You have access HCN regulates cellular processes through posttranslational modification of proteins by S-cyanylation

1 Short title: Hydrogen cyanide regulation by S-cyanylation 1 2 3 4 Corresponding author: 5 Luis C. Romero 6 Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones 7 Científicas and Universidad de Sevilla 8 Avenida Américo Vespucio, 49, 9 41092 Seville, Spain 10 [email protected] 11 12 2 HCN regulates cellular processes through posttranslational 13 modification of proteins by S-cyanylation 14 15 16 Irene García, Lucía Arenas-Alfonseca, Inmaculada Moreno, Cecilia Gotor and Luis C. 17 Romero. 18 19 Instituto de Bioquímica Vegetal y Fotosíntesis, Consejo Superior de Investigaciones 20 Científicas and Universidad de Sevilla, Avenida Américo Vespucio, 49, 41092 Sevilla, 21 Spain 22 23 24 One-sentence summary 25 Hydrogen cyanide can act as a signaling molecule through posttranslational 26 modification of protein cysteine residues, leading to S-cyanylation. 27 . 28 29 Author contributions: 30 I.G. and L.A-A. performed the experiments, I.G. and L.C.R. designed the experiments 31 and analyzed the data; C.G. and L.C.R. supervised the experiments; I.M. provided 32 technical assistance; L.C.R wrote the article with the contributions of all authors. 33 34 Funding: 35 This work was supported in part by the European Regional Development Fund through 36 the Agencia Estatal de Investigación grant BIO2016-76633-P. L. A-A. was supported 37 by the Ministerio de Economia y Competitividad through the program of Formación de 38 Personal Investigador. 39 40 Correspondence: Luis C. Romero. E-mail: [email protected] 41 42 3 ABSTRACT 43 44 Hydrogen cyanide (HCN) is coproduced with ethylene in plant cells and is 45 primarily enzymatically detoxified by the mitochondrial ß-CYANOALANINE 46 SYNTHASE (CAS-C1). Permanent or transient depletion of CAS-C1 activity in 47 Arabidopsis (Arabidopsis thaliana) results in physiological alterations in the plant that 48 suggest that HCN acts as a gasotransmitter molecule. Label-free quantitative proteomic 49 analysis of mitochondrially enriched samples isolated from the wild type and cas-c1 50 mutant revealed significant changes in protein content, identifying 451 proteins that are 51 absent or less abundant in cas-c1 and 353 proteins that are only present or more 52 abundant in cas-c1. Gene ontology classification of these proteins identified proteomic 53 changes that explain the root hairless phenotype and the altered immune response 54 observed in the cas-c1 mutant. The mechanism of action of cyanide as a signaling 55 molecule was addressed using two proteomic approaches aimed at identifying the S- 56 cyanylation of cysteine as a posttranslational modification of proteins. Both the 2- 57 imino-thiazolidine chemical method and the direct untargeted analysis of proteins using 58 LC-MS/MS identified a set of 163 proteins susceptible to S-cyanylation that included 59 SEDOHEPTULOSE 1,7-BISPHOSPHATASE (SBPase), the PEPTIDYL-PROLYL 60 CIS-TRANS ISOMERASE 20-3 (CYP20-3), and ENOLASE 2 (ENO2). In vitro 61 analysis of these enzymes showed that S-cyanylation of SBPase Cys74, CYP20-3 62 Cys259, and ENO2 Cys346 residues affected their enzymatic activity. GO classification 63 and protein-protein interaction cluster analysis showed that S-cyanylation is involved in 64 the regulation of primary metabolic pathways, such as glycolysis, and the Calvin and S- 65 adenosylmethionine cycles. 66 67 4 INTRODUCTION 68 69 Hydrogen cyanide (HCN) is a gaseous acid produced in nature by a wide variety 70 of microorganisms such as fungi, bacteria and algae (Knowles, 1976), as well as in 71 cyanogenic plant food such as almonds (Prunus dulcis), millet sprouts (Panicum 72 miliaceum), cassava roots (Manihot esculenta) or lima beans (Phaseolus lunatus), 73 which are capable of accumulating significant amounts of cyanide, produced by the 74 hydrolysis of cyanogenic glycosides (Hanschen et al., 2014; Baskar et al., 2016; 75 Zidenga et al., 2017). Non-cyanogenic plant species also produce cyanide as a 76 byproduct of some metabolic processes, such as during camalexin and ethylene 77 biosynthesis (Peiser et al., 1984; Yip and Yang, 1988; Bottcher et al., 2009). 78 HCN can be partially dissolved in water and dissociates into H+ and CN– 79 particularly in basic solutions. Because the cytosolic pH is close to 7, it is expected that 80 cyanide would mostly present as hydrogen cyanide in this compartment, based on its pK 81 of 9.3. The moderate lipid solubility and small size of the HCN molecule allow it to 82 rapidly cross membranes and enter subcellular compartments. The anion cyanide is 83 highly toxic because it reacts with Schiff base intermediates and keto compounds to 84 produce cyanohydrins and nitrile derivatives, and because it chelates divalent and 85 trivalent metal ions in metalloproteins. In mitochondria, it can bind to the heme iron of 86 cytochrome c oxidase, inhibiting the functioning of the electron transport chain and 87 acting as a potent inhibitor of respiration (Isom and Way, 1984). Cyanide is also a 88 potent inhibitor of photosynthesis since it binds copper in plastocyanin, thereby 89 inhibiting plastocyanin-dependent electron transport to Photosystem I (Berg and 90 Krogmann, 1975) and dark CO2 assimilation (Bishop and Spikes, 1955; Trebst et al., 91 1960). However, cyanide toxicity in chloroplasts occurs in the dark and is partially 92 reversible by illumination in the presence of an electron acceptor (Bishop and Spikes, 93 1955; Cohen and McCarty, 1976). 94 Free cyanide, produced by metabolic pathways or by the degradation of 95 cyanogenic glycosides, is detoxified by rhodanese or primarily assimilated into amino 96 acids by the action of mitochondrial ß-cyanoalanine synthase (CAS-C1), a pyridoxal 97 phosphate-dependent enzyme that uses cysteine to detoxify cyanide by converting the 98 cyanide and cysteine into hydrogen sulfide (H2S) and β-cyanoalanine (Ressler et al., 99 1969; Hatzfeld et al., 2000). The H2S produced in the mitochondria inhibits cytochrome 100 c oxidase as potently as cyanide and requires detoxification by the mitochondrial O- 101 5 acetylserine(thiol)lyase isoform, which catalyzes the incorporation of sulfide into O- 102 acetylserine to produce cysteine, thus generating a cyclic pathway in the mitochondria 103 (Alvarez et al., 2012; Nicholls et al., 2013). 