orcid.org/0000-0002-7355-3111 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USAFor correspondence (e-mail dmchiniquy@lbl.gov).Search for more papers by this authorWilliam Underwood, orcid.org/0000-0002-9920-8228 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorJason Corwin, Department of Plant Sciences, University of California, Davis, CA, 95616 USASearch for more papers by this authorAndrew Ryan, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorHeidi Szemenyei, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorCandice C. Lim, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorSolomon H. Stonebloom, Joint BioEnergy Institute, Emeryville, CA, 94608 USASearch for more papers by this authorDevon S. Birdseye, Joint BioEnergy Institute, Emeryville, CA, 94608 USASearch for more papers by this authorJohn Vogel, orcid.org/0000-0003-1786-2689 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Joint Genome Institute, Walnut Creek, CA, 94598 USASearch for more papers by this authorDaniel Kliebenstein, Department of Plant Sciences, University of California, Davis, CA, 95616 USASearch for more papers by this authorHenrik V. Scheller, orcid.org/0000-0002-6702-3560 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Joint BioEnergy Institute, Emeryville, CA, 94608 USA Environmental Genomics and Systems Biology Division, Lawrence Berkeley National Laboratory, Berkeley, CA, 94720 USASearch for more papers by this authorShauna Somerville, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this author orcid.org/0000-0002-7355-3111 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USAFor correspondence (e-mail dmchiniquy@lbl.gov).Search for more papers by this authorWilliam Underwood, orcid.org/0000-0002-9920-8228 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorJason Corwin, Department of Plant Sciences, University of California, Davis, CA, 95616 USASearch for more papers by this authorAndrew Ryan, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorHeidi Szemenyei, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorCandice C. Lim, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this authorSolomon H. Stonebloom, Joint BioEnergy Institute, Emeryville, CA, 94608 USASearch for more papers by this authorDevon S. Birdseye, Joint BioEnergy Institute, Emeryville, CA, 94608 USASearch for more papers by this authorJohn Vogel, orcid.org/0000-0003-1786-2689 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Joint Genome Institute, Walnut Creek, CA, 94598 USASearch for more papers by this authorDaniel Kliebenstein, Department of Plant Sciences, University of California, Davis, CA, 95616 USASearch for more papers by this authorHenrik V. Scheller, orcid.org/0000-0002-6702-3560 Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Joint BioEnergy Institute, Emeryville, CA, 94608 USA Environmental Genomics and Systems Biology Division, Lawrence Berkeley National Laboratory, Berkeley, CA, 94720 USASearch for more papers by this authorShauna Somerville, Department of Plant and Microbial Biology, University of California, Berkeley, CA, 94720 USA Energy Biosciences Institute, Berkeley, CA, 94720 USASearch for more papers by this author Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URLShare a linkShare onEmailFacebookTwitterLinked InRedditWechat Summary Powdery mildew (Golovinomyces cichoracearum), one of the most prolific obligate biotrophic fungal pathogens worldwide, infects its host by penetrating the plant cell wall without activating the plant\'s innate immune system. The Arabidopsis mutant powdery mildew resistant5 (pmr5) carries a mutation in a putative pectin acetyltransferase gene that confers enhanced resistance to powdery mildew. Here, we show that heterologously expressed PMR5 protein transfers acetyl groups from [14C]-acetyl-CoA to oligogalacturonides. Through site-directed mutagenesis, we show that three amino acids within a highly conserved esterase domain in putative PMR5 orthologs are necessary for PMR5 function. A suppressor screen of mutagenized pmr5 seed selecting for increased powdery mildew susceptibility identified two previously characterized genes affecting the acetylation of plant cell wall polysaccharides, RWA2 and TBR. The rwa2 and tbr mutants also suppress powdery mildew disease resistance in pmr6, a mutant defective in a putative pectate lyase gene. Cell wall analysis of pmr5 and pmr6, and their rwa2 and tbr suppressor mutants, demonstrates minor shifts in cellulose and pectin composition. In direct contrast to their increased powdery mildew resistance, both pmr5 and pmr6 plants are highly susceptibile to multiple strains of the generalist necrotroph Botrytis cinerea, and have decreased camalexin production upon infection with B.cinerea. These results illustrate that cell wall composition is intimately connected to fungal disease resistance and outline a potential route for engineering powdery mildew resistance into susceptible crop species. Introduction Plant cell walls are a dynamic terrain, undergoing frequent construction and deconstruction to meet the needs of the developing plant (Somerville etal., 2004; Keegstra, 2010). Cell walls must be malleable for growth, sturdy for water transport, flexible to allow movement without breakage and remain able to be deconstructed at precisely timed ripening stages (Cosgrove, 2005). Modifications of specific cell wall polysaccharides, including acetylation and methylation, allow for this plasticity during growth and development, providing flexibility during growth and strength when growth is complete. Pectins, a diverse group of cell wall polysaccharides rich in galacturonic acid residues, are often acetylated or methylated to various degrees, a process that prevents the formation of calcium-mediated interactions, which makes the cell wall more expandable (Liners etal., 1992; Harholt etal., 2010). Upon pathogen attack, pectin fragments elicit a defense response in plants (Lionetti etal., 2012; Bellincampi etal., 2014), making pectin an essential component of the immunity detection system (Ferrari etal., 2013). Specifically, oligogalacturonides (OGAs), which are fragments of the homogalacturonan domains of pectin, are released by the cell wall as degradation byproducts following exposure to pathogen endopolygalacturonases and endopectate lyases. Powdery mildew is a disease caused by biotrophic fungal pathogens that affects over 9000 dicot species and 600 monocot species, including many economically valuable crops; with such a wide host range, it is one of the most prevalent plant pathogens worldwide (Glawe, 2008). To gain access to the host, the powdery mildew fungus uses a combination of pressure and a specialized cocktail of cell wall degrading enzymes that function without triggering the plant\'s elaborate defense system (Schulze-Lefert and Vogel, 2000). Currently, powdery mildew growth is contained by expensive year-round chemical applications and breeder-developed varietals. Modern engineering approaches have the potential to decrease the environmental impact of fungicides, and pectin modification is an effective target in multiple agricultural systems (Ferrari etal., 2008; Osorio etal., 2011; Volpi etal., 2011; Lionetti etal., 2014); however, we are limited in our understanding of pectin biosynthesis and how this modification would affect resistance to other pathogens. Of the estimated 65 enzyme activities required for the synthesis of the variety of pectic polysaccharides (Mohnen, 2008; Caffall and Mohnen, 2009; Harholt etal., 2010), very few have yet been characterized. Previously, 20 mutants resistant to the powdery mildews Golovinomyces cichoracearum and Golovinomyces orontii were identified through an ethyl methanesulfonate (EMS)-generated forward genetics screen in Arabidopsis thaliana (Vogel and Somerville, 2000). One of these mutants, powdery mildew resistant5 (pmr5), is mutated in a gene encoding a member of the trichome birefringence-like (TBL) /DUF231 family of proteins that includes several proteins involved in cell wall biosynthesis and modification. Powdery mildew resistance in pmr5 does not appear to trigger any known defense signaling pathways, including salicylic acid, jasmonic acid and ethylene defense pathways, which suggests that pmr5-mediated resistance may act through a previously uncharacterized defense pathway (Vogel etal., 2004). Characterized proteins from the TBL/DUF231 family include a diversity of cell wall modifiers: AXY4 (acetylation of xyloglucan) (Gille etal., 2011b; Zhu etal., 2014), TBR (trichome birefringence), a gene involved in cellulose biosynthesis (Bischoff etal., 2010), ESK1 (eskimo1) involved in the acetylation of xylan (Xin etal., 2007; Yuan etal., 2013; Urbanowicz etal., 2014), TBL3 and TBL31, which are required for the 3-O-monoacetylation of xylan (Yuan etal., 2016), and TBL10, which O-acetylates rhamnogalacturonanI (Stranne etal., 2018). From another protein family, RWAs (reduced wall acetylation) are likely to function as transporters for acetyl-CoA from the cytosol to the Golgi lumen to provide substrate for acetylation (Manabe etal., 2013), and AXY9 is suggested to produce an acetylated intermediate for the acetylation of xyloglucan (Schultink etal., 2015). Of the four RWA proteins in Arabidopsis, RWA2 is particularly important for pectin acetylation (Manabe etal., 2013). RWAs are also highly homologous to the Cas1p (capsule structure1) protein from Cryptococcus