This is the accepted version of the article: Rodríguez-Concepción, Manuel; D’Andrea, Lucio; Pulido, Pablo. «Control of plastidial metabolism by the Clp protease complex». Journal of experimental botany, Vol. 70, Issue 7 (March 2019), p. 2049-2058. DOI 10.1093/jxb/ery441 This version is avaible at https://ddd.uab.cat/record/205852 under the terms of the license Rodriguez-Concepcion et al. 1 Control of plastidial metabolism by the Clp protease complex 2 3 Manuel RODRIGUEZ-CONCEPCION*,1, Lucio D’ANDREA1,2, Pablo PULIDO1,3 4 5 6 1, Centre for Research in Agricultural Genomics (CRAG) CSIC-IRTA-UAB-UB, 08193 7 Barcelona, Spain. 8 2, Current address: Max-Planck-Institut für Molekulare Pflanzenphysiologie, D-14476 9 Potsdam-Golm, Germany. 10 3, Current address: Department of Plant Sciences, University of Oxford, Oxford OX1 3RB, 11 UK. 12 13 * Corresponding author 14 email, manuel.rodriguez@cragenomica.es 15 16 Date of submission: 07-september-2018 17 Number of Tables: 1 18 Number of Figures: 1 19 Word Count: 5177 (start of introduction to end of acknowledgements) 1 Rodriguez-Concepcion et al. 20 Abstract 21 Plant metabolism is strongly dependent on plastids. Besides hosting the 22 photosynthetic machinery, these endosymbiotic organelles synthesize starch, fatty 23 acids, amino acids, nucleotides, tetrapyrroles, and isoprenoids. Virtually all 24 enzymes involved in plastid-localized metabolic pathways are encoded by the 25 nuclear genome and imported into plastids. Once there, protein quality control 26 systems ensure proper folding of the mature forms and remove irreversibly 27 damaged proteins. The Clp protease is the main machinery for protein degradation 28 in the plastid stroma. Recent work has unveiled an increasing number of client 29 proteins of this proteolytic complex in plants. Notably, a substantial proportion of 30 these substrates are required for normal chloroplast metabolism, including 31 enzymes involved in the production of essential tetrapyrroles and isoprenoids such 32 as chlorophylls and carotenoids. The Clp protease complex acts in coordination 33 with nuclear-encoded plastidial chaperones for the control of both enzyme levels 34 and proper folding (i.e. activity). This communication involves a retrograde 35 signaling pathway, similarly to the unfolded protein response previously 36 characterized in mitochondria and endoplasmic reticulum. Coordinated Clp 37 protease and chaperone activities appear to further influence other plastid 38 processes, such as the differentiation of chloroplasts into carotenoid-accumulating 39 chromoplasts during fruit ripening. 40 41 Keywords: carotenoid, chaperone, chlorophyll, chloroplast, Clp protease, 42 chromoplasts, isoprenoid, proteostasis, tetrapyrrole, unfolded protein response 2 Rodriguez-Concepcion et al. 43 Introduction 44 Plastids are central metabolic factories for plant cells. Besides being the site of 45 carbon, nitrogen, and sulfur assimilation, they synthesize starch, fatty acids, amino 46 acids, nucleotides (purine and pyrimidine bases), tetrapyrroles (including heme 47 and chlorophyll), and isoprenoids (such as carotenoids, and the side chain of 48 chlorophylls). In fact, their classification into different plastid types largely depends 49 on the metabolites that they store: chloroplasts contain chlorophylls, chromoplasts 50 accumulate carotenoids, amyloplasts are starch-rich plastids, and elaioplasts are 51 specialized in storing lipids (Jarvis and Lopez-Juez, 2013; Rolland et al., 2012; 52 Sakamoto et al., 2008). 