104 Despite its toxicity, cyanide has been proposed to act as a regulator of several 105 biological processes such as seed dormancy and germination (Taylorson and Hendricks, 106 1973; Bethke et al., 2006; Garcia et al., 2014), resistance to fungal and viral infection 107 (Chivasa and Carr, 1998; Wong et al., 2002; Seo et al., 2011), susceptibility to the 108 fungus Botrytis cinerea, and increased tolerance to the bacterium Pseudomonas 109 syringae pv. tomato DC3000, as well as to the beet curly top virus (Lozano-Durán et al., 110 2012; Garcia et al., 2013). Arabidopsis (Arabidopsis thaliana) null mutants of the 111 mitochondrial ß-cyanoalanine synthase, CAS-C1, accumulate cyanide to apparently 112 nontoxic levels, as the plants are completely viable, but show a root hairless phenotype 113 suggesting a signaling role in root development (Garcia et al., 2010; Garcia et al., 2014; 114 Arenas-Alfonseca et al., 2018). The root hair defect is phenocopied by the addition of 115 cyanide to the growth medium and reversed by the addition of the antidote 116 hydroxocobalamin (Garcia et al., 2010; Thompson and Marrs, 2012). A fused CAS-C1- 117 GFP protein under the control of the CAS-C1 promoter clearly localized in 118 mitochondria, but showed a tip-preferred localization during root hair growth (Arenas- 119 Alfonseca et al., 2018). Genetic crosses between the cas-c1 mutant and the scn1 or rhd2 120 root hair mutants were performed, and the detailed phenotypic and molecular 121 characterization of the double mutants demonstrated that the scn1 mutation is epistatic 122 to cas-c1, and cas-c1 is epistatic to the rhd2 mutation, indicating that CAS-C1 acts in 123 the early steps of the root hair development process. In addition, CAS-C1 function is 124 independent of ROS production and the direct inhibition of NADPH oxidase by cyanide 125 (Arenas-Alfonseca et al., 2018). 126 Cyanide functions as a signaling molecule in the response to pathogens as 127 indicated by the transient repression of the CAS-C1 transcript and accumulation of 128 cyanide when plants were infected with an avirulent P. syringae pv. tomato DC3000 129 avrRpm1 strain (Garcia et al., 2013). Although transient cyanide accumulation in 130 response to specific pathogens can induce a controlled accumulation of ROS for 131 signaling purposes, a direct effect of cyanide as a signaling molecule cannot be 132 excluded. The cyanide ion can attack cystine peptides or disulfides by nucleophilic 133 displacement on a sulfur atom of the disulfide forming a thiocyanate derivative 134 according to the reaction 135 6 R-S-S-R + CN- ⇆ R-S- + R-S-CN 136 which may be considered a redox reaction (Gawron, 1966). This reaction may 137 occur at intra- and/or inter-chain disulfide linkages at pH 7.0; however, at alkaline pH, 138 the direction of the reaction would be towards the formation of the thiocyanate 139 derivative (Gawron, 1966; Wagner and Davis, 1966). 140 The reaction of cyanide with cystine-containing proteins results in the formation 141 of an S-cyanylated cysteine motif that cycles and is subsequently cleaved to release an 142 amino terminal-peptide at one end and a 2-imino-thiazolidine-4-carboxylyl COOH- 143 terminal peptide at the other (Catsimpoolas and Wood, 1966; Fasco et al., 2007). The 144 reaction of cyanide ions with disulfide bridges within polypeptides has been considered 145 to be similar to that of other small molecules, such as NO or H2S, that react with Cys 146 residues to produce Cys modifications (Yamasaki et al., 2016; Aroca et al., 2017b; 147 Aroca et al., 2018). 148 In this study, we thoroughly examined the function of cyanide and the enzyme 149 ß-cyanoalanine synthase, which regulates the accumulation/detoxification of cyanide, in 150 the physiology of the cell through a proteomic approach. In addition, we analyzed the 151 molecular mechanism by which cyanide can act as a signaling molecule. 152 153 154 RESULTS 155 156 Quantitative proteomic analysis of mitochondrial ß-cyanoalanine synthase CAS- 157 C1 null mutant 158 The absence of ß-cyanoalanine synthase in the mitochondria results in a slight 159 increase in total cyanide in whole seedlings by 1.5-fold in the null cas-c1 mutant and a 160 3-fold accumulation after treatment with the ethylene/cyanide precursor 1- 161 aminocyclopropane-1-carboxylic acid (ACC) (Garcia et al., 2010). To thoroughly 162 investigate the role of CAS-C1 in cellular physiology and decipher the functional 163 mechanism that explains the observed phenotype in the cas-c1 mutant, we characterized 164 the protein composition and relative abundance of root extracts from wild type and cas- 165 c1 mutants using proteomics SWATH-MS (Sequential Window Acquisition of all 166 THeoretical spectra-Mass Spectrometry) technology. 167 Protein samples were isolated from root hydroponic cultures using density- 168 gradient centrifugation aiming to obtain a better enrichment of mitochondrial proteins 169 7 (Table S1). Extracted proteins from three biological replicates of the wild type and three 170 from the cas-c1 mutant line were digested, and the peptide solutions analyzed by a 171 shotgun data-dependent acquisition (DDA) approach to generate the spectral library. 172 After integrating the six datasets, a total of 11,122 peptides (1% FDR and 96% 173 confidence) and 1,734 unique proteins (1% FDR) were identified (Supplemental Dataset 174 1). To quantify them using SWATH acquisition, the same six biological samples were 175 analyzed twice each (technical replicas) by a data-independent acquisition (DIA) 176 method using the LC gradient and LC-MS equipment described to generate the spectral 177 library, but instead using the SWATH acquisition method described in the Materials 178 and Methods section. Therefore, for quantitation, six datasets from wild type and six 179 from cas-c1 were generated and used for the analysis. The fragment spectra were 180 extracted for the twelve runs, and 2,035 ion transitions, 2,901 peptides, and 1,132 181 proteins were quantified. From the 1,132 quantified proteins (Supplemental Dataset 2), 182 551 had significantly different abundances in wild type and cas-c1 (p value < 0.05) 183 (Table S2). From these proteins, 97 were more abundant in the wild type exhibiting a 184 fold change > 1.5, and 71 were more abundant in cas-c1 with a fold change < 0.66. In 185 addition to the differentially abundant proteins, we detected in the spectral library 186 generated by the DDA approach that 354 proteins were only identified in the wild type 187 samples and were below the detection limit in cas-c1 (Table S3), and 282 were only 188 identified in the cas-c1 samples (Table S4). 