Protein Poly(ADP-ribosyl)ation Regulates Immune Gene Expression and Defense Responses

Summary

Fine-tuning of gene expression is a key feature of successful immune responses. However, the underlying mechanisms are not fully understood. Through a genetic screen in model plant Arabidopsis, we reveal that protein poly(ADP-ribosyl)ation (PARylation) post-translational modification plays a pivotal role in controlling plant immune gene expression and defense to pathogen attacks. PARylation is primarily mediated by poly(ADP-ribose) polymerase (PARP), which transfers ADP-ribose moieties from NAD+ to acceptor proteins. The covalently attached poly(ADP-ribose) polymers on the accept proteins could be hydrolyzed by poly(ADP-ribose) glycohydrolase (PARG). We further show that members of Arabidopsis PARPs and PARGs possess differential in vivo and in vitro enzymatic activities. Importantly, the Arabidopsis parp mutant displayed reduced, whereas parg mutant displayed enhanced, immune gene activation and immunity to pathogen infection. Moreover, Arabidopsis PARP2 activity is elevated upon pathogen signal perception. Compared to the lethality of their mammalian counterparts, the viability and normal growth of Arabidopsis parp and parg null mutants provide a unique genetic system to understand protein PARylation in diverse biological processes at the whole organism level.

Introduction

Plants sense the presence of pathogens by the cell surface-localized pattern recognition receptors (PRRs), which perceive evolutionarily conserved pathogen- or microbe-associated molecular patterns (PAMPs or MAMPs), including bacterial flagellin, lipopolysaccharide (LPS), peptidoglycan (PGN), elongation factor Tu (EF-Tu), and fungal chitin [1]–[3]. A 22-amino-acid peptide corresponding to a region near the amino-terminus of flagellin (flg22) is recognized by the Arabidopsis PRR Flagellin-Sensing 2 (FLS2), a leucine-rich repeat receptor-like kinase (LRR-RLK) [4], [5]. Perception of flg22 by FLS2 induces instantaneous association with another LRR-RLK, Brassinosteroid Insensitive 1 (BRI1)-Associated Kinase 1 (BAK1), mainly through ectodomain heterodimerization of flg22-activated FLS2/BAK1 complex [6]–[9]. The receptor-like cytoplasmic kinases (RLCKs), BIK1 and its homolog PBL1, constitutively associate with FLS2 and BAK1, and are released from the receptor complex upon flg22 perception [10]–[12]. BAK1 directly interacts and phosphorylates BIK1 at both serine, threonine and tyrosine residues, thereby activating downstream signaling [12], [13]. In addition, both BAK1 and BIK1 complex with PRR EFR (receptor for EF-Tu) [11], [14], AtPEPR1 (receptor for endogenous danger signal Pep1) [12], [15], [16], and plant brassinosteroid hormone receptor BRI1 [17]–[19]. Activation of PRR complex by the corresponding MAMP triggers a series of defense responses, including rapid activation of MAP kinases (MAPKs) and calcium-dependent protein kinases, transient reactive oxygen species (ROS) production and calcium influx, stomatal closure, callose deposition and massive transcriptional reprogramming [1]–[3]. It has been shown recently that BIK1 is able to phosphorylate plasma membrane-resident NADPH oxidase family member respiratory burst oxidase homolog D (RBOHD), thereby contributing to ROS production [20], [21]. However, it remains largely unknown how PRR complex activation leads to profound immune gene transcriptional reprograming.

Protein poly(ADP-ribosyl)ation (PARylation), an important post-translational modification process, plays a crucial role in a broad array of cellular responses including DNA damage detection and repair, cell division and death, chromatin modification and gene transcriptional regulation [22]–[24] (S1 Fig.). PARylation is primarily mediated by members of poly(ADP-ribose) polymerases (PARPs), which transfer ADP-ribose moieties from nicotinamide adenine dinucleotide (NAD⁺) to different acceptor proteins at glutamate (Glu), aspartate (Asp) or lysine (Lys) residues resulting in the formation of linear or branched poly(ADP-ribose) (PAR) polymers on acceptor proteins (S1 Fig.). PAR activities and PARPs have been found in a wide variety of organisms from archaebacteria to mammals and plants, but they are apparently absent in yeast [25]. Human PARP-1 (HsPARP-1) is the most abundant and ubiquitous PARP among a family of 17 members, and it catalyzes the covalent attachment of PAR polymers on itself (auto-PARylation) and other target proteins, including histones, DNA repair proteins, transcription factors, and chromatin modulators [22]. HsPARP-1 possesses three functional domains with a DNA binding domain at N-terminus, auto-modification domain in the middle and a catalytic domain at C-terminus (S2A Fig.). PARylation is a reversible reaction and the covalently attached PAR on the target proteins can be hydrolyzed to free PAR or mono-(ADP-ribose) by poly (ADP-ribose) glycohydrolase (PARG) [22], [23] (S1 Fig.). PARG contains both endo- and exo-glycohydrolase activities that promote rapid catabolic destruction of PAR of target proteins [26]. There is only one PARG gene in humans with three different isoforms: PARG₉₉ and PARG₁₀₂ in the cytoplasm and PARG₁₁₀ in the nucleus [26]. Mammalian PARG possesses a regulatory and targeting domain (A-domain) at the N-terminus, a mitochondrial targeting sequence (MTS) in the middle and a conserved catalytic domain at the C-terminus [27] (S2B Fig.). The catalytic core containing “GGG-X_6-8-QEE” PARG signature motif interacts with PAR and executes hydrolysis activity [28]. Despite of their apparently opposing activities, members of PARPs and PARGs coordinately regulate protein PARylation and play essential roles in a wide range of cellular processes and contribute to the pathogenicity of various diseases, including cancer, cardiovascular diseases, stroke, metabolic disorders, diabetes and autoimmunity [25].