neoformans, which was identified in a Cas mutant deficient in acetylation of its coat polysaccharide (Janbon etal., 2001). Interestingly, PMR5 has also been identified as homologous to Cas1p as both have the same esterase domain containing an enzymatic triad within the DUF231 domain (Anantharaman and Aravind, 2010). In the DUF231 family, only PMR5 has been implicated in disease resistance, although the freezing tolerance of esk1 and the drought tolerance of TBL10 are both interesting pleiotropic abiotic stress-related traits in this family. In this study, we demonstrate that PMR5 is a protein involved in the acetylation of pectin capable of adding acetyl groups to galacturonic acid (GalA) oligosaccharides. We show through site-directed mutagenesis that three amino acid residues, previously identified as the DUF231 enzymatic triad, are essential to PMR5 function. Furthermore, we show that putative PMR5 orthologs in Hordeum vulgare (barley), Oryza sativa (rice), Sorghum bicolor and Triticum aestivum (wheat) complement the pmr5 mutant, illustrating potential routes for engineering resistance. Additionally, through a forward-genetics screen we show that mutations in two previously characterized cell wall synthesis genes, TBR and RWA2, suppress pmr5-mediated resistance to powdery mildew, offering more insight into the connection between pectin acetylation and disease resistance. Finally, we offer evidence that the same cell walls shifts that improve resistance to the biotrophic powdery mildew fungus compromise resistance to the necrotrophic fungus Botrytis cinerea, highlighting the importance of considering microbial ecology when engineering resistance. Cell walls in pmr5 have altered pectin, decreased acetate and decreased cellulose in 5-week-old leaves As PMR5 is part of the DUF231 family that contains cell wall-modifying enzymes, we first tested whether there were developmental shifts in cell wall sugar content. We conducted a survey of pmr5 mutant cell wall monosaccharide content at 7days post germination (in seedlings), in 3- and 5-week-old leaves, and in young stems (FigureS1). The GalA content in 3-week-old leaves is consistent with the previously reported uronic acid contents of pmr5 in 18-day-old leaves (Vogel etal., 2004). Notably, the levels of the major backbone sugar of pectins, GalA, were similar across a variety of developmental stages and tissues, with the exception of 5-week-old leaf tissue, where we observed several shifts in pectic sugars: 17% decrease in GalA, 9% decrease in Rha, 27% increase in Ara and 12% decrease in Gal, relative to the wild type (Figures1a and S1). In addition, 5-week-old leaves also showed a 12% decrease in cell wall acetyl ester content (Figure1b) and a 23% decrease in cellulose content (Figure1c). Sequential extraction of the 5-week-old leaf cell walls indicated the greatest difference in Rha, Ara and Gal contents to be primarily in the sodium carbonate fraction, a fraction rich in the pectic polysaccharide rhamnogalacturonanI (RG-I) (FigureS2) (Brummell, 2006). RG-I is composed of a backbone of alternating GalA and Rha, with arabinan and galactan side chains. Notably, the decrease in GalA was much larger than the decrease in Rha, indicating that not only RG-I but also homogalacturonan was decreased in the mutant, in addition to decreased cellulose and acetyl ester content. Figure 1Open in figure viewerPowerPoint The cell wall biochemistry of the pmr5 mutant. (a) Shift in cell wall galacturonic acid (GalA) content, indicative of pectin, as the plant grows. Error bars show SDs (n=3). (b) Decreased total wall-bound acetyl ester content of pmr5 cell walls in 5-week-old leaves. Error bars represent SEs (n=8). (c) Cellulose content of 5-week-old leaves. Error bars represent SEs (n=6). Significantly different from the wild type (WT) by Student\'s t-test: ***P 0.001. PMR5 adds acetyl residues to pectin fragments using [14C]-acetyl-CoA as an acetyl donor Based on phylogenetic analysis and cell wall biochemistry, we hypothesized that PMR5 is a pectin acetyltransferase involved in adding acetyl groups to the GalA residues on the homogalacturonan or RG-I chain. Additionally, OGAs (10–15mer) are known to form calcium bridges and are biologically active in the plant defense response (Ferrari etal., 2013). As OGAs are not commercially available, we used a previously developed method (Ferrari etal., 2007) to degrade PGA into small- (2–10mer) and medium-sized (10–25mer) OGAs, purified them for use as substrates in these activity assays, and confirmed their enriched size range by high-performance anion-exchange chromatography (HPAEC) (FigureS3). We synthesized PMR5 cDNA without the transmembrane domain and purified the protein from Escherichia coli cells to use in in vitro activity assays. Few other proteins appeared present at the final elution step on a GelCode™ Blue stained gel, and the band that was detected in high quantities was at ~80kDa, the size of the predicted MBP-PMR5-Myc-His protein (FigureS4). This band was further confirmed to be MBP-PMR5-Myc-His by mass spectrometry sequencing. To determine the biochemical activity of the purified PMR5 protein, we incubated freshly purified PMR5 protein with short- and medium-length OGAs and [14C]-acetyl-CoA at varied pH levels. We used a method modified from Rennie etal. (2012) to separate the unincorporated [14C]-acetyl-CoA by paper chromatography. Negative controls included both reactions pre-mixed with termination buffer at a range of pH values and reactions lacking the acceptor. The PMR5 protein had similar activity levels with short- and medium-length OGAs at pH5.0–5.5, whereas significant activity at pH levels of up to 7.0 was only observed with medium-length OGAs (Figure2). The optimal pH value for the medium-length OGAs was consistent with expectations for an endomembrane protein (Golgi, pH6.2–6.3, endoplasmic reticulum, ER, pH7–7.5), but we cannot explain the lack of detectable activity with short OGAs at the higher pH values. Determining the optimal substrate was limited by the lack of commercially available OGAs, but we were able to confirm that the PMR5 protein transfers acetic acid esters to the OGAs. Figure 2Open in figure viewerPowerPoint Purified PMR5 protein activity. Purified PMR5 protein expressed without the transmembrane domain in E. coli cells shows acetyltransferase activity with [14C]-acetyl-CoA donor and different lengths of oligogalacturonides as acceptors. SE (n=3). Statistical significance was evaluated with an ANOVA analysis followed by a Tukey\'s post hoc test. Means with the same letter are not significantly different at a P-value of 0.05. Based on structural conservation with fungal Cas1p proteins, three amino acids are the enzymatic triad that is essential for the enzymatic acyltransferase function (Anantharaman and Aravind, 2010). We used site-directed mutagenesis to determine whether these amino acids and the associated enzymatic functions are essential for PMR5 function. To monitor PMR5 expression and localization, we first created a PMR5-GFP fusion with the native PMR5 promoter. The transformation of pmr5 with pPMR5::PMR5-GFP resulted in the complementation of pmr5 morphological defects and the restoration of powdery mildew susceptibility (Figure3a). To evaluate putative PMR5 catalytic triad residues, we used site-directed mutagenesis to introduce alanine substitutions at S142, D379 and H382 of the pPMR5::PMR5-GFP construct. Upon transformation of pmr5 with the site-directed mutant constructs the mutant proteins accumulated to similar levels as the wild-type proteins, although the D379A mutant exhibited a somewhat lower level of GFP signal (FigureS9); however, each of the three site-directed mutations (S142A, D379A and H382A) abolished the complementation by the pPMR5:PMR5-GFP construct in the pmr5 mutant background, both in terms of morphological phenotype and powdery mildew resistance (Figure3a). These observations indicate that the putative catalytic triad amino acids are necessary for PMR5 function. Figure 3Open in figure viewerPowerPoint The PMR5 predicted catalytic triad is essential for PMR5 function and may have functional conservation between species. (a) Whole 2 weeks uninfected plant and whole infected plant (10dpi) show that site-directed amino acid mutants fail to complement pmr5 in terms of plant size, leaf shape, and disease resistance. (b) Complementation of pmr5 mutant with four orthologs produces wild type phenotype of leaf size, shape, and powdery mildew disease susceptibility. As indicated by Anantharaman and Aravind (2010), the catalytic triad is well conserved across many phyla (FigureS5). To determine how well conserved PMR5 is across other plant species, using BLAST we found that putative orthologs from barley, rice, sorghum and Vitis vinfera (grapevine) were highly conserved at the amino acid level. In all of these four species the catalytic triad was conserved (FigureS6). To determine whether PMR5 function is conserved in these other species, we cloned the four putative orthologs from barley, rice, sorghum and wheat into a construct driven by the native PMR5 promoter to determine whether these constructs could complement the pmr5 mutant. All four constructs complemented the mutant and restored the leaf phenotype and size, as well as restoring powdery mildew disease susceptibility similar to wild type plants (Figure3b). This suggests that the function of PMR5 is conserved in these