189 The 451 proteins that were absent (354 proteins) or less abundant (97 proteins) 190 in cas-c1 were analyzed based on their assigned functions and classified into 28 191 functional groups (Table S5) using the MapMan nomenclature (Thimm et al., 2004; 192 Klie and Nikoloski, 2012). The most numerous sets corresponded to the general protein 193 group (bin 29), which included 17.1% of the total identified proteins with 77 elements 194 involved in protein degradation (25 elements, primarily of the ubiquitin-proteasome 195 system), protein synthesis (20 elements), subcellular targeting (11 elements) and 196 posttranslational modification (10 elements). Many of the proteins identified are related 197 to pathogen response and biotic stress (Supplemental Fig. S1A), and GO classification 198 highlighted an important group containing proteins involved in signaling (13.3%) with 199 60 elements. Interestingly, 26 proteins of the receptor kinase family functioning in 200 pathogen response are present in this group and include six leucine-rich repeat protein 201 kinases, six domain of unknown function 26 (DUF26) receptor proteins and two lectin 202 kinases (Supplemental Fig. S1B and Table S5). Previous proteomic analyses in potato 203 8 (Solanum tuberosum) and Arabidopsis have also shown the presence of a significant 204 number of protein kinases in plant mitochondria related to communication and signaling 205 (Salvato et al., 2014; Rao et al., 2017). However, some DUF26 receptors are cytosolic 206 or plasma membrane proteins and were detected in this proteomic analysis because 207 although the protein preparation was enriched in mitochondria, it also contained other 208 membranes and organelles. 209 The 353 analyzed proteins that were only present (282 proteins) or more 210 abundant (71 proteins) in cas-c1 compared to the wild type were classified into 28 211 functional groups using MapMan (Table S6). Once again, the most abundant set 212 classified into the protein group with 59 elements involved in protein synthesis (22 213 elements), protein degradation (14 elements), subcellular targeting (9 elements) and 214 posttranslational modification (9 elements). An overrepresentation test of the GO terms 215 using PANTHER DB´s tool (Mi et al., 2017) to examine the abundant proteins showed 216 a 9.52-fold enrichment of proteins involved in the regulation of carbohydrate 217 metabolism, including four nuclear transcription factors NF-YC1, NF-YC3, NF-YC4 218 and NF-YC9; 9.33-fold enrichment in cellular amino acid catabolism and 7.35-fold 219 enrichment in ras-related proteins involved in phagocytosis and membrane trafficking 220 (Table S7). 221 Finally, to determine the functional relevance of the CAS-C1 mutation in the 222 physiology and development of the plant, we used the STRING database v10.5 223 (Szklarczyk et al., 2015) to analyze the protein-protein interaction network of the 353 224 proteins with increased abundance in cas-c1 compared to the wild type to determine 225 functional association of those proteins. Using the highest confidence (interaction score 226 0.900), a total of 295 protein-protein interactions were observed, and they were 227 significantly enriched (p-value < 0.000655) based on the given protein nodes. At least 228 four protein clusters that are biologically connected were clearly distinguished after 229 analysis (Fig. 1). The first cluster contains a subgroup with a complex of 17 230 ribonucleoproteins (yellow nodes) linked to a second cluster of 8 proteins of the 231 spliceosome pathway (red nodes), which is also connected through the E3 ubiquitin 232 ligase MAC3B component of the MOS4-associated complex to a subgroup containing 233 several cyclin-dependent kinases (CDKC1, CDKC2, CDKD1, CDKD3) and ubiquitin- 234 specific protease 12 and 13 (UBP12, UBP13). A third important cluster (blue nodes) 235 centered in ELI3-2 (ELICITOR ACTIVATED GENE 3-2) and CYP84A1 (ferulic-acid 236 5-hydrolase), both involved in lignin biosynthesis, comprises 8 peroxidase proteins 237 9 (PER7, PER16, PER23, PER27, PER32, PER44, PER45 and PER49) and IRX4 238 (cinnamoyl CoA-reductase, involved in the latter stages of lignin biosynthesis). The 239 fourth cluster (green nodes), centered on RAB GTPas-1A, comprises several subgroups 240 with proteins involved in the endomembrane system, regulators of membrane traffic 241 from the Golgi apparatus towards the endoplasmic reticulum, and includes RAB 242 GTPase-1b, 1c and 8C and 5 coatomer subunit proteins. This cluster also contains a 243 subgroup formed by 4 nuclear transcription factors, NF-YC1, NF-YC3, NF-YC4 and 244 NF-YC9 that regulate gene expression by modulating histone acetylation and 245 methylation. 246 247 Identification of S-cyanylation by the chemical 2-imino-thiazolidine method 248 The cyanide molecule possesses physico-chemical characteristics similar to 249 well-established signaling molecules such as NO, CO, or H2S (Gotor et al., 2017). In 250 addition to its affinity for metalloproteins, cyanide does not react with free thiol groups 251 but can react with cystine-containing proteins at cellular neutral and mild alkaline pH 252 and, by analogy to NO or H2S, modify protein residues, which subsequently alters 253 protein function. To explore the function of cyanide as a signaling molecule through 254 protein posttranslational modifications (PTMs), we studied the presence of proteins 255 with S-cyanylated-cysteine residues in cytosolic leaf and root extracts. 256 It is well described that S-cyanylated proteins can be cleaved into two parts, the 257 amino acid backbone from the N-terminus to the cyanilated cysteine residue and the 258 cycled 2-imino-thiazolidine-4-carboxylyl COOH-terminal peptide (Fig. 2A) 259 (Catsimpoolas and Wood, 1966; Fasco et al., 2007), a process that is favored at alkaline 260 conditions. Considering the above chemical properties of S-cyanylated cysteines, we 261 studied in vitro the endogenous presence of this modification in leaf protein extracts 262 from Arabidopsis wild type and cas-c1 mutant lines. Depletion of CAS-C1 activity 263 results in the accumulation of cyanide in 2-week-old seedlings, and this accumulation 264 can be increased by treatment with ACC, the precursor of ethylene and cyanide, via 265 ACC oxidases (Garcia et al., 2010). Therefore, the presence of endogenous S- 266 cyanylated proteins was analyzed in 2-week-old wild-type and cas-c1 leaves treated 267 with 100 µM ACC for 24 h by comparing the mobility shift or spot volume of the 268 proteins after cleavage with 1 M NH4OH (Fig. 2B) by the 2-imino-thiazolidine chemical 269 method. The protein samples from 2-week-old leaves were treated with NH4OH for 1 h 270 or 0.01 M NaOH as a control and separated by two-dimensional (2D) isoelectric 271 16 can act by forming adducts with the metal centers in proteins that contain iron or cobalt. 