The Arabidopsis genome encodes three members of PARPs, AtPARP1 (At2g31320), AtPARP2 (At4g02390) and AtPARP3 (At5g22470) and two members of PARGs, AtPARG1 (At2g31870) and AtPARG2 (At2g31865) [23], [29] (S2 Fig.). AtPARP1 (it was originally named as AtPARP2) shares the conserved domain structure with HsPARP-1, whereas AtPARP2 (it was originally named as AtPARP1) and AtPARP3 more closely resemble HsPARP-2 and HsPARP-3 [29] (S2A Fig.). As their mammalian counterparts, plant PARPs are implicated in DNA repair, cell cycle and genotoxic stress [29]-[32]. Importantly, plant PARPs play an essential role in response to abiotic stresses. Transgenic Arabidopsis or oilseed rape (Brassica napus) plants with reduced PARP gene expression were more resistant to various abiotic stresses, including drought, high light and heat, partially attributed to a maintained energy homeostasis of reduced NAD⁺ and ATP consumption and alternation in plant hormone abscisic acid (ABA) levels in the transgenic plants [33], [34]. The two Arabidopsis PARG genes, AtPARG1 and AtPARG2, which were likely derived from a tandem duplication event, locates next to each other on the same chromosome [23]. AtPARG1 (TEJ) was originally identified as a regulator of circadian rhythm and flowering in Arabidopsis [35]. Interestingly, the AtPARG2 gene was robustly induced by the treatments of MAMPs and various pathogens [36]. The plants carrying mutation in AtPARG1, but not AtPARG2, showed the elevated elf18 (a 18-amino-acid peptide of EF-Tu)-mediated seedling growth inhibition and phenylpropanoid pigment accumulation, suggesting a negative role of Arabidopsis PARG in certain plant immune responses [37]. Similar to AtPARP1, AtPARG1 also plays a role in plant drought, osmotic and oxidative stress tolerance [38]. In contrast to the extensive research efforts on PARPs/PARGs in animal systems, the biochemical activities and molecular actions of plant PARPs/PARGs remain poorly characterized.

To elucidate the signaling networks regulating immune gene activation, we developed a sensitive genetic screen with an ethyl methanesulfonate (EMS)-mutagenized population of Arabidopsis transgenic plants carrying a luciferase reporter gene under the control of the FRK1 promoter (pFRK1::LUC). The FRK1 (flg22-induced receptor-like kinase 1) gene is a specific and early immune responsive gene activated by multiple MAMPs [39], [40]. A series of mutants with altered pFRK1::LUC activity upon flg22 treatment were identified and named as Arabidopsis genes governing immune gene expression (aggie). In this study, we isolated and characterized the aggie2 mutant, which exhibited elevated immune gene expression upon multiple MAMP treatments. Map-based cloning coupled with next generation sequencing revealed that Aggie2 encodes AtPARG1. Extensive biochemical analysis demonstrates that both AtPARP1 and AtPARP2 carry poly(ADP-ribose) polymerase activity, whereas AtPARG1, but not AtPARG2, possesses poly(ADP-ribose) glycohydrolase activity in vivo and in vitro. Significantly, the enzymatic activity of AtPARP2 is enhanced upon flg22 perception, suggesting the potential involvement of protein PARylation in MAMP-triggered immunity. The aggie2 mutation (G450R) occurs at a highly conserved PARG residue which is essential for both Arabidopsis AtPARG1 and human HsPARG enzymatic activity. Consistent with the negative role of AtPARG1 in plant innate immunity, AtPARP1 and AtPARP2 positively regulate immune gene activation and plant resistance to virulent bacterial pathogen infection. Our results indicate that the reversible posttranslational PARylation process mediated by AtPARPs and AtPARGs plays a crucial role in mounting successful innate immune responses upon MAMP perception in Arabidopsis.

Results

The aggie2 mutant displays enhanced immune gene expression

The aggie2 mutant isolated from a genetic screen of the EMS-mutagenized pFRK1::LUC transgenic plants exhibits elevated FRK1 promoter activity upon flg22 treatment compared to its parental line, pFRK1::LUC (WT) (Fig. 1A). The elevated luciferase activity in the aggie2 mutant was observed over a 48-hr time course period upon flg22 treatment (Fig. 1B). Notably, the aggie2 mutant did not display detectable enhanced FRK1 promoter activity in the absence of flg22 treatment, suggesting its specific regulation in plant defense. In addition to flg22, other MAMPs, including elf18, LPS, PGN and fungal chitin, also elicited the enhanced FRK1 promoter activity in the aggie2 mutant (Fig. 1C), indicating that Aggie2 functions as a convergent component downstream of multiple MAMP receptors. Consistently, the aggie2 mutant displayed the enhanced FRK1 promoter activity in response to the non-pathogenic bacterium Pseudomonas syringae pv. tomato (Pst) DC3000 hrcC defective in type III secretion of effectors, and a non-adaptive bacterium P. syringae pv. phaseolicola NPS3121 (Fig. 1D). The pathogenic bacterium Pst DC3000 failed to activate pFRK1::LUC, likely due to the suppression function of multiple effectors secreted from virulent bacterium [40]. Pathogen infection or purified MAMPs could induce callose deposits in leaves or cotyledons of Arabidopsis, which has emerged as an indicator of plant immune responses [41]. We compared callose deposits by aniline blue staining in WT and aggie2 mutant plants upon flg22 treatment. The aggie2 mutant deposited more callose than WT plants 12 hr after flg22 treatment, and the size of each callose deposit appeared bigger in the aggie2 mutant than that in WT plants (Fig. 1E).

We also detected MAPK activation and ROS production, two early events triggered by multiple MAMPs, in WT and aggie2 mutant. The flg22-induced MAPK activation detected by an α-pERK antibody did not show significant and reproducible difference in WT and aggie2 seedlings (Fig. 1F), suggesting that Aggie2 acts either independently or downstream of MAPK cascade. The flg22-induced ROS burst appeared to be similar in the aggie2 mutant compared to that in WT plants (Fig. 1G). We did not observe reproducible disease alternation in the aggie2 mutant compared to WT plants in response to Pst DC3000 infection either by hand-infiltration or spray-inoculation with various inoculums and conditions (S3A Fig.). Among 7 times of disease assays with Pst DC3000 hand-infiltration, we observed that aggie2 was slightly more resistant than WT plants for 4 times, whereas we did not see the significant difference between aggie2 and WT for other 3 times (S3A Fig). By contrast, the aggie2 mutant showed enhanced susceptibility to a necrotrophic fungus Botrytis cinerea compared to WT plants as evidenced by symptom development and lesion progression after infection (S3B Fig).

We further detected endogenous FRK1 expression in flg22-treated seedlings of WT and aggie2 mutant with quantitative reverse transcription-polymerase chain reaction (qRT-PCR) analysis. The FRK1 expression was significantly elevated in the aggie2 mutant compared to that of WT pFRK1::LUC transgenic plants at both 30 min and 90 min after flg22 treatment (Fig. 1H). Similarly, the expression of several other early MAMP marker genes, including MYB15 and At2g17740 was also enhanced in the aggie2 mutant (Fig. 1H). Taken together, the results indicate that Aggie2 negatively regulates the expression of certain flg22-induced genes.