four species and that site-directed mutagenesis of one of the three amino acids in the catalytic triad could effectively abolish PMR5 function in these other species. Collectively, these discoveries open the possibility of using CRISPR-Cas9 engineering to improve powdery mildew resistance into a wide variety of crop species plants by targeted swapping and mutation of the PMR5 gene. To identify additional genes involved in pmr5-mediated powdery mildew resistance, we conducted a genetic suppressor screen of pmr5. Bulked pmr5 seeds were EMS mutagenized and the progeny were screened as described by Vogel and Somerville (2000) for the restoration of susceptibility to powdery mildew. Twenty suppressors of pmr5 were isolated from independent pools of M2 seeds. Of these 20 suppressors, two mapped to another Cas1p protein family member, At3g06550 (RWA2), and one mapped to a DUF231 family member, At5g06700 (TBR1). Characterized mutant lines for rwa2 and tbr1 were crossed with pmr5 to confirm that these genes were responsible for the suppression of pmr5 resistance (Figure4a). Both RWA2 and TBR1 are characterized as genes affecting the acetylation of plant cell wall polysaccharides. One other powdery mildew-resistant mutant, pmr6, was previously characterized as having reduced esterification of cell wall pectin by Fourier-transform infrared (FTIR) spectroscopy (Vogel etal., 2002). To test whether these two cell wall-related suppressors of pmr5 resistance also could suppress pmr6 resistance, tbr1 and rwa2 mutations were crossed into a pmr6 background. We found that pmr6rwa2 and pmr6tbr1 regained the susceptibility to powdery mildew (FigureS8). We then examined the cell wall composition of all four double mutants (pmr5rwa2, pmr5tbr1, pmr6rwa2 and pmr6tbr1) to determine whether there were any changes that could be correlated with the resistance phenotype. Monosaccharide composition was analyzed in 3-week-old leaves, which was a time point that showed few differences between pmr5 and the wild type in our developmental cell wall survey and was the age of the leaf at the time of all powdery mildew inoculations. The double mutants show shifts in the pectic sugars (generally more galacturonic acid, less galactose and less arabinose than the single mutants), although it is difficult to determine whether these effects are additive, as the rwa2 and tbr1 single mutants also have some shifts relative to the wild type, pmr5 and pmr6 plants (TableS1a). Likewise, the cellulose content of these single and double mutants was also decreased, but not in a clearly additive way (TableS1b). This suggests that cellulose content itself is not responsible for resistance in pmr5 and pmr6. The galacturonic acid content of the wild type was closer to pmr5 and pmr6 than to their double mutants, and so by itself cannot explain the resistance phenotype, and the other pectic sugar shifts were even less clear to be consistently present in resistant mutants and absent in susceptible ones. Figure 4Open in figure viewerPowerPoint Powdery mildew disease phenotype in double mutants identified through a suppressor screen. (a) Late infection leaf phenotype of double mutants identified through suppressor screen 10dpi. (b) Early infection phenotype in using the ratio of fungal to plant gDNA 5 dpi in double mutants in the oligogalacturonide defense pathway. Statistical significance was evaluated with an ANOVA analysis followed by a Tukey\'s post hoc test. Means with the same letter are not significantly different at a P-value of 0.001. Error bars are SD (n = 12). To determine whether an OGA pre-treatment could improve resistance to G.cichoracearum, pmr5 and the double mutants pmr5rwa2 and pmr5tbr1 were treated with OGAs of 10–25mer prior to powdery mildew inoculation, as described by Ferrari etal. (2007). The Col-0 wild type had a substantial decrease in the ratio of fungal to plant gDNA, indicating that the OGA pre-treatment does provide temporary protection from G.cichoracearum infection (Figure5b). Additionally, rwa2 shows high susceptibility to powdery mildew and had the greatest decrease in fungal growth from the OGA pre-treatment. The pmr5tbr1 double mutant had slightly less powdery mildew growth with the OGA pre-treatment, and the pmr5rwa2 double mutant did not gain any additional protection from powdery mildew infection with the OGA pre-treatment. Figure 5Open in figure viewerPowerPoint Powdery mildew disease phenotype in single and double mutants connected to oligogalacturonide defense response. (a) Late infection phenotype 10 dpi of powdery mildew, (b) Early infection phenotype with and without an oligogalacturonide (OGA) pre-treatment 24 h before infection (n = 12). The OGA pre-treatment confers resistance to WT but does not enhance resistance in pmr5. Statistical significance was evaluated using an ANOVA analysis followed by a Tukey\'s post hoc test. Means with the same letter are not significantly different at a P value of 0.05. (c) Early infection phenotype quantified using Gc fungal spore counts 5 dpi in double mutants in oligogalacturonide defense pathway (n = 3). Key: Error bars are SD. Significantly different from WT untreated by t test ***P 0.001. Next, we examined two mutants in the OGA defense pathway, pad3, a mutant in the enzyme that catalyzes the final step in camalexin biosynthesis, and wak1, a wall-associated kinase identified as an intermembrane receptor to extracellular OGAs, and we crossed these two mutations into the pmr5 background (Figure5a). Fungal spores were measured at 5days post-inoculation (5dpi) with G.cichoracearum to gain insight into the relative levels of powdery mildew infection (Figure5c). Much like rwa2, pad3 shows very high susceptibility to powdery mildew. Interestingly, when pad3 is crossed into pmr5, the pmr5/pad3 double mutant retains powdery mildew resistance. Similarly, wak1 was highly susceptible to powdery mildew, and this mutation did not suppress the pmr5-mediated resistance. Despite the high susceptibility of the wak1 and pad3 single mutants, defects in this pathway could not abolish pmr5-mediated resistance, indicating that the pathway for which WAK1 and PAD3 detects signals at the cell wall and transmits them through camalexin synthesis is not required for pmr5-mediated resistance. pmr5 and pmr6 are more susceptible to multiple strains of the necrotrophic pathogen B.cinerea, and produce less camalexin upon infection Biotrophic and necrotrophic pathogens have strikingly different modes of virulence that require different plant innate immune system responses (Glazebrook, 2005); however, the cell wall is typically hypothesized to provide a barrier against both pathogen types, with limited evidence for cell wall resistance mechanisms differentiating between pathogens. Additionally, the quantitative resistance of the plant to generalist necrotrophic pathogens is dependent on the pathogen genotype (Corwin etal., 2016a). To examine if pmr5-mediated resistance protects against necrotrophic pathogens, we inoculated pmr5 and pmr6 plants, and suppressors of pmr5-mediated resistance with four different strains of B.cinerea and measured virulence. We also measured camalexin production, a well-established Arabidopsis defense compound against nectrotrophic pathogens. We found that both pmr5 and pmr6 mutants had larger lesion areas and produced less camalexin during infection for all isolates of B.cinerea relative to the wild type (Figure6). In contrast to pmr5 and pmr6, the interaction of B.cinerea with the rwa2 mutant was highly dependent upon the genotype of the pathogen (Manabe etal., 2011). Specifically, we found lesion size in rwa2 to be similar to that in wild type with Apple517 and B05.10 strains, with a non-significant lower lesion size with Supersteak and UKRazz strains (Figure6a). Similarly, we found elevated camalexin levels in a rwa2 background only for the UKRazz isolate (Figure6b). In the pmr5rwa2 double mutant, lesion size and camalexin production proved to be largely additive for each isolate, showing an intermediate phenotype between the hyper-susceptible pmr5 mutant and the hypo-susceptible rwa2 mutant. Interestingly, tbr1 and pmr5tbr1 were more susceptible to all strains of B.cinerea except UKRazz. This may also be explained by the concomitant increase in camalexin content (Figure6b). Taken together, this suggests that although pmr5 and pmr6 are both more resistant to powdery mildew, a biotrophic pathogen, this comes with a trade-off to being more susceptible to B.cinerea, a generalist necrotrophic pathogen. Additionally, the lower resistance to B.cinerea is correlated with a concomitant decrease in the defense compound camalexin. Figure 6Open in figure viewerPowerPoint Lesion area and camalexin content of single and double mutants. (a) pmr5 and pmr6 are more susceptible to multiple strains of Botrytis cinerea. The pmr5rwa2 double mutant has an intermediate phenotype of lesion area. SE (n 5). (b) Camalexin leaf content following B.cinerea inoculation. Nearly all mutants produce less camalexin than the wild type (WT) control. Camalexin production is inversely proportional to lesion area in most genotypes. SE (n 5). Key: significantly different from WT by ANOVA *P 0.05, **P 0.01, ***P 0.001, significantly different among fungal genotypes by ANOVA +P 0.05. We report evidence suggesting that PMR5 is a functional acetyltransferase that mediates pectin acetylation on GalA residues, and this leads to altered