476 The cyano-iron complex in the heme prosthetic groups of metalloproteins inhibits the 477 activity of enzymes such as cytochrome c oxidase. A second mechanism involves the 478 posttranslational modification of protein cysteine residues leading to the S-cyanylation 479 of the protein. This process was thought not to take place under physiological 480 conditions in any organism, however S-cyanylated-Cys residues in human plasma 481 proteins, such as immunoglobulin G and serum albumin, are detected after cyanide 482 poisoning or in smoking persons and are used as biomarkers of potential cyanide 483 exposure (Fasco et al., 2007; Grigoryan et al., 2016). We have here identified S- 484 cyanylated-Cys residues in the proteins of leaf and root samples, in addition to 485 characterizing the molecular changes produced by genetic ß-cyanoalanine synthase 486 depletion and concomitant cyanide accumulation. Using two different physicochemical 487 approaches, the 2-imino-thiazolidine chemical method and direct untargeted analysis of 488 proteins using LC-MS/MS, we found that this modification occurs during physiological 489 growth conditions in leaf and root tissues, but can be increased by stimulating the 490 biosynthesis of ethylene and the concomitant cyanide production by treatment with the 491 ACC precursor. Although the number of S-cyanylated proteins detected is not high 492 (Table 1), we are probably underestimating the true number since we are only 493 identifying abundant proteins. Many PTMs such as carbonylation, nitrosylation, 494 persulfuration or nitration are identified using chemical labelling methods or by specific 495 antibodies, but so far we do not have available a method to enrich selectively for S- 496 cyanylated-Cys proteins that will allow us to purify and identify them (Alvarez et al., 497 2011; Moller et al., 2011; Zhang et al., 2014; Aroca et al., 2017a). 498 499 Role of S-cyanylation in cellular processes 500 Both methods used to identify the S-cyanylated Cys residues detected the ENO2 501 protein encoded by the LOS2/ENO2 locus, which functions in glycolysis for the 502 dehydration of 2-phosphoglycerate to produce phosphoenolpyruvate or acts as a 503 repressor of cMyc transcription by alternative translation of the locus to produce MBP-1 504 (Lee et al., 2002; Eremina et al., 2015). GO classification, enrichment and clustering 505 analysis of the 163 proteins that were identified to be susceptible to S-cyanylation 506 highlighted the role of these proteins in primary carbon fixation, glycolysis, the TCA 507 cycle and in general, carbon metabolism, but also in chaperone-assisted protein folding 508 and SAM-cycle/DNA methylation (Fig. 9). The in vitro analyses performed with some 509 17 of the identified S-cyanylated proteins showed that S-cyanylation of CYP20-3 results in 510 the partial inhibition of its activity but results in the activation of ENO2. Since cyanide 511 action against cysteine residue only takes place over oxidized cysteines in the form of 512 disulfide bridges, the effect of S-cyanylation depends on the redox state of the target 513 protein. CYP20-3 can form two disulfide bridges, which are required for its isomerase 514 activity and are subject to regulation by thioredoxin (Motohashi et al., 2003), but S- 515 cyanylation of Cys206 in its oxidized state results in the irreversible inactivation of its 516 activity. Most of the enzymes of central carbon metabolism, photosynthetic carbon 517 fixation, glycolysis and the TCA cycle are known to be light/dark regulated; therefore, 518 they are inhibited in the dark by the oxidation of active site cysteines, but reactivated in 519 the light by the action of ferredoxin/thioredoxin systems (Buchanan and Balmer, 2005; 520 Geigenberger et al., 2005). Therefore, cyanide modification of these enzymes in their 521 oxidized state must result in the irreversible inactivation of their activity as shown with 522 CYP20-3. In this case, we can hypothesize that S-cyanylation acts as a switch to 523 definitively stop the light/dark regulation of this activity under some specific conditions. 524 ENO2 activity, the ninth step of glycolysis, is higher during oxidative conditions 525 and during KCN treatment, which indicates that S-cyanylation switches on the enzyme 526 activity (Fig. 8 and (Anderson et al., 1998)). Thus, S-cyanylation can accelerate the 527 degradation of sugar for ATP production to compensate for the inhibition of the 528 cytochrome oxidase pathway. The cas-c1 mutant shows higher transcript levels of the 529 alternative oxidase AOX1a and higher respiration rates, but the levels of ATP are 530 unaltered (Garcia et al., 2010; Arenas-Alfonseca et al., 2018). Thus, transient cyanide 531 accumulation associated with developmental or environmental stress can potentially 532 generate metabolic perturbations through the inhibition of enzymes of central carbon 533 metabolism and the induction of the AOX gene that controls the production of signaling 534 molecules such as ROS. 535 A second cyanide signaling mechanism might be the direct effect of S- 536 cyanylation on protein localization. Many of the enzymes identified as susceptible to S- 537 cyanylation, such as the glycolytic enzymes, including ENO2 and GAPDH, and 538 proteins of the SAM-cycle, are moonlighting proteins involved either in metabolic 539 pathways and gene transcription, but they lack nuclear localization sequences (Lee et 540 al., 2002; Zaffagnini et al., 2013; Boukouris et al., 2016). Several of the enzymes of the 541 glycolytic pathway are redox regulated and are more active in their reduced form, such 542 as glyceraldehyde 3-phosphate dehydrogenase (Holtgrefe et al., 2008). However, 543 18 posttranslational modification of the cytosolic GAPDH GapC isoform by S- 544 persulfidation of Cys160 slightly regulates its activity, but is critical to determine the 545 cytosolic/nuclear partitioning of the enzyme (Aroca et al., 2015; Aroca et al., 2017b). In 546 addition to its enolase activity, the LOS2/ENO2 locus can be alternatively translated 547 from a second AUG translation initiation codon at position +93 to form the protein C- 548 MYC BINDING PROTEIN 1 (MBP-1), which was found to bind c-myc-like elements in 549 the promoter of the STZ/ZAT10 gene acting as a transcriptional repressor (Lee et al., 550 2002). Both ENO2 and MBP-1 contain the Cys346 identified as being susceptible to S- 551 cyanylation (Fig. 7). Although we have not tested the effect of S-cyanylation in the 552 nuclear localization of ENO2 or the MBP-1 protein, previous transcriptomic analyses 553 show the repression of STZ/ZAT10 in the root tissue of cas-c1, suggesting the 554 predominant nuclear localization of MBP-1 (Garcia et al., 2010). 