Aggie2 encodes a putative poly(ADP-ribose) glycohydrolase

To isolate the causative mutation in aggie2, we crossed aggie2 (in the Col-0 accession background) with the Ler accession and mapped aggie2 to an 88 kilobase pair (kb) region between markers F20M17 and F22D22 on Chromosome 2 (Fig. 2A). We then performed Illumina whole genome sequencing of aggie2 and WT pFRK1::LUC transgenic plants. The comparative sequence analysis identified a G to A mutation at the position 1348 bp of At2g31870 within this 88 kb region. The mutation was further confirmed by Sanger sequencing of the genomic DNA of At2g31870. At2g31870 encodes AtPARG1 and the mutation in the aggie2 mutant causes an amino acid change of Glycine (G) at 450 to Arginine (R) (G450R) (Fig. 2B). The G450 in AtPARG1 resides in a highly conserved region at the C-terminus with unknown function. Notably, this residue is invariable in different species of plants and animals, including Arabidopsis, poplar, tomato, maize, sorghum, rice, moss, rat, mouse, human and fruit fly, suggesting the essential role of this residue in PARG functions (Fig. 2B).

To confirm that the G450R lesion in AtPARG1 is the causative mutation in aggie2, we complemented the aggie2 mutant with a construct carrying AtPARG1 cDNA fused with a FLAG epitope tag under the control of its native promoter (pAtPARG1::AtPARG1-FLAG). Two homozygous T3 transgenic lines, one line with relatively low (C2-3) and another line with moderate (C4-1) expression of AtPARG1-FLAG, were chosen for complementation assays. Both lines restored WT level of pFRK1::LUC activity upon flg22 treatment either imaged with an EMCCD camera (Fig. 2C) or quantified by a luminometer (Fig. 2D), confirming that the enhanced FRK1 promoter activity in aggie2 is caused by the mutation in AtPARG1. We also isolated T-DNA insertion line of AtPARG1, parg1-1 (SALK_147805) and parg1-2 (SALK_116088), and examined flg22-induced immune gene activation. Similar to the aggie2 mutant, parg1-1 and parg1-2 displayed the elevated activation of FRK1, MYB15 and At2g17740 after flg22 treatment compared to WT Col-0 plants (Fig. 2E). PARP inhibitor disrupted MAMP-induced cell wall lignification [37]. We found that both parg1-1 and aggie2 mutants showed the enhanced accumulation of lignin biosynthesis precursors, O-4-linked-coniferyl and sinapyl aldehydes, upon flg22 treatment by Wiesner staining (Fig. 2F). The complementation line C2-3 restored accumulation of these lignin biosynthesis precursors to the WT level (Fig. 2F). Consistent with a previous report [36], the transcript of AtPARG2, but not AtPARG1, was induced by flg22 treatment (S4A Fig.).

AtPARP1 and AtPARP2 carry poly(ADP-ribose) polymerase activity in vitro

AtPARG1 encodes a putative poly(ADP-ribose) glycohydrolase with a predicated activity to remove poly(ADP-ribose) polymers on the acceptor proteins catalyzed by poly(ADP-ribose) polymerases (PARPs). To elucidate the biochemical activity and function of AtPARGs, we first characterized the function of AtPARPs and established in vivo and in vitro protein PARylation assays. The Arabidopsis genome encodes three PARPs, AtPARP1, AtPARP2 and AtPARP3, with each consisting of a conserved PARP catalytic domain and a variable DNA binding domain (S2 Fig.). AtPARP1 and AtPARP3 carry zinc-finger domains for DNA binding, which is similar with human HsPARP-1, whereas AtPARP2 contains two SAP domains with putative DNA binding activity. The SAP domain was named after scaffold attachment factor A/B (SAF-A/B), apoptotic chromatin condensation inducer in the nucleus (Acinus) and protein inhibitors of activated STAT (PIAS), which all have DNA and chromatin binding ability and regulate chromatin structure and/or transcription [42]. Analysis of their tissue expression pattern suggests that AtPARP1 and AtPARP2 are expressed in leaves, whereas AtPARP3 is primarily expressed in developing seeds (S4B-S4C Fig.). Thus, we focused on AtPARP1 and AtPARP2 for the functional studies.

We first tested whether AtPARP1 and AtPARP2 carry poly(ADP-ribose) polymerase activity with recombinant proteins of AtPARP1 and AtPARP2 fused with Maltose Binding Protein (MBP). In the presence of activated DNA, both AtPARP1 and AtPARP2 could catalyze PARylation reaction by repeatedly transferring ADP-ribose groups from NAD⁺ to itself (auto-PARylation) as appeared a ladder-like smear with high-molecular-weight proteins in a Western blot using an α-PAR antibody which detects the PAR polymers of PARylated proteins (Fig. 3A). Apparently, AtPARP2 exhibited stronger in vitro enzymatic activity than AtPARP1 when detected by α-PAR antibody. The enzymatic activity of AtPARP1 and AtPARP2 was blocked by 3-AB, a competitive inhibitor of PARP (Fig. 3A). The activity of AtPARP2 is comparable with that of human HsPARP-1 (S5A Fig.). In addition, both AtPARP1 and AtPARP2 were able to transfer ADP-ribose from Biotin-NAD⁺ to itself and a relatively discrete band could be detected by horseradish peroxidase (HRP) conjugated streptavidin (Fig. 3B). The specificity of PARP activity was confirmed with 3-AB treatment, which dramatically reduced auto-PARylation. Similar with the observation using α-PAR antibody, AtPARP2 exhibited stronger in vitro enzymatic activity than AtPARP1 when detected by streptavidin-HRP for biotinylated NAD⁺. We further developed a PARylation assay with radiolabeled ³²P-NAD⁺ as the ADP-ribose donor (Fig. 3C). Clearly, both AtPARP1 and AtPARP2 were able to transfer ADP-ribose from ³²P-NAD⁺ to itself as shown with SDS-PAGE autoradiograph (Fig. 3C). The formation of relatively discrete band was likely caused by these assay conditions, which favor synthesis of short polymers due to limited amount of NAD⁺ [43]. Together, the data support that both AtPARP1 and AtPARP2 are active poly(ADP-ribose) polymerases in vitro. It has been shown that human HsPARP-1 could modify linker histone H1 proteins and thereby create a chromatin structure more accessible to RNA polymerase II (RNAPII) to regulate transcription [44]. We further examined whether AtPARP2 was also able to PARylate Arabidopsis histone proteins. As detected with radiolabeled ³²P-NAD⁺, AtPARP2 could PARylate two Arabidopsis histone proteins H1.1 and H1.3 (Fig. 3D). It is possible that Arabidopsis PARPs may use a similar mechanism for transcriptional regulation.