pathogen resistance. This is evidenced by the observation that the pmr5 mutant shows decreased acetyl esters extracted from the cell walls in 5-week-old leaves and altered pectic sugar composition, and that the PMR5 protein purified from E.coli shows an increased number of OGA products with a radiolabeled acetyl group. PMR5 belongs to the DUF231 protein family that has other members previously characterized to be involved in cell wall acetylation. Finally, PMR5 has an esterase domain that is evolutionarily conserved with the C.neoformans Cas1p protein (FigureS5) involved in acetylating the fungal coat (Janbon etal., 2001). The acetylation of the fungal coat by Cas1p is essential for the virulence of Cryptococcus in animal hosts. The protein is also homologous with the Cas1 mammalian protein involved in the acetylation of sialic acid in certain glycolipids (Arming etal., 2011). This provides important insight into the previously uncharacterized function of PMR5. The activity on OGAs suggests that PMR5 acetylates homogalacturonan domains of pectin, unlike the recently reported TBL10, which is specific for RG-I domains (Stranne etal., 2018). Neither pmr5 nor tbl10 is devoid of acetylation of the respective pectin domains, suggesting that additional members of the large TBL family are involved in pectin acetylation. To what extent the acetyltransferases are redundant or acetylate specific subdomains of homogalacturonan and RG-I remains to be determined. Since the initial characterization of pmr5 in 2000, the underlying mechanism for powdery mildew resistance in this Arabidopsis mutant has remained a mystery. We hypothesized that pmr5 powdery mildew resistance resulted from an increase in biologically active OGAs being solubilized by the fungus upon fungal cell wall degradation, leading to the activation of the OGA defense pathway (Ferrari etal., 2013). This hypothesis would be consistent with previous findings that pmr5-mediated resistance is not dependent on known resistance pathways (Vogel etal., 2004), as the OGA defense pathway is not dependent on the activation of salicylic acid (SA), jasmonic acid (JA) or ethylene defense pathways (Ferrari etal., 2007). We demonstrated that OGA pre-treatment protected Arabidopsis from G.cichoracearum infection in wild-type plants (Figure5b). One study in wheat found that the wheat powdery mildew (Blumeria graminis f.sp. tritici) had decreased infection with an OGA pre-treatment that was acetylated as compared with the unacetylated OGAs (Randoux etal., 2010). The defense pathway activated by OGAs requires PAD3, the enzyme that catalyzes the final step in camalexin biosynthesis (Ferrari etal., 2013), and also uses WAK1, a wall-associated kinase that is the receptor for biologically active oligosaccharides that activate this pathway (Brutus etal., 2010; Kohorn and Kohorn, 2012); the pmr5pad3 double mutant as well as the pmr5wak1 double mutant do not abolish pmr5-mediated resistance (Figure5a,c), indicating that pmr5-mediated resistance is not dependent on the activation of the OGA defense pathway, disproving our original hypothesis. Nevertheless, given the high susceptibility of both wak1 and rwa2, and that exogenous application of OGAs decreases powdery mildew infection in the wild type (Figure5b), it appears that this pathway already provides some baseline defense against powdery mildew. Although pmr mutants are less susceptible to powdery mildew infection, both pmr5 and pmr6 mutants are more susceptible to B.cinerea. This could be attributed in part to reduced camalexin production following B.cinerea inoculation (Figure6a,b). The pmr5tbr1 double mutant and suppressor of pmr5-mediated resistance and the tbr1 single mutant are also more susceptible to B.cinerea, with a concomitant lower camalexin production upon infection. Necrotrophs have a larger number of cell wall degrading enzymes than biotrophs (Kämper etal., 2006; O\'Connell etal., 2012), and the acetylation of wall polysaccharides affects the association with cellulose and extractability of the cell wall (Selig etal., 2009). It is possible that the relative susceptibility of these mutants to B.cinerea results from altered pectic fragments that activate the camalexin synthesis pathway, but it also possible that the altered cellulose content (Figure6b; Table S1b) makes the cell wall more easily hydrolyzed by the fungus. The B.cinerea fungus may be demolishing the cell walls in the mutant faster, leading to less stress detection (Hamann, 2012) and less camalexin production. This increased stress detection hypothesis would not explain why the rwa2 mutant increases resistance to just a subset of specific B.cinerea strains, however (Manabe etal., 2011) (Figure6a). This shows that there is significant genetic variation among isolates of Botrytis for their virulence in RWA-mediated resistance and PMR5-mediated susceptibility, which could be investigated through genetic mapping in the fungus. Cell wall modifications can lead to defense priming that can alter pathogen penetration and growth (Underwood, 2012). Although rwa2 has similar monosaccharide and cellulose alterations as the other mutants examined here (TableS1a,b), it is clear that the altered acetylation of cell wall polysaccharides leads to improved resistance through improved stress detection with certain B.cinerea strains, although the exact mechanism remains elusive. Indeed, the rwa2 mutant exhibits a massive induction of defense-related genes in the absence of pathogen infection (Nafisi etal., 2015). Pectin acetylation, as demonstrated for pectin methylesterification, could be a pectin integrity maintenance mechanism that the plant has evolved to render pectin less accessible to the action of hydrolases produced by Botrytis strains (Lionetti etal., 2017). We found that the acetate ester content of pmr5 leaves was 12% lower than that of the wild type. This is a relatively minor shift to cause such large disease phenotypes as increasing resistance to powdery mildew and decreasing resistance to B.cinerea. This illustrates how even slight modifications to cell wall structure lead to trade-offs in fungal disease resistance and highlight the importance of considering microbial ecology when engineering resistance. It also suggests that some plant cell wall composition formulations may have evolved for defense against particular pathogens in environments with that selective pressure, and some may have evolved for an average level of resistance to commonly found pathogens. We show that PMR5 can add acetyl residues to OGAs, and that an exogenously applied OGA treatment to wild-type leaves decreased powdery mildew infection. We show that pmr5 is more susceptible to Botrytis infection and produces less camalexin upon infection, providing more evidence linking the OGA defense pathway to camalexin production; however, two genes essential to this pathway, wak1 and pad3 (the final biosynthetic enzyme for camalexin production), when crossed with pmr5 did not abolish the pmr5-mediated resistance, indicating that the camalexin response is not solely responsible for the powdery mildew resistance phenotype. A suppressor screen for the return to susceptibility mapped two of these suppressors to cell wall genes, RWA2 (reduced wall acetylation2) and TBR (trichome birefringence), suggesting that pmr5-mediated resistance is connected to cell wall structure and/or degradation upon pathogen attack. We found that the three predicted amino acid residues in the catalytic triad are essential for complementing the pmr5 mutant, and that these amino acids are highly conserved in other species. Putative PMR5 homologs from barley, grape, rice and sorghum complemented the pmr5 mutant. The latter two results suggest a future route for engineering resistance. Arabidopsis thaliana Heynh. (L) accession Columbia-0 (Col-0) was used for the control line in this study. Arabidopsis seeds were sown on soil (Pro-Mix HP; PRO-MIX, https://www.pthorticulture.com) and stratified for 3days at 4°C. Plants to be infected with powdery mildew (G.cichoracearum, race UCSC1) were grown in growth chambers at 22°C with 70% relative humidity (RH) and a 14-h photoperiod. The Arabidopsis mutants used in this study include: pmr5 (ABRC stock: CS6579), pmr6-1 (ABRC stock: CS6354), tbr-1 (ABRC stock: CS3741), rwa2-3 (SALK_013562), wak1 (SALK_107175) and pad3 (SALK_026585C). Homozygous plants were confirmed by polymerase chain reaction (PCR) with gene-specific primers (TableS2). Expression levels of PMR5 were quantified using qPCR in 3-week-old leaves of the wild type and the pmr5 mutant, and showed that PMR5 was expressed ~2.7-fold lower in pmr5 than in the wild type (FigureS7). Arabidopsis plants were transformed using Agrobacterium tumefaciens GV3101 via the floral-dip method (Clough and Bent, 1998). For selection, seeds were germinated on half-strength MS media with 25μgml−1 kanamycin. Resistant plants were transferred to soil and grown as described above. Construction of barley, grape, rice and sorghum PMR5 ortholog complementation constructs Sequences for the cDNA for H.vulgare, O.sativa and V.vinifera were synthesized by Life Technologies (ThermoFisher Scientific, https://www.thermofisher.com), as found in the NCBI database (https://www.ncbi.nlm.nih.gov). The PMR5 gene from Sorghum bicolor was obtained by amplification from cDNA extracted from young leaves. Products were sequenced to confirm their identity. Constructs were made