555 In addition to the glycolytic enzymes, we also detected a large number of S- 556 cyanylated proteins involved in the S-adenosyl-L-methionine (SAM) cycle and DNA 557 and histone methylation such as methionine synthase MS1 and MS2, S-adenosyl-L- 558 homocysteine hydrolase SAHH1/HOG1, SAHH2, and DNA 559 METHYLTRANSFERASE 2 (DMT2). Methylation occurs in the nucleus through the 560 metabolite S-adenosylmethionine, which functions as the primary methyl group donor, 561 and the action of methytransferases that transfer the methyl group of SAM producing S- 562 adenosyl-L-homocysteine (SAH), which is a potent inhibitor of methyltransferase 563 activity (Sauter et al., 2013). SAHH1/HOG is localized either in the cytoplasm or 564 nucleus, but it lacks any nuclear localization signal (Lee et al., 2012). Null T-DNA 565 insertional mutants in SAHH1gene, hog1 mutant, shows embryo-lethal phenotypes 566 (Rocha et al., 2005). However, an sahh1 mutant that produces the normal SAHH1 567 protein, but in reduced amounts, maintains its fertility level but shows a deficiency in 568 root hair development (Wu et al., 2009). Since the S-cyanylation of SAHH1 does not 569 result in the severe embryo-lethality of cas-c1, the Cys modification may be affecting 570 the nuclear localization of the protein rather than inhibiting the SAHH1 activity. Based 571 on all previous data, we suggest that the effect of S-cyanylation on protein localization 572 is plausible and deserves further investigation. 573 574 CONCLUSION 575 In this study, we demonstrate that the accumulation of cyanide by the repression 576 of the ß-cyanoalanine synthase activity may induce a nonenzymatic PTM into the Cys 577 19 motif. Although many nonenzymatic PTMs have been proposed to be indicators of 578 different cellular stresses and used as biomarkers of aging, such as the oxidation of 579 several amino acids, carbonylation or carbamylation, these modifications can also occur 580 in nonstressed and healthy cells and can regulate and influence nonpathogenic cellular 581 processes (Moller et al., 2011; Harmel and Fiedler, 2018). Unlike other nonenzymatic 582 PTMs such as nitrosylation, disulfide formation and acylation, S-cyanylation seems to 583 be a nonreversible modification since reductants cannot reverse its effect in the 584 analyzed proteins. The fact that S-cyanylaton occurs on oxidized cysteines in the form 585 of disulfide bridges can cause a change in the properties and function of the proteins. 586 However, a much more extensive study on the effect of this PTM will be necessary to 587 fully understand the functionality of this modification. 588 589 590 MATERIALS AND METHODS 591 592 Plant material and growth conditions 593 Arabidopsis (Arabidopsis thaliana) wild type ecotype Col-0 and the cas-c1 T- 594 DNA insertion mutant (cas-c1; SALK_103855) (Garcia et al., 2010) were grown in soil 595 in a photoperiod of 16 h of white light (120 mol m-2 s-1) at 20 ºC and 8 h of dark at 18 596 ºC or in hydroponic culture in a photoperiod of 8 h of white light (120 mol m-2 s-1) at 597 20 ºC and 16 h of dark at 18 ºC (Bermudez et al., 2010). For the hydroponic cultures, 598 seeds were surface-sterilized, incubated for 4 days at 4º C and germinated for one week 599 in rockwool. Seedlings were introduced in the wells of hydroponic boxes containing 5 L 600 of nutrient solution. The composition of the nutrient solution was as follows: 1 mM 601 KH2PO4, 1 mM MgSO4 · 7H2O, 15 µM MnSO4 · H2O, 1 µM CuSO4 · 5 H2O, 1 µM 602 ZnSO4 · 7 H2O, 30 µM H3BO3, 28 nM (NH4)6Mo7O24 · 4 H2O, 100 nM CoSO4 · 7 H2O, 603 10 mM Ca(NO3)2 · 4 H2O and 90 mM sequestrene 138 Fe G100 (Syngenta, Spain). All 604 nutrient solutions were changed twice per month. 605 606 Mitochondrial protein extraction 607 Mitochondria-enriched samples were isolated from Arabidopsis roots using 608 differential and density-gradient centrifugation as described (Struglics et al., 1993) but 609 with modifications. A total of 30 g of Arabidopsis hydroponic root tissue was ground 610 using a mortar and pestle with liquid nitrogen and resuspended into an extraction buffer 611 20 containing 15 mM MOPS (pH 7.4), 1.5 mM EDTA, 0.25 M sucrose, 0.4% (w/v) BSA 612 and 0.6% (w/v) PVP-40, using 3 ml of buffer g-1 fresh weight, and the homogenates 613 were filtered through two layers of Miracloth (Calbiochem). The filtered homogenate 614 was centrifuged at 2200 g for 5 min and the supernatant transferred to a new tube, 615 centrifuged again sequentially at 3200 g and 4500 g each time for 5 min until a non- 616 green supernatant was obtained. This final supernatant was centrifuged at 18000 g for 617 30 min. The pellet was resuspended in 1 ml of gradient buffer (20 mM MOPS, pH 7.2, 618 0.6 M sucrose). The separation of the cellular organelles was obtained using a self- 619 generated 45 and 27% Percoll gradient. The gradient was centrifuged at 20000 g for 20 620 min in an ultracentrifuge using an SW28.1 Ti rotor (Beckman) at 4°C. The 621 mitochondrial band was taken out, diluted 10 times with the wash buffer (0.3 M 622 sucrose, 1 mM EDTA, 10 mM MOPS at pH 7.2), and centrifuged at 20000 g for 15 min 623 four times to wash the organelles free from Percoll. The last pellet was resuspended in 624 300 µL of lysis buffer (50 mM potassium phosphate buffer, pH 7.0, and 0.1% (v/v) 625 Triton X-100) and the mitochondrial samples disrupted using repeating freeze-thaw 626 cycles and ultrasonic lysis for 1.5 min. The samples were centrifuged at 20000 g for 15 627 min. Proteins from the supernatant were precipitated with 10% (v/v) trichloroacetic acid 628 and ice-cold acetone and resuspended in 20 mM Tris-HCl, pH 8.0. 