Flg22 induces AtPARP2 activity in vivo

We further developed an in vivo PARylation assay with transiently expressed AtPARP2 tagged with an HA epitope at the C-terminus in Arabidopsis protoplasts. After feeding the cells with ³²P-NAD⁺, the AtPARP2 proteins were immunoprecipitated with an α-HA antibody and separated in SDS-PAGE. A band corresponding to the predicated molecular weight of AtPARP2 was observed with autoradiograph, indicating in vivo AtPARP2 activity (Fig. 3E). This band is specific to AtPARP2 since it was absent in the vector control transfected cells. Strikingly, the flg22 treatment enhanced AtPARP2 in vivo PARylation activity as detected by increased band intensity with autoradiograph. Apparently, the flg22-mediated enhancement of AtPARP2 activity was not due to the increase of protein expression after treatment (Fig. 3E). The data demonstrate that AtPARP2 possesses poly(ADP-ribose) polymerase activity in vivo and AtPARP2-mediated protein PARylation is regulated by flg22 signaling. We further examined AtPARP2-GFP localization with Agrobacterium-mediated Nicotiana benthamiana transient assay. A strong fluorescence signal from AtPARP2-GFP was exclusively detected in the nucleus (Fig. 3F), which is consistent with its potential role in DNA repair, chromatin modulation and transcriptional regulation.

AtPARG1, but not AtPARG2, is a functional PARG enzyme

We next tested whether AtPARG1 and AtPARG2 possess poly(ADP-ribose) glycohydrolase activity (Fig. 4A). We isolated and purified AtPARG1 and AtPARG2 proteins fused with glutathione S-transferase (GST) expressed from E. coli, and established an in vitro PARG assays to examine whether AtPARGs could remove PAR from auto-PARylated AtPARP2 in vitro. As shown in Fig. 4B, AtPARG1 diminished the formation of the ladder-like smear of auto-PARylated AtPARP2 detected in a Western blot with an α-PAR antibody, suggesting the PARG activity of AtPARG1 towards AtPARP2. However, AtPARG2 appeared to be inactive towards auto-PARylated AtPARP2 in this assay (Fig. 4B). Similarly, AtPARG1, but not AtPARG2, could remove PAR polymers from auto-ADP-ribosylated AtPARP2 as detected with ³²P-NAD⁺ autoradiograph (Fig. 4C). We further examined whether AtPARG2 may possess PARG activity specifically towards AtPARP1 but not AtPARP2. As shown in Fig. 4C, AtPARG2 did not remove PAR polymers from auto-ADP-ribosylated AtPARP1. The 6xHistidine (His6)-tagged AtPARG2 also did not display in vitro enzymatic activity (S5B Fig.). Similar to the above assays using in vitro expressed AtPARG1 proteins (Fig. 4B & 4C), the immunoprecipitated AtPARG1 expressed in Arabidopsis protoplasts almost completely removed PAR polymers from in vitro PARylated AtPARP2 (Fig. 4D). Furthermore, AtPARG1 was able to remove PAR polymers from auto-PARylated human HsPARP-1 (S5C Fig.). Similarly, human HsPARG was also able to remove PAR polymers from AtPARP2 (S5D Fig.), suggesting the functional conservation of human and Arabidopsis PARPs/PARGs.

To test whether AtPARGs carry enzymatic activity in vivo, HA-tagged AtPARG1 or AtPARG2 was co-expressed with FLAG-tagged AtPARP2 transiently expressed in Arabidopsis protoplasts. After feeding the protoplasts with ³²P-NAD⁺, AtPARP2 activity was detected with autoradiograph after immunoprecipitation with an α-FLAG antibody. Significantly, co-expression of AtPARG1, but not AtPARG2, substantially removed PAR polymers from in vivo PARylated AtPARP2 (Fig. 4E). The expression level of AtPARG1 and AtPARG2 was similar in protoplasts as detected by an α-HA Western blot (Fig. 4E). Taken together, our data indicate that AtPARG1 has in vivo and in vitro poly(ADP-ribose) glycohydrolase activity and AtPARG2 activity was not detected. Consistently, the parg1 mutant, but not parg2 mutant, accumulated higher PAR polymers than Col-0 with dot blotting of nuclear proteins by α-PAR antibody (S5E Fig.). Subcellular localization study indicates that AtPARG1-GFP and AtPARG2-GFP reside mainly in nucleus, but also in plasma membrane and cytoplasm when transiently expressed in Arabidopsis protoplasts (Fig. 4F).

Notably, AtPARG1 protein possesses a PARG signature motif with the conserved sequence of “GGG-X₇-QEE”. Mutation of E273 (the last E in the signature motif) in AtPARG1 to glycine (E273G) blocked its enzymatic activity, implicating the importance of this signature motif in PARG enzymatic activity (Fig. 5A). Examination of AtPARG2 sequence revealed that AtPARG2 has a polymorphism in the PARG signature motif. Instead of the conserved sequence “GGG-X₇-QEE”, AtPARG2 possesses “GGL-X₇-QEE” (Fig. 4A and 5A). The importance of this residue was shown by that the mutation of G264 (third G in the signature motif) in AtPARG1 to leucine (G264L) blocked its PARG enzymatic activity (Fig. 5A). We further determined whether lack of enzymatic activity of AtPARG2 (Fig. 4B, 4C & 4E) is due to this polymorphism in the PARG signature motif. We mutated leucine (L275) in AtPARG2 to glycine and generated the conserved “GGG-X₇-QEE” motif. However, AtPARG2L275G mutant with a perfectly conserved PARG signature motif still did not show any detectable poly(ADP-ribose) glycohydrolase activity (Fig. 5A). The data suggest that the polymorphism of the PARG signature motif in AtPARG2 is not the sole determinant of its lack of detectable enzymatic activity and additional polymorphisms/deletions also account for its loss of PARG functions. There are only about 52% amino acid identity and 66% similarity between AtPARG1 and AtPARG2 (S6 Fig.).