using the Gibson assembly technique, with one fragment consisting of the AtPMR5 upstream promoter region (1000bp), including the 5′ untranslated region (5′-UTR) and the second fragment consisting of the PMR5 homolog from each organism, with and without a stop codon. These fragments were introduced into pMDC99 to produce an untagged PMR5 as well as into pMDC107 to produce the GFP-tagged PMR5. Both plasmids were digested with PacI and AscI to produce the vector backbone for assembly. Assembly was performed using a Gibson assembly cloning kit from New England Biolabs (https://www.neb.com). Genomic DNA from Arabidopsis ecotype Col-0 was isolated using a previously described CTAB DNA prep (Lukowitz etal., 1996). To generate the pPMR5:PMR5-GFP construct, the genomic region containing the PMR5 coding sequence and the promoter 1kb upstream of the coding sequence was amplified from Arabidopsis Col-0 genomic DNA by PCR using the primers listed in TableS2. The amplified product was electrophoresed on a 1% agarose gel, cut from the gel using a razor blade, and purified from the gel slice using a Zymoclean Gel DNA Recovery kit (Zymo Research, https://www.zymoresearch.com), according to the manufacturer\'s protocol. The PMR5 product was cloned into a Gateway entry vector using a pCR8/GW/TOPO cloning kit (Life Technologies, ThermoFisher Scientific) according to the manufacturer\'s protocol. The pPMR5:PMR5 fragment was subsequently transferred to the vector pMDC107 (Curtis and Grossniklaus, 2003) to fuse GFP in-frame to the C terminus of PMR5 using LR Clonase (Life Technologies, ThermoFisher Scientific), according to the manufacturer\'s instructions. Putative esterase catalytic triad mutants PMR5S142A-GFP, PMR5D379A-GFP and PMR5H382A-GFP were generated by site-directed mutagenesis of the pPMR5:PMR5-GFP construct described above using a Quikchange II XL site-directed mutagenesis kit (Agilent Technologies, https://www.agilent.com) with mutagenesis primers, according to the manufacturer\'s protocol (TableS2). For Arabidopsis transformation, all vector constructs were transformed into Agrobacterium strain GV3101 (Koncz and Schell, 1986) by electroporation. The cDNA for SbPMR5 was amplified by PCR using gene-specific primers (TableS2) from first-strand cDNA made from pooled rice samples. Coding sequences for genes were cloned using Gateway technology (Invitrogen, ThermoFisher Scientific) and Gateway-compatible primers (TableS2), as follows: PCR reaction products were gel-purified using the MinElute Gel extraction kit (Qiagen, https://www.qiagen.com) and used for Gibson assembly reactions using the LR clonase enzyme mix (Invitrogen, ThermoFisher Scientific). Plant leaf samples were observed on a confocal microscope system consisting of a Leica DMI 6000B inverted microscope (Leica Microsystems, https://www.leica-microsystems.com) fitted with a Yokogawa CSU-X1 spinning disk confocal head (Yokogawa Electric Corporation, https://www.yokogawa.com) and a Photometrics QuantEM 512SC EM-CCD camera (Photometrics, https://www.photometrics.com). Microscope control and image acquisition were accomplished using metamorph (
Molecular Devices, https://www.
moleculardevices.com). Pieces were cut from 3-week-old Arabidopsis leaves and mounted in H2O on microscope slides for imaging. Leaf samples were observed using a 63× water immersion objective. GFP was excited with a 488-nm diode laser and observed using a 525±25-nm emission filter. Image processing was performed using imagej (rsbweb.nih.gov/ij/) and photoshop (Adobe, https://www.adobe.com). z-projections were prepared from 50–150 optical sections (z-distance 0.3μm) by maximum projection using imagej. We performed several experiments to test the putative PMR5 pectin acetyltransferase activity. In our initial experiments, we infiltrated Nicotiana tabacum (Nicotiana benthamiana) leaves with a vector containing 35S:PMR5-GFP, confirming expression by microscopy, and isolating the microsomal fraction for an activity assay. We used [14C]-acetyl-CoA as the donor and endogenous substrates as acceptors; however, we were not able to detect a difference in acetyltransferase activity in the PMR5-infiltrated lines above the relatively high level of endogenous acetyltransferase activity. As a result of this high background acetyltransferase activity, we next established a two-step protein affinity purification system in E.coli. The PMR5 coding sequence was synthesized without the first 29 amino acids (starting at nucleotide 88) with a Myc and His tag at the C-terminal end. Transmembrane domain predictions for PMR5 are from amino acids 6–21, so this effectively deleted the transmembrane domain, making the protein soluble. This was cloned into the pMAL-c2 vector for the pMAL protein fusion and E.coli protein purification system, and transformed into OrigamiB E.coli cells (Novagen, now Merck, http://www.merckmillipore.com). A 6-ml starter culture with rich media+glucose (1L:10g tryptone, 5g yeast extract, 5g NaCl, 2g glucose, 100μgml−1 ampicillin) was grown overnight at 37°C, then added to 3L of rich media+glucose and grown at 37°C to an OD600 of 0.5, induced with 0.3m isopropyl β-D-1-thiogalactopyranoside (IPTG), and grown at 16°C for 16h before pelleting cells and resuspending them in lysis buffer (50mm NaH2PO4, pH7.6, 150mm NaCl, 5% glycerol, 5mm imidazole, 1mm 2-mercaptoethanol, 1 EDTA-free Protease Inhibitor Cocktail Tablet; Roche, https://www.roche.com). Cells were lysed using an Avestin E3 Emulsiflex homogenizer and cell debris was pelleted (JA-20 rotor, 48 400 g, 1.5h, 4°C). Lysate was run through a 10-ml equilibrated column with Ni-NTA resin, washed once with lysis buffer and then eluted with 200mm imidazole. Flow through was added to an equilibrated amylose resin column, which was washed with a column length of lysis buffer and eluted with lysis buffer+100mm maltose. The 80-kDa bands generated from MBP-PMR5-His purification were cut out from a sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gel and digested following the UC Berkeley QB3 proteomics/mass spectrometry protocol (http://qb3.berkeley.edu/pmsl/wp-content/uploads/2016/06/In-gel-digestion-protocol.pdf). Briefly, gel fragments were washed with a 100mm ammonium bicarbonate wash, reduced with 45mm dithiothreitol (DTT), incubated with 100mm iodoacetamide for irreversible alkylation, then washed with a 50:50 mix of acetonitrile and ammonium bicarbonate. Gel pieces were dehydrated with acetonitrile and then dried by vacuum centrifuge. Gel pieces were rehydrated with 25mm ammonium bicarbonate containing modified trypsin (Promega, https://www.promega.com) to digest overnight at room temperature (20–24°C). Peptides were recovered in the supernatant from the overnight digest and then further extracted with 60% acetonitrile/0.1% formic acid and 100% acetonitrile. Samples were analyzed by one-dimensional LC/MS-MS in the UC Berkeley QB3 proteomics/mass spectrometry lab. Acetyltransferase activity assays were based on the protocols described by Lee etal. (2007) and Rennie etal. (2012). Protein was purified as described above, with an added step of concentrating the final protein to 500μl in a 10000 molecular weight cut-off (MWCO) Vivaspin concentrator (Viva Products, https://www.vivaproducts.com). Purified protein was quantified on a NanoDrop (NanoDrop 8000 Spectrophotometer, ThermoFisher Scientific), and per reaction, 2μg protein was added to buffer containing: 50mm buffer [2-(N-morpholino)ethanesulfonic acid (MES) buffer for pH5.0, 5.5, 6.0 and 6.5 and HEPES buffer for pH7.0, 7.5 and 8.0], 400mm sucrose, 5mm MnCl2, 4μg oligogalacturonides (10–25mer), 2μl (20nCiμl−1) [14C]-acetyl-CoA (1480Bq per reaction, specific activity, 40–60mCimmol−1; Perkin Elmer, https://www.perkinelmer.com) in a total reaction volume of 50μl. After incubation at 24°C for 2h, the reaction was stopped by adding 5μl of termination buffer [0.3m acetic acid containing 20mm ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA)]. The solution was spotted onto Whatman 3MM chromatography paper (Whatman, now GE Healthcare Life Sciences, https://www.gelifesciences.com), which was then developed for 4h in 95% EtOH/1m ammonium acetate, 2:1 (v/v) as the solvent. The radiolabeled oligosaccharides retained at the original spot were cut out and vortexed in 1ml 100mm NaOH, then 200μl 1m acetic acid to neutralize the NaOH, and then 4ml of scintillation fluid was added (Ecoscint XR; National Diagnostics, https://www.nationaldiagnostics.com). The level of activity was determined using a scintillation counter set to measure 14C counts for 2min (Beckman LS 6500; GMI Inc, https://www.gmi-inc.com/). Negative protein controls were heat treated at 100°C for 30min prior to the addition to reactions. Reactions were set up the same day as protein purification from harvested E.coli cells. The OGAs with a degree of polymerization of 10–25 were prepared by partial hydrolysis of polygalacturonic acid (PGA), based on a protocol described by Bellincampi etal. (2000). In a 1-L bottle, 0.4g PGA (P3889; Sigma-Aldrich, https://www.sigmaaldrich.com) was added to 400ml of water and brought to a pH of 4.4 using 1N NaOH. The solution was autoclaved for 45min, allowed to cool, and brought to a pH of 2 with 1N HCl, stirring constantly, to precipitate the higher dp OGAs. The solution was then centrifuged at 12000g for 20min, 363ml of supernatant was transferred to a 500-ml bottle, and lower dp OGAs (10–25mers) were