629 630 Identification of protein S-cyanylation using the 2-imino-thiazolidine chemical 631 method 632 Arabidopsis cytosol-enriched leaf protein extract was isolated to perform 2D 633 electrophoresis. For this, 200 mg of frozen leaf tissue was ground in liquid nitrogen 634 using a mortar and pestle. The resulting powder was homogenized in 2 mL of water and 635 ultracentrifuged at 176000 g for 1 h at 4 °C. The protein concentration in the 636 supernatant was then quantified and used for the ammonium hydroxide treatment. Four 637 hundred micrograms of protein were incubated with 1 M of ammonium hydroxide or 638 0.01 M of sodium hydroxide for 1 h at 20ºC. Proteins were then dried in a speed 639 vacuum, resuspended in 300 µL of 1 mM Tris - 10 mM EDTA, pH 7.5, and precipitated 640 with 10% TCA (v/v) overnight at 4ºC, centrifuged at 20000 g for 15 min at 4ºC, and the 641 pellet washed twice with 100% acetone. After the final centrifugation step, the protein 642 pellet was air-dried and resolubilized in 8 M urea, 2% (w/v) CHAPS, 50 643 mM dithiothreitol, 0.2% (v/v) ampholytes (Bio-Lyte 3-10 buffer, Bio-Rad), and a trace 644 of bromophenol blue. 645 21 The proteins were applied to a linear immobilized pH gradient strip, pH 4 to 7 646 during rehydration. Isoelectric focusing was performed at 22°C in the Protean IEF Cell 647 (Bio-Rad) in three steps: (1) 250 V for 15 min, (2) 10,000 V for 1 h, and (3) 10,000 V 648 up to 40,000 V. The intensity was fixed at 50 μA per immobilized pH gradient strip to 649 avoid overheating the system. For the second dimension analysis, the equilibrated gel 650 strips were placed on top of vertical polyacrylamide gels (12% w/v acrylamide and 651 SDS) (Laemmli, 1970). Electrophoresis was conducted at 240 V and 35 mA per gel for 652 5 h using the Protean II xi Cell (Bio-Rad). Protein spots were visualized after 653 Coomassie brilliant blue staining. 654 The 2D images were analyzed using laser densitometry scanning with a GS-800 655 calibrated densitometer (Bio-Rad), and three high-quality gels from independent protein 656 extracts and conditions were analyzed using the PDQuest software version 7.0 (Bio- 657 Rad). Protein spots of interest were excised, trypsin digested and identified using 658 MALDI-mass spectrometry analysis at the Proteomic Facility of the Institute of Plant 659 Biochemistry and Photosynthesis as previously described (Bermudez et al., 2012) 660 661 Identification of S-cyanylated proteins using LC-MS/MS analysis 662 To identify S-cyanylated proteins, tryptic digestion was performed in solution of 663 root extracts, and peptide analysis was carried out using a nano liquid chromatography 664 system coupled to a high-speed mass spectrometer at the Proteomics Facility of the 665 Centro Nacional de Biotecnología, Spain. An aliquot of 40 μg of protein from each 666 treatment group was digested with sequencing-grade modified trypsin (Sigma-Aldrich) 667 at 37 ºC overnight on a shaker. Three biological replicates for both conditions (wild 668 type and cas-c1) were analyzed using the same setup and the parameters previously 669 described (Aroca et al., 2017a). 670 The mass spectrometry data have been deposited in the ProteomeXchange 671 Consortium via the PRIDE (Vizcaino et al., 2016) partner repository with the dataset 672 identifier PXD009812. 673 Protein function analysis and classification was performed using MapMan 674 (Thimm et al., 2004; Klie and Nikoloski, 2012) and clustering analysis by using the 675 STRING database (Szklarczyk et al., 2015). The protein-protein association network 676 was assessed in STRING using interaction evidences coming from experimental data on 677 protein-protein interactions, interaction prediction from co-expression analysis, and 678 22 known pathways and protein complexes from curated databases and co-occurrence in 679 publications (Szklarczyk et al., 2015; Szklarczyk et al., 2017) 680 681 Protein relative quantitation by SWATH-MS acquisition and analysis 682 Protein samples from mitochondrial extracts for SWATH-MS measurements 683 were alkylated and trypsin-digested as described (Vowinckel et al., 2013; Ortea et al., 684 2018), and performed at the Proteomic Facility of the Insituto Maimónides de 685 Investigación Biomédica de Córdoba, Spain. 686 A data-dependent acquisition (DDA) approach using nano-LC-MS/MS was first 687 performed to generate the SWATH-MS spectral library as described (Ortea et al., 688 2018). 689 The peptide and protein identifications were performed using Protein Pilot 690 software (version 5.0.1, Sciex) with the Paragon algorithm. The searching was 691 conducted against a revised UniProt Swiss-Prot Arabidopsis thaliana protein database 692 (August 2016), specifying iodoacetamide with other possible Cys modifications. The 693 false discovery rate (FDR) was set to 0.01 for both peptides and proteins. The MS/MS 694 spectra of the identified peptides were then used to generate the spectral library for 695 SWATH peak extraction using the add-in for PeakView Software (version 2.1, Sciex) 696 MS/MSALL with SWATH Acquisition MicroApp (version 2.0, Sciex). Peptides with a 697 confidence score above 99% (as obtained from the Protein Pilot database search) were 698 included in the spectral library. 699 For relative quantitation using SWATH analysis, the same samples used to 700 generate the spectral library were analyzed using a data-independent acquisition (DIA) 701 method. Each sample (2 μL) was analyzed using the LC-MS equipment and LC 702 gradient described above to build the spectral library but instead used the SWATH-MS 703 acquisition method. The method consisted of repeating a cycle that consisted of the 704 acquisition of 34 TOF MS/MS scans (230 to 1500 m/z, 100 ms acquisition time) of 705 overlapping sequential precursor isolation windows of 25 m/z width (1 m/z overlap) 706 covering the 400 to 1250 m/z mass range with a previous TOF MS scan (400 to 1250 707 m/z, 50 ms acquisition time) for each cycle. The total cycle time was 3.5 s. 708 The targeted data extraction of the fragment ion chromatogram traces from the 709 SWATH runs was performed by PeakView (version 2.1) with the MS/MSALL with 710 SWATH Acquisition MicroApp (version 2.0). This application processed the data using 711 the spectral library created from the shotgun data. Up to 10 peptides per protein and 7 712 23 fragments per peptide were selected, based on signal intensity. Any shared and modified 713 peptides were excluded from the processing. Windows of 12 min and 20 ppm width 714 were used to extract the ion chromatograms. SWATH quantitation was attempted for all 715 proteins in the ion library that were identified by ProteinPilot with an FDR below 1%. 