The aggie2 mutation occurs at a conserved and essential PARG residue

We further addressed whether the aggie2 (G450R) mutation affected its PARG activity. Significantly, the aggie2 (G450R) mutant of AtPARG1 completely abolished its enzymatic activity detected by either α-PAR antibody (Fig. 4B) or ³²P-NAD⁺ autoradiograph-based assay (Fig. 4C). Notably, the G450 in AtPARG1 is highly conserved among PARGs of different species (Fig. 2B). Interestingly, the corresponding mutation in human HsPARG (G867R) also abolished its activity towards HsPARP-1 and AtPARP2, suggesting the essential role of this highly conserved residue in different PARGs (Fig. 5B and S5D Fig.). The phenylalanine (F) at position 227 in bacterium Thermomonospora curvata PARG is implicated in positioning the terminal ribose and the mutation of which rendered the enzyme inactive [28]. Surprisingly, mutation of the corresponding residue F457 to glycine (F457G) in AtPARG1 did not affect its enzymatic activity (Fig. 5A), suggesting a possible distinct function mediated by this residue in different PARGs and potentially divergent evolution.

AtPARPs positively regulate plant immunity

We tested the involvement of AtPARPs in plant innate immunity and immune gene activation. Because of the potential functional redundancy of AtPARP1 and AtPARP2 [30], [31], we performed disease assay and analyzed defense gene expression in atparp1atparp2 (atparp1/2) double mutant. The atparp1/2 mutant plants were more susceptible to virulent P. syringae pv. maculicola ES4326 (Psm) infection compared to WT plants as indicated by more than 10 fold increase of bacterial growth in the atparp1/2 mutant (Fig. 6A). The disease symptom development was more pronounced in the atparp1/2 mutant than WT plants (Fig. 6A). Similarly, the atparp1/2 mutant plants showed the enhanced susceptibility with bacterial growth and symptom development to the infections by Pst DC3000 and a less virulent bacterium Pst DC3000ΔavrPtoavrPtoB (Fig. 6B & S7 Fig.). In addition, the atparp1/2 mutant plants showed the reduced induction of MAMP marker genes, including FRK1 and At2g17740, compared to WT plants at 90 min after flg22 treatment (Fig. 6C). Together, these data indicate that AtPARP1 and AtPARP2 are positive regulators in plant immunity and defense gene activation to bacterial infections.

Discussion

Protein PARylation mediated by PARPs and PARGs is an important, but less understood posttranslational modification process implicated in the regulation of diverse cellular processes and physiological responses [26]. In this study, an unbiased genetic screen revealed that Arabidopsis AtPARG1 plays an important role in regulating immune gene expression upon pathogen infection. We established and performed extensive in vitro and in vivo biochemical assays of PARP and PARG enzymatic activities. We have shown for the first time that Arabidopsis AtPARP1 and AtPARP2 are able to transfer ADP-ribose moieties from NAD⁺ to itself and acceptor proteins in vitro and in vivo. Thus, they are bona fide poly(ADP-ribose) polymerases. Interestingly, in contrast to their mammalian counterparts, AtPARP2 is more enzymatically active than AtPARP1. Significantly, MAMP perception promotes substantial enhancement of AtPARP2 enzymatic activity in vivo, reconciling the biological importance of PARPs/PARGs in regulating immune gene expression. AtPARG1, but not AtPARG2, is able to remove PAR polymers from PARylated proteins in vivo and in vitro and it is a bona fide poly(ADP-ribose) glycohydrolase. The Arabidopsis parg1 (aggie2) mutant plants exhibited elevated expression of several MAMP-induced genes and callose deposition. Conversely, the Arabidopsis atparp1/2 mutant showed reduced expression of MAMP-induced genes and enhanced susceptibility to virulent Pseudomonas infections. Thus, the data suggest that protein PARylation positively regulates certain aspects of plant immune responses. Notably, the viability and normal growth of Arabidopsis parp and parg null mutants represent a unique opportunity to study protein PARylation regulatory mechanisms in diverse biological processes at the whole organismal level.

Our results lend support to a previous study that treatment of pharmacological inhibitor of PARPs, 3-AB, disrupted elf18- and/or flg22-induced callose and lignin deposition, pigment accumulation and phenylalanine ammonia lyase activity [37]. However, the flg22-induced defense genes (FRK1 and WRKY29) were not affected by 3-AB treatment [37]. Our study with Arabidopsis parg and parp genetic mutants revealed a previously unrecognized function of protein PARylation in regulating immune gene expression upon pathogen infection. This is consistent with the general role of human PARPs and PARG in transcriptional regulation and chromatin modification [43], [45] and further substantiates the hypothesis that plant PARPs could ameliorate the cellular stresses caused by antimicrobial defenses (e.g. the effects of elevated ROS levels) [23]. Interestingly, ADP-ribosylation has also been exploited by pathogens as a means to quell plant immunity. Two Pseudomonas syringae effectors, HopU1 and HopF2, mono-ADP-ribosylate RNA-binding protein GRP7 and MAPK kinase MKK5 respectively, and interfere with their activities in plant defense transcription regulation and signaling [46], [47].

Unlike mammals and most other animals that encode a single PARG gene, the Arabidopsis genome encodes two adjacent PARG genes, AtPARG1 and AtPARG2, as well as a pseudogene At2g31860. Surprisingly, only AtPARG1, but not AtPARG2, possesses detectable poly(ADP-ribose) glycohydrolase activity in vitro and in vivo with our extensive biochemical assays. Sequence analysis identified a polymorphism in the conserved PARG signature motif “GGG-X₇-QEE”, where the third G is replaced with an L in AtPARG2. The PARG signature motif is absolutely required for its enzymatic activity as mutations at this motif in AtPARG1 completely abolished its activity. However, creation of the conserved signature motif in AtPARG2 was unable to gain its PARG activity suggesting that other polymorphisms in AtPARG2 are also responsible for its lack of enzymatic activity. Consistent with our biochemical assays, the PAR polymer concentration was much higher in atparg1 mutant than that in WT plants. A similar conclusion was reached on tej mutant, which carries a G262E mutation in the invariable signature motif of AtPARG1 [35]. The atparg1, but not atparg2 mutant, affected elf18-induced seedling growth inhibition and pigment formation, and sensitivity to DNA-damaging agent [37]. Interestingly, AtPARG2 is substantially induced in multiple plant-pathogen interactions [36] and it is required for plant resistance to B. cinerea infections [37]. Thus, despite of lacking detectable enzymatic activity, AtPARG2 may still play certain role in plant immunity. It is possible that AtPARG2 may regulate AtPARG1 activity. It is also possible that AtPARG2 has evolved novel functions in plant immune responses. Several other plant species, including rice, poplar, tomato and maize, are also predicted to encode multiple PARGs [23] (S8 Fig.). Unlike Arabidopsis PARGs, different PARG members in other species have invariant signature motif. For example, all three PARGs in poplar contain GGG-X₇-QEE signature motif (S8 Fig.). However, a few other species such as Eutrema salsugineum, Capsella rubella, Phaseolus vulgaris, Oikopleura dioica, and Xenopus laevis, contain PARGs with an AtPARG2-like signature GGL-X₇-QEE. It remains unknown how many PARGs are enzymatic active in the species with multiple PARGs.

