conducted as recommended by the manufacture's protocol. Briefly, the membrane (PIP Strips or Membrane Lipid Arrays; Echelon Bioscience Inc) was incubated in 3% fatty acid free BSA (Sigma-Aldrich) in a mixture of phosphate-buffered saline and 0.1% Tween 20 (PBST) for 1 h at room temperature (RT) and then incubated in the 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 (1:10000; KPL) was used as a secondary antibody. Binding of proteins to phospholipids was visualized by incubation with a chemiluminescent substrate.

Lipid extraction and TLC
Four-week-old N. benthamiana plants were inoculated with RCNMV via agroinfiltration. At 2 dai, 0.33 g of infiltrated leaves were ground in liquid nitrogen and extracted in 900 μl of water. Total lipids were extracted by adding 3 ml CHCl 3 /CH 3 OH (2:1, v/v) to each sample. The samples were vortexed and centrifuged at 1690g, at 4°C for 10 min. The organic phase was recovered and dried under nitrogen gas stream. Lipids were dissolved in 100 μl CHCl 3 /CH 3 OH (2:1, v/v). Subsequently, 5 μl of the samples were analyzed on TLC plates (Merck, Germany). The chromatography was performed using CHCl 3 /CH 3 OH/formic acid/acetic acid (12:6:0.6:0.4, v/ v). Plates were air-dried, soaked in 10% CuSO4, and charred at 180°C for 10 min to visualize lipids.
To identify lipid species, the air-dried TLC plates were stained for an hour in a 0.03% Coomassie Brilliant Blue R-250 solution containing 20% of methanol and 0.5% of acetic acid. Destaining of the plates was performed with 20% methanol containing 0.5% acetic acid for 5 min. After drying the plates for a few minutes, the blue bands of interest were scratched out and transferred to glass tubes. The scratched silica gels were mixed with chloroform/methanol (3/7, v/v), followed by the centrifugation at 1,690g, at 4°C for 10 min. The supernatants were subjected to the LC-MS analysis using LCMS-IT-TOF mass spectrometer (Shimadzu, Kyoto, Japan). A TSK gel ODS-100Z column (2.0 × 150 mm, 5 μm, Tosoh, Tokyo, Japan) was eluted isocratically with acetonitrile/methanol/2-propanol/water (6/131/110/3, by volume) containing 19.6 mM of ammonium formate and 0.2% of formic acid at a flow rate of 0.2 mL/min. The MS was performed using an electrospray ionization interface operated in negative ion mode, under the following conditions: CDL temperature, 200°C; block heater temperature, 200°C; nebulizing gas (N 2 ) flow, 1.5 L/min. The MS data were acquired in the range of m/z 600 to 1,000 using 10 msec ion accumulation time. The MS 2 data were acquired in the range of m/z 125 to 500, using 50 msec ion accumulation time and automatic precursor ion selection in the range of m/ z 650 to 750. CID parameters were follows: energy, 50%; collision gas (argon) 50%.

Quantitative RT-PCR analysis
Total RNA extracted from N. benthamina leaves or protoplasts were subjected to reverse transcription using PrimeScript RT reagent Kit (Takara) using oligo-dT and random primers according to manufacturer's protocol. Real-time PCR was carried out using SYBR Premix Ex Taq (RR420A, Takara). Primers used for real-time PCR analysis were listed in S2 Table. Quantitative analysis of each mRNA was performed using a Thermal cycler Dice Real Time System TP800 (Takara).

Accession numbers
NbPLDα and NbPLDβ were registered through DDBJ and accession numbers LC033851 and LC033852, respectively, were given on March 11 2015.
Supporting Information S1 Table. Table. List of primers used in RT-qPCR analysis. (EPS) S1 Fig. Six-His-and FLAG-tagged p27 (p27-HF) and p88 pol (p88 pol -HF) support RNA2 replication in N. benthamiana. Total RNA was extracted from Agrobacterium-infiltrated leaves expressing p27 plus p88 plus RNA2 or p27-HF plus p88-HF plus RNA2 at 2 days after infiltration, and the accumulations of positive-or negative-stranded RNA2 were analyzed by northern blotting. Ethidium bromide-stained rRNA was used as a loading control and is shown below the northern blots. Protoplasts were inoculated with in vitro transcribed RNA1 and RNA2. The inoculated protoplasts were incubated at 20°C for 18 hours in the presence of n-butanol. Accumulation of RCNMV RNAs was analyzed by northern blotting. Ethidium bromide-stained ribosomal RNAs (rRNAs) are shown below the northern blots, as loading controls. sgRNA, subgenomic RNA; SR1f, a small RNA fragment that derived from non-coding region of RNA1 [47]. has no effects on 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 hours in the presence of phospholipids (0, 1, 2.5, or 5 μM). Accumulation of RCNMV RNA was analyzed by northern blotting. Ethidium bromide-stained RNAs (rRNA) is shown below the northern blots, as loading controls. The numbers below the images represent the relative accumulation levels (means ± standard error) of viral RNAs (RNA1 and RNA2, respectively) using the Image Gauge program, which were calculated based on three independent experiments. sgRNA, subgenomic RNA; SR1f, a small RNA fragment that derived from noncoding region of RNA1 [47].