precipitated by adding 25ml 1m NaOAc and 112ml 100% ETOH. This solution (pH6) was placed at 4°C overnight and centrifuged for 30min at 17400g, the supernatant was poured off, and the pellets were resuspended in 50ml of water, which was dialyzed against 4L of water for 2days, changing the water twice. This solution was lyophilized and stored as powder in −20°C. The enrichment of the size ranges of the OGAs in solution (1mgml−1) was confirmed by HPAEC on a Dionex ICS3000 CarboPac PA200 system (ThermoFisher Scientific) equipped with a pulsed amperometric detector (PAD), based on Hotchkiss and Hicks (1990) (FigureS3). For Arabidopsis plants, 3- and 5-week-old leaf tissue as well as 6-week-old stem tissue was collected, frozen in liquid nitrogen and freeze-dried overnight. Alcohol insoluble residue (AIR) preparation and de-starching was performed according to the methods described by Yin etal. (2011). For monosaccharide composition analysis, 5mg was hydrolyzed in 2m trifluoroacetic acid (TFA) at 120°C for 1h. The released monosaccharides were separated by HPAEC on a Dionex ICS3000 system equipped with a PAD as described by Harholt etal. (2006). For cellulose content analysis, 5mg of AIR was treated with 72% H2SO4 for 1h at 30°C with shaking and diluted with 715μl water and autoclaved at 120°C for 1h. Samples were diluted 200 times with water and run on the HPAEC-PAD system for quantification. All samples were analyzed on a CarboPac PA-20 column (ThermoFisher Scientific) eluted with 2mm KOH in order to resolve and quantify Fuc, Gal, Glc, Xyl and Man. Samples without concentrated sulfuric acid pre-treatment were further analyzed on a CarboPac PA-20 column eluted with 18mm of KOH in order to resolve and quantify Rha and Ara. For acetic acid quantification, 10mgml−1 AIR was saponified by adding an equal volume of 1m NaOH and incubated for 1h at 26°C with shaking (600rpm). The de-esterified samples were neutralized with an equal volume of 1m HCL, pelleted for 10min at 20800g, and the acetic acid content of the supernatant was determined using the Acetic Acid Kit (K-Acet; Megazyme, https://www.megazyme.com), as described by Gille etal. (2011a). De-starched AIR (5mg) was suspended in 0.5ml 0.05m CDTA (pH6.5) for 24h at room temperature on a thermomixer. The suspension was centrifuged at 48000g at 4°C and the pellet was washed twice with deionized water. The pellet was subsequently sequentially extracted using 0.05m Na2CO3 containing 0.01m NaBH4 for 24h at 4°C and washed twice with deionized water. The CDTA and Na2CO3 extracts represent the pectin-rich fractions. For the hemicellulose-rich fraction, the residual pellet was extracted with 4m KOH for 24h at room temperature with shaking on a thermomixer. This was centrifuged at 48000g at 4°C to collect the supernatant, which was adjusted to pH5 with glacial acetic acid and dialyzed against deionized water and lyophilized. Each fraction was then TFA hydrolyzed and run on the HPAEC-PAD system for monosaccharide quantification as described above. Squash, variety Kuta (Park Seed, https://parkseed.com), was used as a host for the production of G.cichoracearum UCSC1 inoculum. The inoculum was prepared by touching squash plants with infected squash leaves 10–12days before inoculation. Arabidopsis plants were inoculated at 18–21days post-germination by placing a 1.3-m settling tower over two flats and tapping between one and three squash leaves over the top of the settling tower. After allowing the spores to settle for 5min, plants were placed in a high-humidity growth chamber (Adam and Somerville, 1996). For the OGA pre-treatment of Arabidopsis seedlings, the protocol was based on that described by Ferrari etal. (2007). Seedlings (18day old) grown as described above were treated with OGAs at 24h and 4h prior to inoculation with powdery mildew. Plants were sprayed with 200μgml−1 OGA solution, leaving the lid on the plant tray for 3h after each treatment. Ten days after inoculation, a quantitative disease resistance (QDR) score for each plant was measured as described by Meyer etal. (2005) and Wang etal. (2018), with wild-type plants included in each inoculation batch as a comparison: 0=no disease phenotype; 1=few visible conidia; 2=visible conidia but less than wild type; 3=conidia equivalent to wild type; and 4=more conidia than wild type. For lines that consistently scored 1, 2 or 4, additional more quantitative assays were completed, including spore counts and qPCR of fungal to plant gDNA. The protocol for quantifying fungal spores on each genotype was adopted from Weßling and Panstruga (2012). Leaves infected with G.cichoracearum (6dpi) were harvested, taking 500mg of leaves per genotype. Water was added (15ml) and vortexed for 30s. The spore solution was filtered through Miracloth (Merck) to remove large debris and leaves. The resulting spore solution was pelleted (5min, 4000g), removing the supernatant, and resuspended in 50μl 10% glycerol. Spores were counted in 10 1-mm2 fields of a Neubauer-improved haemocytometer (Marienfeld, https://www.marienfeld-superior.com) and the results were averaged over at least three slides, with at least three biological replicates per genotype. Spore counts were normalized to the initial weight of the leaves collected. The protocol for quantifying fungal to plant genomic DNA was as described by Weßling and Panstruga (2012). Leaves infected with G.cichoracearum (5dpi) from approximately 40 plants per biological replicate per genotype were harvested and flash-frozen in liquid nitrogen. Genomic DNA was isolated as previously described (Brouwer etal., 2003). Each sample was ground by adding a 1-mm metal ball and the frozen material was agitated in a MM400 mixer mill (Retsch, https://www.retsch.com) for 1min at 30Hz. After grinding, 300μl of lysis buffer [2.5m LiCl, 50mm Tris-HCl, 62.5mm Na2-ethylenediamine tetraacetic acid (EDTA), and 4.0% Triton X-100, pH8.0] and 300μl phenol:chloroform:isoamyl alcohol (25:24:1 v/v) were added and the samples vortexed. Samples were centrifuged (5min, 16000g) and the supernatant was precipitated by two volumes of 100% ethanol, pelleted by centrifugation again, washed in 70% ethanol, air-dried and then resuspended in 50μl of water. DNA quality and concentration were assessed on a NanoDrop (NanoDro 8000 Spectrophotometer, ThermoFisher Scientific). For Q-PCR, 15-μl samples were prepared using the Brilliant Sybr Green QPCR Reagent Kit (Stratagene, https://go.strategene.org), according to the manufacturer\'s instructions. Three technical replicates per sample and a primer concentration of 0.4μm were used. The qPCR program was: denaturation at 95°C for 3min, 40 repeats of 95°C for 20s, 61°C for 20s and 72°C for 15s. A melting curve analysis was completed from 55°C to 95°C in 0.5°C steps with a 10-s dwell time. The ratio of G.cichoracearum to Arabidopsis genomic DNA was calculated using the ΔΔCt method (Pfaffl, 2001). Primers for Q-PCR are listed in TableS2. Total RNA was extracted using the RNeasy plant mini kit (Qiagen), following the manufacturer\'s instructions. RNA preparations were treated with DNase1 (Qiagen) to remove traces of DNA contamination. RNA (1μg) was used for reverse transcription with the Transcriptor high-fidelity cDNA synthesis kit (Roche) and oligo dT primers. After synthesis, the cDNA reaction was diluted four times in RNAse-free water, and 2μl was used for RT-PCR using gene-specific primers (TableS2). Molecular phylogenetic analysis of the PC-esterase domain by maximum-likelihood method The evolutionary history was inferred by using the maximum-likelihood method using the JTT matrix-based model (Jones etal., 1992). The tree with the highest log likelihood (-2467.1125) is shown (FigureS5). The initial tree(s) for the heuristic search was obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using a JTT model, and then selecting the topology with superior log-likelihood value. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site (next to the branches). The analysis involved 28 amino acid sequences. All positions containing gaps and missing data were eliminated. There were a total of 39 positions in the final data set. Evolutionary analyses were conducted in mega7 (Kumar etal., 2016). Assays for Botrytis susceptibility were completed as previously described (Denby etal., 2004; Kliebenstein etal., 2005). In brief, a total of 20 plants per mutant and a wild-type control (Col-0) were grown in a randomized complete block design for 5weeks. Plants were sown on soil (Sunshine Mix #1; Sun Gro Horticulture, http://www.sungro.com) under full-spectrum lights using short-day conditions (10h light). At 5weeks of age, the first true, mature leaves of each plant were collected and placed on growing flats containing 1% Phytagar. Detached leaves were infected using 4μl of a half-strength organic grape juice control or a spore suspension containing 10sporesperμl of one of four genotypically diverse isolates (Apple517, B05.10, Supersteak or UKRazz). Infected leaves were incubated for 72h at room temperature under constant light. Experiments were fully replicated using different growing chambers with 2weeks between plantings. Pictures of infected leaves were taken at 72hpi with a 1-cm reference scale on the flat and leaves were placed in 400μl of 90% methanol for camalexin extraction. To measure the lesion area, pictures were analyzed using the r package ebimage to isolate the lesion from the leaf and to calculate