Although there are 17 PARPs in mammals, the parp-1parp-2 double mutant mice are not viable and die at the onset of gastrulation, suggesting the essential role of protein PARylation during early embryogenesis [48]. The lethality of parp-1parp-2 double mutant mice might be due to genomic instability. However, Arabidopsis atparp1/2 double mutant is largely morphologically similar with WT plants and does not display any obvious growth defects. Although Arabidopsis atparp1/2 double mutant was hypersensitive to genotoxic stress, they did not have significant changes in telomere length nor end-to-end chromosome fusions [30]. Albeit mainly expressed in developing seeds, AtPARP3 may have redundant functions with AtPARP1 and AtPARP2 in maintaining genome stability. It remains interesting whether atparp1/2/3 triple mutant will exert abnormal plant growth and development. Consistent with the essential function of PARylation during embryogenesis, PARG-deficient mice and Drosophila are embryonic lethal which is probably due to the accumulation of PAR polymers and uncontrolled PAR-dependent signaling [49], [50]. The normal plant growth phenotype of atparg1 mutant might be due to the redundant function of AtPARG2. However, our extensive biochemical analysis indicates that AtPARG1, but not AtPARG2, accounts for most of PARG enzymatic activity. As AtPARG1 and AtPARG2 reside next to each other on the same chromosome, it is challenging to generate the double mutant. It remains possible that other PAR-degrading enzymes with distinct sequences exist in Arabidopsis. In vertebrate, ADP-ribosyl hydrolase 3 (ARH3), a structurally distinct enzyme from PARG, could also degrade PAR polymers associated with the mitochondrial matrix [26].

We observed that AtPARP2 activity was rapidly and substantially stimulated by flg22 treatment. In line with this observation, it has been shown that bacterial infections induced the increase of PAR polymers in Arabidopsis [37]. It is well established that damaged DNA stimulates PARP activity. Recent studies have shown that pathogen treatments induce DNA damage [51], [52], which could potentially serve as a trigger to activate PARP. Treatments with virulent or avirulent Pst strains for hours could induce DNA damage in Arabidopsis as detected by abundance of histone γ-H2AX, a sensitive indicator of DNA double-strand breaks or by DNA comet assays [51]. Prolonged pathogen treatment is often accompanied with the elevated accumulation of plant defense hormone salicylic acid (SA). It has also been shown that SA can also trigger DNA damage in the absence of a genotoxic agent [53]. However, treatments of flg22 or elf18 did not induce detectable DNA damage [51]. In addition, flg22-mediated stimulation of AtPARP2 activity occurs rather rapidly and within 30 min after treatment. Apparently, flg22 signaling could directly activate AtPARP2. It is well known that human HsPARP-1 is regulated by different posttranslational modification processes, such as phosphorylation, ubiquitination, SUMOylation and cleavage [22]. HsPARP-1 could be activated by phosphorylated MAPK ERK2 in a broken DNA-independent manner, thereby enhancing ERK-induced Elk1 phosphorylation, core histone acetylation, and transcription of the Elk1-target genes [54]. MAPK cascade plays a central role functioning downstream of multiple MAMP receptors. It will be interesting to test whether flg22-activated MAPKs directly modulate PARP and/or PARG activities.

Our genetic and biochemical analyses revealed that PARP/PARG-mediated PAR dynamics regulates immune gene expression in Arabidopsis. Mammalian PARPs/PARG regulate gene expression through a variety of mechanisms including modulating chromatin, functioning as transcriptional co-regulators and mediating DNA methylation [55]. PARylation of histone lysine demethylase KDM5B maintains histone H3 lysine 4 trimethyl (H3K4me3), a histone mark associated with active promoters, by inhibiting KDM5B demethylase activity and interactions with chromatin. In addition, HsPARP-1 is able to promote exclusion of H1 and opening of promoter chromatin, which collectively lead to a permissive chromatin environment that allows loading of the RNAPII machinery [45]. HsPARG is also able to promote the formation of a chromatin environment suitable for retinoic acid receptor (RAR)-mediated transcription by removing PAR polymer from PARylated H3K9 demethylase KDM4D/JMJD2D thereby activating KDM4D/JMJD2D to inhibit H3K9me2, a histone mark associated with transcriptional repression [43]. Arabidopsis PARPs and PARGs are localized in the nucleus, and AtPARP2 could PARylate Histone H1. It is plausible to speculate that similar modes of action of protein PARylation-mediated transcriptional regulation exist in plants. Future identification of PARP/PARG targets (promoters and proteins) and PAR-associated proteins, especially during plant immune responses, will elucidate how protein PARylation modulates plant immune gene expression.

Materials and Methods

Plant and pathogen materials and growth conditions

Arabidopsis accession Col-0, pFRK1::LUC transgenic plants, aggie2 mutant, atparg1-1 (SALK_147805), atparg1-2 (SALK_16088), atparg2 (GABI-Kat 072B04), atparp1/atparp2 (GABI-Kat 692A05/SALK_640400), pPARG1::PARG1-FLAG transgenic plants were grown in soil (Metro Mix 366) at 23°C, 60% humidity and 75 µE m⁻²s⁻¹ light with a 12-hr light/12-hr dark photoperiod. Four-week-old plants were used for protoplast isolation and transient expression assays according to the standard procedure [56]. Seedlings were germinated on ½ Murashige and Skoog (MS) plate containing 1% sucrose, 0.8% Agar and grown at 23°C and 75 µE m^-2s⁻¹ light with a 12-hr light/12-hr dark photoperiod for 12 days, transferred to a 6-well tissue culture plate with 2 ml H₂O for overnight, and then treated with 100 nM flg22 or H₂O for indicated time.