716 The extracted ion chromatograms were then generated for each selected fragment ion. 717 The peak areas for the peptides were obtained by summing the peak areas from the 718 corresponding fragment ions. PeakView computed an FDR and a score for each 719 assigned peptide according to the chromatographic and spectral components. Only 720 peptides with an FDR below 5% were used for protein quantitation. Protein quantitation 721 was calculated by adding the peak areas of the corresponding peptides. To test for 722 differential protein abundance between the two groups, MarkerView (version 1.2.1, 723 Sciex) was used for signal normalization. 724 The mass spectrometry data have been deposited in the ProteomeXchange 725 Consortium via the PRIDE (Vizcaino et al., 2016) partner repository with the dataset 726 identifier PXD009929. 727 728 Expression and purification of recombinant His-tagged proteins 729 The complementary DNAs encoding the mature chloroplastic SBPase 730 (At3g55800), the mature chloroplastic CYP20-3 (At3g62030), and the complete 731 cytosolic ENO2 (At2g36530) were cloned into the pDEST17 vector (Invitrogen) with 732 Gateway Technology (Invitrogen) to express an N-terminal 6-His-tagged protein using 733 the Escherichia coli expression system. For SBPase and ENO2 protein expression, 734 transformed E. coli BL21 cell cultures at an optical density at 600 nm of 0.4 were 735 treated with 0.5 mM isopropyl--D-thiogalactopyranoside and incubated for an 736 additional 2 h at 37°C. CYP20-3 was also obtained by IPTG induction, but in this case 737 the induction was for 4 h at 30ºC. Purification was performed using nickel resin binding 738 under nondenaturing conditions using a Ni-NTA Purification System (Invitrogen) 739 according to the manufacturer’s instructions. Recombinant protein production and 740 purification were assessed by SDS-PAGE using 12% (w/v) polyacrylamide gels and 741 Coomassie brilliant blue staining. 742 743 Electrophoretic mobility 744 Nickel-affinity purified recombinant CYP20-3 was incubated at 4 ºC with 745 potassium cyanide, hydrogen peroxide, DTT or combinations of them for different time 746 24 periods and different concentrations as indicated, followed by a further incubation with 747 the alkylating agent 2-iodoacetamide at a concentration of 100 mM for 30 min. 748 Subsequently, 20 μg of the samples were subjected to SDS-PAGE using 15% (w/v) 749 polyacrylamide gels, either under nonreducing or reducing conditions depending on the 750 presence of 2-mercaptoethanol in the loading buffer. After electrophoresis, the proteins 751 were visualized using Coomassie Brilliant Blue staining. 752 753 Enzyme activity assays 754 The PPIase activity of the nickel-affinity purified recombinant CYP20-3 was 755 measured using a chymotrypsin-coupled assay as previously described (Motohashi et 756 al., 2003) with some modifications. An assay mixture containing 44 mM HEPES-NaOH 757 (pH 8.0), 1 µg of the recombinant CYP20-3 and 0.6 mg of chymotrypsin was incubated 758 for 5 min at 1 ºC. To initiate the reaction, 75 μM of the substrate N-succinyl-Ala-Ala- 759 Pro-Phe-p-nitroanilide (Sigma), previously dissolved in trifluoroethanol and LiCl, was 760 added. The absorbance at 367 nm was immediately monitored using UV 761 spectrophotometry for 1.5 min. The specific activities were calculated by using the 762 extinction coefficient of 13.4 mM-1 cm-1. 763 The enolase activity of the nickel-affinity purified recombinant ENO2 was 764 measured using a coupled assay system of lactate dehydrogenase and pyruvate kinase. 765 Enolase activity was monitored as a decrease in NADH absorbance at 340 nm and 766 estimated by using the molar extinction coefficient of NADH of 6.22 mM-1 cm-1 767 (Fukano and Kimura, 2014). 768 In all cases, one unit of activity is defined as the amount of enzyme that 769 catalyzed the production of 1 μmol product min−1. 770 771 Accession Numbers 772 Sequence data from this article can be found in the Web site 773 http://www.arabidopsis.org/ for Arabidopsis genes with locus identifiers provided in 774 this study. 775 776 Supplemental Data 777 The following supplemental materials are available. 778 Supplemental Dataset 1. Proteins identified by shotgun data dependent acquisition in 779 mitochondrial samples 780 25 Supplemental Dataset 2. Quantified proteins using the SWATH acquisition method in 781 mitochondrial samples 782 Supplemental Dataset 3. Identified proteins in ACC treated (R+) and untreated (R-) 783 root samples 784 Supplemental Table S1. Overrepresentation test in mitochondrial proteins identified in 785 the spectral library in mitochondrial samples 786 Supplemental Table S2. Proteins with different abundance in mitochondrial samples 787 between wild type and cas-c1 with p value <0.05 788 Supplemental Table S3. Identified proteins in the spectral library only present in 789 mitochondrial wild-type samples and absent in cas-c1 790 Supplemental Table S4. Identified proteins in the spectral library only present in 791 mitochondrial cas-c1 samples and absent in wild type 792 Supplemental Table S5. Functional classification of less abundant or absent proteins in 793 mitochondrial cas-c1 samples 794 Supplemental Table S6. Functional classification of proteins more abundant or only 795 present in mitochondrial cas-c1 samples 796 Supplemental Table S7. Overrepresentation test of the more abundant proteins in 797 mitochondrial cas-c1 samples 798 Supplemental Table S8. S-cyanylated proteins identified in wild-type and cas-c1 799 leaves as determined by the chemical 2-imino-thiazolidine method 800 Supplemental Table S9. Overrepresentation test of S-cyanylated proteins identified in 801 wild-type and cas-c1 leaves 802 Supplemental Table S10. Total proteins identified as susceptible to S-cyanylation in 803 leaf and root tissues 804 Supplemental Table S11. SBPase predicted ion types for S-cyanyl modified peptide. 