the lesion area. Camalexin content was determined as previously described (Corwin etal., 2016a,b). In brief, leaves were ground in methanol, centrifuged to remove precipitate, and 300μl of supernatant was transferred to a fresh micro-titer plate. A total of 50μl of extract was sampled and run on an Agilent 1100 series HPLC equipped with a Agilent Lichrocart 250-4 RP18e 5μm column (Agilent, https://www.agilent.com). The separation of camalexin was accomplished using an acetonitrile and water gradient according to the following program: 5-min gradient from 63% to 69% acetonitrile, 30-s gradient from 69% to 99% acetonitrile, 2min at 99% acetonitrile and a post-run equilibration of 3.5min at 63% acetonitrile. The detection of camalexin was achieved with an Agilent fluorescence detector (FLD) with excitation of 318nm and emission detection at 385nm. A standard curve using purified camalexin was used to identify and quantitate camalexin from in vivo samples. A Student\'s t-test was used to determine the significance of differences found in cell wall biochemistry, as measured in the activity assays, with the levels of significance indicated as: *P 0.05, **P 0.01 and ***P 0.001. For differences in spore counts and quantitative gDNA qPCR measurements, the significance was determined with an analysis of variance (ANOVA) test followed by a Tukey\'s post-hoc test. For Botrytis test results, significance testing for the effect of each cell wall mutant was determined separately for lesion area (TableS3) and camalexin content (TableS4) by including all individual alleles in a single general linear model (GLM). F-tests were derived from these linear models using the following expression: where Y is the dependent phenotype (lesion area or camalexin). The main fixed effects are denoted as E, I, G, R, P, S and T, which represent the experimental block, infection status (i.e. uninfected control vs infected), fungal genotype, RWA2-3 allele, PMR5 allele, PMR6 allele and TBR1-1 allele, respectively, with e=1,2; i=1,2; g=1…4; r=1,2; p=1,2; s=1,2; and t=1,2. The fungal genotype was nested within the infection status in order to avoid overestimating the significance of the treatment. All P values were derived from a type-II sum of squares and the residual error was assumed to follow a multivariate normal distribution [~N(0,2)]. We thank Gustavo Garcia and Jasmin Chau Tran for technical assistance. We thank Clarice de Azevedo Souza, Nadav Sorek and Trevor Yeats for helpful discussions. We thank Bradley Dotson for pmr5tbr1 seed stock. This work was supported by the Energy Biosciences Institute and NSF IOS 1339125 to DK. This work was part of the DOE Joint BioEnergy Institute (http://www.jbei.org) and the DOE Joint Genome Institute (http://www.jgi.org) supported by the US Department of Energy, Office of Science, Office of Biological and Environmental Research, through contract DE-AC02-05CH11231 between Lawrence Berkeley National Laboratory and the US Department of Energy. SS, DK and HS supervised this study. DC, WU, JC, AR, HS, CCC and DB collected the data, and DC and JC performed the data analysis. JV mutagenized pmr5 and screened for disease suppressor mutants. DC wrote the article with input from the other authors. Figure S1. Cell wall characterization of the pmr5 mutant in multiple tissues and time points. Figure S2. Sequentially extracted cell walls of 5-week-old leaves in the pmr5 mutant. Figure S3. Chromatography profile of OGAs solution. Figure S4. PMR5 purification from E.coli. Figure S5. Molecular phylogenetic analysis of previously identified PC-esterase domain members by maximum-likelihood method. Figure S6. Conservation of catalytic triad in other species. Figure S7. Expression of PMR5 in 3-week-old leaves of wild type and pmr5. Figure S8. Powdery mildew disease phenotype in single and double pmr6 mutants. Figure S9. Microscopy of the PMR5-GFP protein in three site-directed mutants in the pmr5 mutant background. Table S1. Cell wall characterization of two cell wall mutant suppressors of powdery mildew disease resistance. Table S2. List of primers used for genotyping, gene cloning, and quantitative PCR. Table S3. ANOVA table (type-II tests) for lesion area of pmr5 and two suppressors of powdery mildew resistance. Table S4. ANOVA table (type-II tests) for camalexin content of pmr5 and two suppressors of powdery mildew resistance. Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article. Adam, L. and Somerville, S.C. (1996) Genetic characterization of five powdery mildew disease resistance loci in Arabidopsis thaliana. Plant J. 9(3), 341– 356. Wiley Online Library Anantharaman, V. and Aravind, L. (2010) Novel eukaryotic enzymes modifying cell-surface biopolymers. Biol. Direct, 5, 1. Arming, S., Wipfler, D., Mayr, J., Merling, A., Vilas, U., Schauer, R., Schwartz-Albiez, R. and Vlasak, R. (2011) The human Cas1 protein: a Sialic Acid-Specific O-Acetyltransferase? Glycobiology, 21(5), 553– 564. Bellincampi, D., Cervone, F. and Lionetti, V. (2014) Plant cell wall dynamics and wall-related susceptibility in plant–pathogen interactions. Front. Plant Sci. 5, 228. https://doi.org/10.3389/fpls.2014.00228. Bellincampi, D., Dipierro, N., Salvi, G., Cervone, F. and De Lorenzo, G. (2000) Extracellular H2O2 induced by oligogalacturonides is not involved in the inhibition of the auxin-regulated rolB gene expression in tobacco leaf explants. Plant Physiol. 122(4), 1379– 1386. Bischoff, V., Nita, S., Neumetzler, L., Schindelasch, D., Urbain, A., Eshed, R., Persson, S., Delmer, D. and Scheible, W.-R. (2010) TRICHOME BIREFRINGENCE and its homolog AT5G01360 encode novel plant-specific DUF231 proteins required for cellulose biosynthesis in Arabidopsis. Plant Physiol. 153(2), 590– 602. Brouwer, M., Lievens, B., Van Hemelrijck, W., Van den Ackerveken, G., Cammue, B.P. and Thomma, B.P. (2003) Quantification of disease progression of several microbial pathogens on Arabidopsis thaliana using real-time fluorescence PCR. FEMS Microbiol. Lett. 228(2), 241– 248. Wiley Online Library Brummell, D.A. (2006) Cell wall disassembly in ripening fruit. Funct. Plant Biol. 33(2), 103– 119. Brutus, A., Sicilia, F., Macone, A., Cervone, F. and De Lorenzo, G. (2010) A domain swap approach reveals a role of the plant wall-associated Kinase 1 (WAK1) as a receptor of oligogalacturonides. Proc. Natl Acad. Sci. USA, 107(20), 9452– 9457. Caffall, K.H. and Mohnen, D. (2009) The structure, function, and biosynthesis of plant cell wall pectic polysaccharides. Carbohyd. Res. 344(14), 1879– 1900. Clough, S.J. and Bent, A.F. (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16(6), 735– 743. Wiley Online Library Corwin, J.A., Copeland, D., Feusier, J., Subedy, A., Eshbaugh, R., Palmer, C., Maloof, J. and Kliebenstein, D.J. (2016a) The quantitative basis of the Arabidopsis innate immune system to endemic pathogens depends on pathogen genetics. PLoS Genet. 12(2), e1005789. Corwin, J.A., Subedy, A., Eshbaugh, R. and Kliebenstein, D.J. (2016b) Expansive phenotypic landscape of botrytis cinerea shows differential contribution of genetic diversity and plasticity. Mol. Plant Microbe Interact. 29(4), 287– 298. Cosgrove, D.J. (2005) Growth of the plant cell wall. Nat. Rev. Mol. Cell Biol. 6(11), 850– 861. Curtis, M.D. and Grossniklaus, U. (2003) A gateway cloning vector set for high-throughput functional analysis of genes in planta. Plant Physiol. 133(2), 462– 469. Denby, K.J., Kumar, P. and Kliebenstein, D.J. (2004) Identification of Botrytis Cinerea susceptibility loci in Arabidopsis Thaliana. Plant J. 38(3), 473– 486. Wiley Online Library Ferrari, S., Galletti, R., Denoux, C., De Lorenzo, G., Ausubel, F.M. and Dewdney, J. (2007) Resistance to Botrytis Cinerea induced in Arabidopsis by elicitors is independent of salicylic acid, ethylene, or jasmonate signaling but requires PHYTOALEXIN DEFICIENT3. Plant Physiol. 144(1), 367– 379. Ferrari, S., Galletti, R., Pontiggia, D., Manfredini, C., Lionetti, V., Bellincampi, D., Cervone, F. and De Lorenzo, G. (2008) Transgenic expression of a fungal endo-polygalacturonase increases plant resistance to pathogens and reduces auxin sensitivity. Plant Physiol. 146(2), 669– 681. Ferrari, S., Savatin, D.V., Sicilia, F., Gramegna, G., Cervone, F. and De Lorenzo, G. (2013) Oligogalacturonides: plant damage-associated molecular patterns and regulators of growth and development. Front. Plant Sci. 4, 49. https://doi.org/10.3389/fpls.2013.00049. Gille, S., Cheng, K., Skinner, M.E., Liepman, A.H., Wilkerson, C.G. and Pauly, M. (2011a) Deep sequencing of Voodoo Lily (Amorphophallus Konjac): an approach to identify relevant genes involved in the synthesis of the Hemicellulose Glucomannan. Planta, 234(3), 515– 526. Gille, S., de Souza, A., Xiong, G., Benz, M., Cheng, K., Schultink, A., Reca, I.-B. and Pauly, M. (2011b) O-Acetylation of Arabidopsis Hemicellulose Xyloglucan requires AXY4 or AXY4L, proteins with a TBL and DUF231 domain. Plant Cell, 23(11), 4041– 4053. Glawe, D.A. (2008) The powdery mildews: a review of the World\'s most familiar (yet Poorly Known) plant pathogens. Annu. Rev. Phytopathol. 46, 27– 51. Glazebrook, J. (2005) Contrasting mechanisms of defense against biotrophic and necrotrophic pathogens. Annu. Rev. Phytopathol. 43, 205– 227. Hamann, T. (2012) Plant cell wall integrity maintenance as an essential component of biotic stress response mechanisms. Front. Plant Sci. 3, 77. Harholt, J., Jensen, J.K., Sørensen, S.O., Orfila, C., Pauly, M. and Scheller, H.V. (2006) ARABINAN DEFICIENT 1 Is a putative arabinosyltransferase involved in biosynthesis of pectic arabinan in Arabidopsis. Plant Physiol. 