Pseudomonas syringae pv. tomato (Pst) DC3000, hrcC, ΔavrPtoavrPtoB, P. syringae pv. maculicola ES4326 (Psm), or P. syringae pv. phaseolicola NPS3121 strains were cultured overnight at 28°C in the KB medium with 50 µg/ml rifampicin or streptomycin. Bacteria were harvested by centrifugation, washed, and adjusted to the desired density with 10 mM MgCl₂. Leaves of 4-week-old plants were hand-infiltrated with bacterial suspension using a 1-ml needleless syringe and collected at the indicated time for luciferase activity or bacterial growth assays. To measure bacterial growth, two leaf discs were ground in 100 µl H₂O and serial dilutions were plated on TSA medium (1% Bacto tryptone, 1% sucrose, 0.1% glutamic acid, 1.5% agar) with appropriate antibiotics. Bacterial colony forming units (cfu) were counted 2 days after incubation at 28°C. Each data point is shown as triplicates. Botrytis cinerea strain BO5 was cultured on Potato Dextrose Agar (Difco) and incubated at room temperature. Conidia were re-suspended in distilled water and spore concentration was adjusted to 2.5 × 10⁵ spores/ml. Gelatin (0.5%) was added to conidial suspension before inoculation. Leaves of six-week-old plants were drop-inoculated with B. cinerea at the concentration of 2.5 × 10⁵ spores/ml. Lesion size was measured 2 days post-inoculation.

Mutant screening, map-based cloning and next generation sequencing

The pFRK1::LUC construct in a binary vector was transformed into Arabidopsis Col-0 plants. The homozygous transgenic plants with flg22-inducible pFRK1::LUC were selected for mutagenesis. The seeds were mutagenized with 0.4% ethane methyl sulfonate (EMS). Approximately 6,000 M2 seedlings were screened for their responsiveness to flg22 treatment. The seedlings were germinated in liquid ½ MS medium for 14 days, and then transferred to water for overnight and treated with 10 nM flg22. After 12 hr flg22 treatment, the individual seedlings were transferred to a 96-well plate, sprayed with 0.2 mM luciferin and kept in dark for 20 min. The bioluminescence from induced pFRK1::LUC expression was recorded by a luminometer (Perkin Elmer, 2030 Multilabel Reader, Victor X3). The candidate mutants with altered flg22 responsiveness were recovered on ½ MS plate for 10 days, and then transferred to soil for seeds.

The aggie2 mutant was crossed with Arabidopsis Ler accession, and an F2 population was used for map-based cloning. Mapping with 270 F2 plants with aggie2 mutant phenotype placed the causal mutation in an 88 kb region between marker F20F17 and F22D22 on chromosome 2. The aggie2 genomic DNA was sequenced with the 100 nt paired-end sequencing on an Illumina HiSeq 2000 platform at Texas AgriLife Genomics and Bioinformatics Service (TAGS) (College Station, TX, USA). Ten-fold genome coverage was obtained with 11M reads. The Illumina reads were analyzed using CLC Genomics Workbench 6.0.1 software. By mapping to Col-0 genomic sequence (TAIR10 release), SNPs were identified as candidates of aggie2 mutation. In the aforementioned 88 kb region, a G to A mutation at the position of 1348 nt of At2g31870 was identified with 100% frequency. The mutation was confirmed by Sanger sequencing of aggie2 genomic DNA.

Plasmid constructs for protoplasts and transgenic plants

The AtPARP1, AtPARP2, AtPARG1, AtPARG2 and Histone H1.1 (At1g06760) genes were amplified from Arabidopsis Col-0 cDNA and cloned into a plant transient expression vector (pHBT vector) with an HA, FLAG or GFP epitope tag at the C-terminus via restriction sites NcoI or BamHI and StuI respectively. The oligos used to amplify aforementioned cDNAs are listed in S1 Table. The target genes were confirmed by Sanger sequencing. The cloned genes in plant expression vector were then sub-cloned into protein fusion vectors, pGEX-4T (Pharmacia, USA), pMAL-c (NEB, USA) or pET28a (EMD Millipore, USA), for protein expression in bacteria. For Histone H1.3 (At2g18050), we ordered cDNA from ABRC (G13366) and cloned it into a modified pMAL-c via SfiI site. Point mutations were introduced by site-directed mutagenesis PCR. The AtPARG1 promoter (1163 bp upstream of start codon ATG) was amplified from the genomic DNA of Col-0 and digested with KpnI and NcoI. The AtPARG1-FLAG-NOS terminator fragment was released from pHBT-AtPARG1-FLAG via NcoI and EcoRI digestion. The two fragments were ligated and sub-cloned into a binary vector, pCAMBIA2300 via KpnI and EcoRI sites to yield expression construct (pAtPARG1::AtPARG1-FLAG). The resulting binary vector was transformed into aggie2 via Agrobacterium-mediated transformation.

The primers for cloning and point mutations were listed in the S1 Table.

In vitro and in vivo PARP and PARG assays

Expression and purification of GST, His6 and MBP fusion proteins were performed according to the manufacturer's manuals. For in vitro auto-PARylation reaction, 1.2 µg of MBP-AtPARP2 or MBP-AtPARP1 proteins were incubated in a 20 µl reaction with 1 × PAR reaction buffer (50 mM Tris-HCl, pH8.0, 50 mM NaCl) with 0.2 mM NAD⁺, and 1 × activated DNA (Trevigen, USA). To inhibit PAR reaction, 2.5 mM PARP inhibitor, 3-Aminobenzamide (3-AB, Sigma, USA), was added to the reaction. The reactions were kept at room temperature for 30 min and stopped by adding SDS loading buffer. To detect PARG activity, about 1.0 µg of purified GST, GST-AtPARG1 or GST-AtPARG2 proteins together with 2.5 mM 3-AB were added to auto-PARylated AtPARP2 proteins derived from the above PAR reactions and incubated at room temperature for another 30 min. PARylated proteins were separated in 7.5% SDS-PAGE and detected with an α-PAR polyclonal antibody (Trevigen, USA). For Biotin NAD⁺ PAR assay, 25 µM Biotin-NAD⁺ (Trevigen, USA) was added to replace NAD⁺ in the reaction described above. The PAR polymer formation was detected by Streptavidin-HRP (Pierce, USA). For in vitro ³²P-NAD⁺-mediated PAR assays, 1.0 µg of MBP-AtPARP2 or MBP-AtPARP1 proteins were incubated in a 20 µl reaction in the buffer containing 50 mM Tris-HCl, pH8.0, 4 mM MgCl₂, 300 mM NaCl, 1 mM DTT, 0.1 µg/ml BSA, 1 × activated DNA, 1 µCi ³²P-NAD⁺ (Perkin Elmer, USA) and 100 nM cold NAD⁺ for 30 min at room temperature. For Histone PARylation assays, 2.0 µg of MBP-H1.1 or MBP-H1.3 proteins were added in the above reactions. The radiolabeled proteins were separated in SDS-PAGE and visualized by autoradiography.