805 The ions detected in the spectrum are highlighted in red color. This table refers to 806 Figure 4 807 Supplemental Table S12. CYP20-3 predicted ion types for the modified peptide. The 808 ions detected in the spectrum are highlighted in red color. This table refers to Figure 5 809 Supplemental Table S13. ENO2 predicted ion types for the modified peptide. The ions 810 detected in the spectrum are highlighted in red color. This table refers to Figure 7 811 Supplemental Figure S1. Schematic image of the signaling pathway and components 812 involved in biotic stress that are absent or less abundant in mitochondrial cas-c1 813 samples. A) Blue squares represent proteins classified in biotic stress according to GO 814 32 Bethke, P.C., Libourel, L.G.L., Reinohl, V., Jones, R.L. (2006). Sodium 1026 nitroprusside, cyanide, nitrite, and nitrate break Arabidopsis seed dormancy in a 1027 nitric oxide-dependent manner. Planta 223: 805-812. 1028 Bishop, N.I., Spikes, J.D. (1955). Inhibition by Cyanide of the Photochemical Activity 1029 of Isolated Chloroplasts. 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A mutant deficient in S-adenosylhomocysteine hydrolase in Arabidopsis 1229 shows defects in root-hair developmentThis paper is one of a selection of papers 1230 published in a Special Issue from the National Research Council of Canada – 1231 Plant Biotechnology Institute. Botany 87: 571-584. 1232 Yamasaki, H., Watanabe, N.S., Sakihama, Y., Cohen, M.F. (2016). An Overview of 1233 Methods in Plant Nitric Oxide (NO) Research: Why Do We Always Need to 1234 Use Multiple Methods? Methods Mol Biol 1424: 1-14. 1235 Yip, W.K., Yang, S.F. (1988). Cyanide metabolism in relation to ethylene production 1236 in plant tissues. Plant Physiol 88: 473-476. 1237 Zaffagnini, M., Fermani, S., Costa, A., Lemaire, S.D., Trost, P. (2013). Plant 1238 cytoplasmic GAPDH: redox post-translational modifications and moonlighting 1239 properties. Front Plant Sci 4: 450. 1240 Zhang, D., Macinkovic, I., Devarie-Baez, N.O., Pan, J., Park, C.M., Carroll, K.S., 1241 Filipovic, M.R., Xian, M. (2014). Detection of protein S-sulfhydration by a tag- 1242 switch technique. Angew Chem Int Ed Engl 53: 575-581. 1243 Zidenga, T., Siritunga, D., Sayre, R.T. (2017). Cyanogen Metabolism in Cassava 1244 Roots: Impact on Protein Synthesis and Root Development. Front Plant Sci 8: 1245 220. 1246 1247 1248 Figure 1. Protein-protein interaction network of the abundant proteins in mitochondrial cas-c1 samples . Round nodes represents proteins, and lines represent interactions. Dashed lines represent inter-cluster edges. Node modules indicated in red, yellow, blue and green colors were identified by k-means clustering PER7 PER45 PER32 PER27 PER16 PER23 PER44 PER49 AB Figure 2. Cyanylation and in vitro analysis in leaf samples.A) Chemical cleavage of the peptide chains on the N-terminal side of cyanylated cysteine residues. S-cyanylated peptide at alkaline pH results in the cleavage of the protein into two peptides: the N-terminal backbone at the Cys-CN residue and a cycled iminothiazoline derivative peptide. B) Representative 2D IEF-SDS PAGE protein profile of protein extracts of 2-week-old Arabidopsis cas-c1 mutant leaves treated with 100 µMACC for 24 h. Proteins were treated with 0.01 M NaOH (control samples) or 1 M NH4OH and were separated in the first dimension in gel strips (7 cm) with a linear pH gradient 4–7 and in 12% acrylamide vertical gels in the second dimension and stained with Coomassie-blue G-250. White circles show the mobility shift of a representative protein (SBPase). SBPase SBPase Control (0.01 M NaOH) Cleavage (1 M NH4OH) 47 pI 47 pI 15 - - 15 - 25 25 - 37 - - 37 - 50 50 - - 75 kDa Figure 3. Protein-protein interaction network of identified S-cyanylated proteins in leaf and root tissues.Round nodes represents proteins, and lines edges represent interactions. Dashed lines represent inter-cluster edges. Node modules indicated in red, yellow, blue and green colors were identified by k-means clustering. ATMS2 Figure 4. Identification of sedoheptulose 1, 7-bisphosphatase using mass spectrometry. (A) The protein was identified with a sequence coverage of 67%. The identified peptides are shown in red and the peptide containing S-cyanylated Cys74 is underlined. The modified Cys residue is shown in blue color. (B) Mass spectrum from the LC-MS/MS analysis of the tryptic peptide containing Cys74 of SBPase . Predicted ion types and the detected ions are shown in Table S11. ASedoheptulose 1, 7-bisphosphatase Protein sequence coverage: 67% Peptide score: 123 METSIACYSR GILPPSVSSQ RSSTLVSPPS YSTSSSFKRL KSSSIFGDSL 50 RLAPKSQLKA TKAKSNGAST VTKCEIGQSL EEFLAQATPD KGLRTLLMCM 100 GEALRTIAFK VRTASCGGTA CVNSFGDEQL AVDMLADKLL FEALQYSHVC 150 KYACSEEVPE LQDMGGPVEG GFSVAFDPLD GSSIVDTNFT VGTIFGVWPG 200 DKLTGITGGD QVAAAMGIYG PRTTYVLAVK GFPGTHEFLL LDEGKWQHVK 250 ETTEIAEGKM FSPGNLRATF DNSEYSKLID YYVKEKYTLR YTGGMVPDVN 300 QIIVKEKGIF TNVTSPTAKA KLRLLFEVAP LGLLIENAGG FSSDGHKSVL 350 DKTIINLDDR TQVAYGSKNE IIRFEETLYG TSRLKNVPIG VTA 393 B b10 m/z y8 y9 y10 y11 y13 1000800 1250 1500 1750 0 2.5 5.0 7.5 10.0 12.5 b10 64SNGASTVTKC[CN] EIGQSLEEFLAQATPDK90 y9 y11 y10 y13 Ion intensity m/z b2++ b2 y3 b5 b6 b7 bo8 b9 b10 y09 b12 b13 y11 y15 b15 b16 500 1000 1500 100 80 60 40 20 0 Figure 5. Identification of peptidyl-prolyl cis-trans isomerase CYP20-3 using mass spectrometry. (A) The protein was identified with a sequence coverage of 61%. The identified peptides are shown in red, and the peptide containing S-cyanylated Cys206 is underlined. The modified Cys residue is shown in blue color. (B) Mass spectrum from the LC-MS/MS analysis of the tryptic peptide containing Cys206 of CYP20-3. Predicted ion types and the detected ions are shown in Table S12. A Peptidyl-prolyl cis-trans isomerase CYP20-3 Protein sequence coverage: 61% Peptide score: 75.6 MASSSSMQMV HTSRSIAQIG FGVKSQLVSA NRTTQSVCFG ARSSGIALSS 50 RLHYASPIKQ FSGVYATTKH QRTACVKSMA AEEEEVIEPQ AKVTNKVYFD 100 VEIGGEVAGR IVMGLFGEVV PKTVENFRAL CTGEKKYGYK GSSFHRIIKD 150 FMIQGGDFTE GNGTGGISIY GAKFEDENFT LKHTGPGILS MANAGPNTNG 200 SQFFICTVKT SWLDNKHVVF GQVIEGMKLV RTLESQETRA FDVPKKGCRI 250 YACGELPLDA 260 y4 181H T G P G I L S M A N A G P N T N G S Q F F I C[CN] T V K209 y3 y4 y7 y9 y11 y15 b2b5b6b7b8b9b10 b12 b13 b15 b16 B Ion intensity Google Scholar: Author Only Title Only Author and Title Gotor, C., Laureano-Marín, A.M., Arenas-Alfonseca, L., Moreno, I., Aroca, Á., García, I., Romero, L.C. (2017). Advances in Plant Sulfur Metabolism and Signaling. In Progress in Botany Vol. 78, F M Cánovas, U Lüttge,R Matyssek, eds (Cham: Springer International Publishing), pp. 45-66. 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