140(1), 49– 58. Harholt, J., Suttangkakul, A. and Scheller, H.V. (2010) Biosynthesis of pectin. Plant Physiol. 153(2), 384– 395. Hotchkiss, A.T. Jr and Hicks, K.B. (1990) Analysis of oligogalacturonic acids with 50 or fewer residues by high-performance anion-exchange chromatography and pulsed amperometric detection. Anal. Biochem. 184(2), 200– 206. Janbon, G., Himmelreich, U., Moyrand, F., Improvisi, L. and Dromer, F. (2001) Cas1p is a membrane protein necessary for the O-Acetylation of the Cryptococcus Neoformans Capsular Polysaccharide. Mol. Microbiol. 42(2), 453– 467. Wiley Online Library Jones, D.T., Taylor, W.R. and Thornton, J.M. (1992) The rapid generation of mutation data matrices from protein sequences. Comput. Appl. Biosci. 8(3), 275– 282. Kämper, J., Kahmann, R., Bölker, M. etal. (2006) Insights from the genome of the biotrophic fungal plant pathogen Ustilago Maydis. Nature, 444(7115), 97– 101. Kliebenstein, D.J., Rowe, H.C. and Denby, K.J. (2005) Secondary metabolites influence Arabidopsis/Botrytis interactions: variation in host production and pathogen sensitivity. Plant J. 44(1), 25– 36. Wiley Online Library Kohorn, B.D. and Kohorn, S.L. (2012) The cell wall-associated Kinases, WAKs, as pectin receptors. Front. Plant Sci. 3, 88. https://doi.org/10.3389/fpls.2012.00088. Koncz, C. and Schell, J. (1986) The promoter of TL-DNA gene 5controls the tissue-specific expression of chimaeric genes carried by a novel type of Agrobacterium binary vector. Mol. Gen. Genet. 204, 383. https://doi.org/10.1007/BF00331014. Kumar, S., Stecher, G. and Tamura, K. (2016) MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol. Biol. Evol. 33(7), 1870– 1874. Lee, C., O\'Neill, M.A., Tsumuraya, Y., Darvill, A.G. and Ye, Z.-H. (2007) The irregular xylem9 mutant is deficient in xylan xylosyltransferase activity. Plant and Cell Physiol. 48(11), 1624– 1634. https://doi.org/10.1093/pcp/pcm135. Liners, F., Thibault, J.-F. and Van Cutsem, P. (1992) Influence of the degree of polymerization of Oligogalacturonates and of Esterification pattern of pectin on their recognition by monoclonal antibodies. Plant Physiol. 99(3), 1099– 1104. Lionetti, V., Cervone, F. and Bellincampi, D. (2012) Methyl esterification of pectin plays a role during plant-pathogen interactions and affects plant resistance to diseases. J. Plant Physiol. 169(16), 1623– 1630. Lionetti, V., Raiola, A., Cervone, F. and Bellincampi, D. (2014) Transgenic expression of pectin methylesterase inhibitors limits tobamovirus spread in tobacco and Arabidopsis. Mol. Plant Pathol. 15(3), 265– 274. Wiley Online Library Lionetti, V., Fabri, E., De Caroli, M., Hansen, A.R., Willats, W.G.T., Piro, G. and Bellincampi, D. (2017) Three pectin methylesterase inhibitors protect cell wall integrity for Arabidopsis immunity to Botrytis. Plant Physiol. 173(3), 1844– 1863. Lukowitz, W., Meyer, U. and Jürgens, G. (1996) Cytokinesis in the Arabidopsis embryo involves the syntaxin-related KNOLLE gene product. Cell, 84, 61– 71. Manabe, Y., Nafisi, M., Verhertbruggen, Y. etal. (2011) Loss-of-function mutation of REDUCED WALL ACETYLATION2 in Arabidopsis leads to reduced cell wall acetylation and increased resistance to Botrytis Cinerea. Plant Physiol. 155(3), 1068– 1078. Manabe, Y., Verhertbruggen, Y., Gille, S. etal. (2013) RWA proteins play vital and distinct roles in cell wall O-Acetylation in Arabidopsis Thaliana. Plant Physiol. 163, 1107– 1117. Meyer, D., Lauber, E., Roby, D., Arlat, M. and Kroj, T. (2005) Optimization of pathogenicity assays to study the Arabidopsis Thaliana-Xanthomonas Campestris Pv. Campestris Pathosystem. Mol. Plant Pathol. 6(3), 327– 333. Wiley Online Library Mohnen, D. (2008) Pectin structure and biosynthesis. Curr. Opin. Plant Biol. 11(3), 266– 277. Nafisi, M., Stranne, M., Fimognari, L. etal. (2015) Acetylation of cell wall is required for structural integrity of the leaf surface and exerts a global impact on plant stress responses. Front. Plant Sci. 6, 550. O\'Connell, R.J., Thon, M.R., Hacquard, S. etal. (2012) Lifestyle transitions in plant pathogenic Colletotrichum Fungi deciphered by genome and transcriptome analyses. Nat. Genet. 44(9), 1060– 1065. Osorio, S., Bombarely, A., Giavalisco, P., Usadel, B., Stephens, C., Aragüez, I., Medina-Escobar, N., Botella, M.A., Fernie, A.R. and Valpuesta, V. (2011) Demethylation of Oligogalacturonides by FaPE1 in the fruits of the wild strawberry Fragaria Vesca triggers metabolic and transcriptional changes associated with defence and development of the fruit. J. Exp. Bot. 62(8), 2855– 2873. Pfaffl, M.W. (2001) A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 29(9), e45. Randoux, B., Renard-Merlier, D., Mulard, G., Rossard, S., Duyme, F., Sanssené, J., Courtois, J., Durand, R. and Reignault, P. (2010) Distinct defenses induced in wheat against powdery mildew by acetylated and nonacetylated oligogalacturonides. Phytopathology, 100(12), 1352– 1363. Rennie, E.A., Hansen, S.F., Baidoo, E.E.K., Hadi, M.Z., Keasling, J.D. and Scheller, H.V. (2012) Three members of the Arabidopsis glycosyltransferase family 8 are Xylan Glucuronosyltransferases1[W][OA]. Plant Physiol. 159(4), 1408– 1417. Schultink, A., Naylor, D., Dama, M. and Pauly, M. (2015) The role of the plant-specific ALTERED XYLOGLUCAN9 protein in Arabidopsis cell wall Polysaccharide O-Acetylation. Plant Physiol. 167(4), 1271– 1283. Schulze-Lefert, P. and Vogel, J. (2000) Closing the ranks to attack by powdery mildew. Trends Plant Sci. 5(8), 343– 348. Selig, M.J., Vinzant, T.B., Himmel, M.E. and Decker, S.R. (2009) The effect of lignin removal by alkaline peroxide pretreatment on the susceptibility of corn stover to purified cellulolytic and xylanolytic enzymes. Appl. Biochem. Biotechnol. 155(1–3), 397– 406. Somerville, C., Bauer, S., Brininstool, G. etal. (2004) Toward a systems approach to understanding plant cell walls. Science, 306(5705), 2206– 2211. Stranne, M., Ren, Y., Fimognari, L. etal. (2018) TBL10 is required for O-Acetylation of pectic Rhamnogalacturonan-I in Arabidopsis Thaliana. Plant J. 96(4), 772– 785. Wiley Online Library Underwood, W. (2012) The plant cell wall: a dynamic barrier against pathogen invasion. Front. Plant Sci. 3, 85. https://doi.org/10.3389/fpls.2012.00085. Urbanowicz, B.R., Peña, M.J., Moniz, H.A., Moremen, K.W. and York, W.S. (2014) Two Arabidopsis proteins synthesize acetylated Xylan in vitro. Plant J. 80(2), 197– 206. Wiley Online Library Vogel, J. and Somerville, S. (2000) Isolation and characterization of powdery mildew-resistant Arabidopsis mutants. Proc. Natl Acad. Sci. USA, 97(4), 1897– 1902. Vogel, J.P., Raab, T.K., Schiff, C. and Somerville, S.C. (2002) PMR6, a Pectate Lyase-Like gene required for powdery mildew susceptibility in Arabidopsis. Plant Cell, 14(9), 2095– 2106. Vogel, J.P., Raab, T.K., Somerville, C.R. and Somerville, S.C. (2004) Mutations in PMR5 result in powdery mildew resistance and altered cell wall composition. Plant J. 40(6), 968– 978. Wiley Online Library Volpi, C., Janni, M., Lionetti, V., Bellincampi, D., Favaron, F. and D\'Ovidio, R. (2011) The ectopic expression of a pectin methyl esterase inhibitor increases pectin methyl esterification and limits fungal diseases in wheat. Mol. Plant Microbe Interact. 24(9), 1012– 1019. Wang, M., Roux, F., Bartoli, C., Huard-Chauveau, C., Meyer, C., Lee, H., Roby, D., McPeek, M.S. and Bergelson, J. (2018) Two-way mixed-effects methods for joint association analysis using both host and pathogen genomes. Proc. Natl Acad. Sci. USA, 115(24), E5440– E5449. Weßling, R. and Panstruga, R. (2012) Rapid quantification of plant-powdery mildew interactions by qPCR and conidiospore counts. Plant Methods, 8(1), 35. https://doi.org/10.1186/1746-4811-8-35. Xin, Z., Mandaokar, A., Chen, J., Last, R.L. and Browse, J. (2007) Arabidopsis ESK1 encodes a novel regulator of freezing tolerance. Plant J. 49(5), 786– 799. Wiley Online Library Yin, L., Verhertbruggen, Y., Oikawa, A. etal. (2011) The cooperative activities of CSLD2, CSLD3, and CSLD5 are required for normal Arabidopsis development. Mol. Plant, 4(6), 1024– 1037. Yuan, Y., Teng, Q., Zhong, R. and Ye, Z.-H. (2013) The Arabidopsis DUF231 domain-containing protein ESK1 mediates 2-O- and 3-O-Acetylation of Xylosyl residues in Xylan. Plant Cell Physiol. 54(7), 1186– 1199. Yuan, Y., Teng, Q., Zhong, R. and Ye, Z.-H. (2016) TBL3 and TBL31, two Arabidopsis DUF231 domain proteins, are required for 3-O-Monoacetylation of Xylan. Plant Cell Physiol. 57(1), 35– 45. Zhu, X.F., Sun, Y., Zhang, B.C., Mansoori, N., Wan, J.X., Liu, Y., Wang, Z.W., Shi, Y.Z., Zhou, Y.H. and Zheng, S.J. (2014) TRICHOME BIREFRINGENCE-LIKE27 affects aluminum sensitivity by modulating the O-Acetylation of Xyloglucan and Aluminum-binding capacity in Arabidopsis. Plant Physiol. 166(1), 181– 189. The full text of this article hosted at iucr.org is unavailable due to technical difficulties. Please check your email for instructions on resetting your password. If you do not receive an email within 10 minutes, your email address may not be registered, and you may need to create a new Wiley Online Library account. Can\'t sign in? Forgot your username? Enter your email address below and we will send you your username If the address matches an existing account you will receive an email with instructions to retrieve your username