For in vivo PAR assays, 500 µl Arabidopsis protoplasts at the concentration of 2 × 10⁵/ml were transfected with 100 µg of plasmid DNA of pHBT-AtPARP2-HA. After 12 hr incubation, the protoplasts were treated with 100 nM flg22 for 30 min and fed with 1 µCi ³²P-NAD⁺ for 1 hr. The protoplasts were then lysed in IP buffer (50 mM Tris-HCl, pH7.5, 150 mM NaCl, 5 mM EDTA, 1% Triton, 1 × protease inhibitor, 1 mM DTT, 2 mM NaF and 2 mM Na₃VO₄) and the AtPARP2-HA proteins were immunoprecipitated with α-HA antibody (Roche, USA) and protein-G-agarose (Roche, USA) in a shaker for 3 hr at 4°C. In vivo PARylated proteins enriched on the beads were then separated in 10% SDS-PAGE and visualized by autoradiography. For in vivo PARG assay, AtPARG1-HA or AtPARG2-HA plasmid DNA was co-transfected with AtPARP2-FLAG plasmid DNA into protoplasts, and expressed for 12 hr. The protoplasts were fed with ³²P-NAD⁺ and subjected to immunoprecipitation as described above. The AtPARP2-FLAG proteins were immunoprecipitated with α-FLAG agarose gel (Sigma, USA), separated in 10% SDS-PAGE and visualized by autoradiography. The expression of AtPARPs and AtPARGs was detected with Western blot (WB) using the corresponding antibodies.

Detection of PAR polymers from protein extract of nuclei

The 12-day old seedlings grown on ½ MS plates were harvested and ground into fine powder in liquid nitrogen. Isolation of nuclei with Honda buffer was performed according to published procedure [57]. Nuclear proteins were released in 1xPBS buffer with 1% SDS and spotted on nitrocellulose membrane. The protein loaded on the membrane was normalized by using α-Histone H3 antibody (Abcam, USA), and the PAR polymers were detected by α-PAR antibody. The relative PAR level was determined by calculating the ratio of PAR signal to Histone H3 signal after quantification of hybridization intensity with ImageJ software.

RNA isolation and RT-PCR

For RNA isolation, 12-day-old seedlings grown on ½ MS plate were transferred to 2 ml H₂O in a 6-well plate to recover for 1 day, and then treated with 100 nm flg22 for 30 or 90 min. RNA was extracted using TRIzol reagent (Life Technologies, USA) and quantified with NanoDrop. The RNA was treated with RQ1 RNase-free DNase I (Promega, USA) for 30 min at 37°C, and then reverse transcribed with M-MuLV Reverse Transcriptase (NEB, USA). Real-time RT-PCR was carried out using iTaq Universal SYBR Green Supermix (Bio-Rad, USA) on 7900HT Fast Real-Time PCR System (Applied Biosystems, USA). The primers used to detect specific transcript by real-time RT-PCR are listed in S2 Table.

Callose deposition

Leaves of six-week-old plants grown in soil were hand-inoculated with 0.5 µM flg22 or H₂O for 12 hr. The leaves were then transferred into FAA solution (10% formaldehyde, 5% acetic acid and 50% ethanol) for 12 hr, de-stained in 95% ethanol for 6 hr, washed twice with ddH₂O, and incubated in 0.01% aniline blue solution (150 mM KH₂PO4, pH 9.5) for 1 hr. The callose deposits were visualized with a fluorescence microscope. Callose deposits were counted using ImageJ 1.43U software (http://rsb.info.nih.gov/ij/).

Lignin deposition

Leaves of six-week-old plants grown in soil were surface-sterilized by 70% ethanol, rinsed with H₂O and incubated with 100 nM flg22 or H₂O for 12 hr. The leaves were then de-stained in 95% ethanol with 2% chloroform for 12 hr and 95% ethanol for 6 hr, washed twice with 95% ethanol, and incubated in 2% phloroglucinol solution (20% ethanol, 20% HCl) for 5 min. The images were scanned by HP officejet Pro 8600 Premium.

MAPK assay

Ten-day-old seedlings germinated on ½MS plate were transferred to 2ml H₂O in a 6-well plate to recover for 1 day, and then treated with 100 nM flg22 for 5, 15 or 45 min. The seedlings were grinded in IP buffer. The cleared lysate was mixed with SDS sample buffer and loaded onto 12.5% SDS-PAGE. Activated MAPKs were detected with α-pErk1/2 antibody (Cell Signaling, USA).

ROS analyses

ROS burst was determined by a luminol-based assay. At least 10 leaves of four-week-old Arabidopsis plants for each genotype were excised into leaf discs of 0.25 cm², followed by an overnight incubation in 96-well plate with 100 µl of H₂O to eliminate the wounding effect. H₂O was replaced by 100 µl of reaction solution containing 50 µM luminol and 10 µg/ml horseradish peroxidase (Sigma, USA) supplemented with or without 100 nM flg22. The measurement was conducted immediately after adding the solution with a luminometer (Perkin Elmer, 2030 Multilabel Reader, Victor X3), with a 1.5 min interval reading time for a period of 30 min. The measurement values for ROS production from 20 leaf discs per treatment were indicated as means of RLU (Relative Light Units).

GFP localization assay

Arabidopsis protoplasts were transfected with various GFP-tagged pHBT constructs as indicated in the figures. Fluorescence signals in the protoplasts were visualized under a confocal microscope 12 hr after transfection. To construct 35S::AtPARP2-GFP binary plasmid for Agrobacterium-mediated transient assay, the NcoI-PstI fragment containing AtPARP2-GFP was released from pHBT-35S::AtPARP2-GFP and ligated into pCB302 binary vector. For tobacco transient expression, Agrobacterium tumefaciens strain GV3101 containing pCB302-35S::AtPARP2-GFP was cultured at 28°C for 18 hr. Bacteria were harvested by centrifugation at a speed of 3500 rpm and re-suspended with infiltration buffer (10 mM MES pH = 5.7, 10 mM MgCl₂, 200 µM acetosyringone). Cell solution at OD600 = 0.75 was used to infiltrate 3-week-old Nicotiana benthamiana leaves. Fluorescence signals were detected 2 days post-infiltration. Fluorescence images were taken with Nikon-A1 confocal laser microscope systems and images were processed using NIS-Elements Microscope Imaging Software. The excitation lines for imaging GFP, RFP and chloroplast were 488, 561 and 640 nm, respectively.

Supporting Information

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