53 Similar to mitochondria, plastids are endosymbiotic organelles that retain their own 54 genome (plastome). The Arabidopsis thaliana plastome contains 45 RNA-coding 55 genes and 87 protein-coding genes, but only two of them are involved in processes 56 other than plastome gene expression or photosynthesis: accD encodes the 57 carboxytransferase beta subunit of the acetyl-CoA carboxylase complex (which 58 catalyzes the first committed step in fatty acid biosynthesis) and clpP1 encodes a 59 proteolytic subunit of the stromal Clp (Caseinolytic protease) complex (Sato et al., 60 1999). Therefore, virtually all proteins involved in plastidial metabolism (including 61 enzymes) are encoded by the nuclear genome and imported into plastids. Most of 62 these plastid-targeted proteins bear an N-terminal targeting signal called plastid 63 transit peptide. They are synthesized in the cytosol and transported into plastids in 64 an energy-dependent process mediated by molecular machines (translocons) 65 located in the outer and inner envelope membranes (Sakamoto et al., 2008). At 66 the inner membrane their transit peptide is cleaved and the mature forms of the 67 imported proteins are properly folded, assembled, and sorted to their particular 68 subplastidial destination by a separate targeting process. Inside plastids, protein 69 quality control (PQC) systems formed by chaperones and proteases promote 70 correct protein folding and remove irreversibly damaged proteins. To ensure an 71 appropriate supply of functional proteins, retrograde (plastid-to-nucleus) signaling 72 pathways adjust nuclear gene expression to particular plastid needs, e.g. in 73 response to environmental challenges (Chan et al., 2016; Grimm et al., 2014). 74 Regulated proteolysis additionally adjust protein levels and hence activity to the 3 Rodriguez-Concepcion et al. 75 specific requirements of individual plastids (Nishimura et al., 2016, 2017; 76 Sakamoto, 2006). 77 78 The multimeric Clp complex is the main stromal protease 79 Among the several proteases found in plant plastids, the serine-type Clp protease 80 complex is the main machinery for processive degradation of proteins in the plastid 81 stroma (Nishimura et al., 2016; Nishimura and van Wijk, 2015). One of the roles of 82 this housekeeping protease appears to be the removal of proteins that do not fold 83 properly or lose their native conformation and become misfolded in the stroma 84 (Nishimura and van Wijk, 2015; Zybailov et al., 2009). Because misfolded proteins 85 tend to aggregate, the Clp protease complex might also prevent the formation of 86 potentially toxic protein aggregates (Llamas et al., 2017; Nishimura and van Wijk, 87 2015; Zybailov et al., 2009). Additional PQC roles suggested for the Clp protease 88 include the survey of proteins translocated into plastids, clearing proteins that 89 remain unprocessed or misfolded after import but also removing proteins that 90 aggregate and become stuck in the import channel (Flores-Perez et al., 2016; 91 Huang et al., 2016; Sjogren et al., 2014). 92 Similar to other prokaryotic-type ATP-dependent plastidial proteases (such as 93 FtsH and Lon), the Clp complex contains two differentiated domains: a barrel- 94 shaped proteolytic core with the catalytic center positioned inside the chamber to 95 prevent unspecific protein degradation and a chaperone ring that delivers the 96 protein substrates. But unlike FtsH and Lon, in the Clp protease these two 97 characteristic domains are separated into individual multiprotein subcomplexes 98 (Figure 1). The subunit composition of the Clp protease is similar in all plastid types 99 (D'Andrea et al., 2018; Peltier et al., 2004). The proteolytic core (ClpRP) is formed 100 by two asymmetric rings (P and R) and stabilized by plant-specific ClpT1 and 101 ClpT2 proteins (Kim et al., 2015; Sjogren and Clarke, 2011). The P ring is formed 102 by a 1:2:3:1 ratio of proteolytically active subunits ClpP3, ClpP4, ClpP5 and ClpP6. 103 The R ring contains the plastome-encoded ClpP1 subunit but also non-catalytic 104 ClpR1, ClpR2, ClpR3 and ClpR4 proteins in a 3:1:1:1:1 ratio (Nishimura et al., 105 2016; Nishimura and van Wijk, 2015; Olinares et al., 2011; Peltier et al., 2004). 106 Proteins entering the catalytic ClpRP barrel are assumed to be degraded into 4 Rodriguez-Concepcion et al. 107 peptide fragments (5-10 residues) that are ejected from the core cavity through 108 lateral pores (Peltier et al., 2004). 109 Substrate access to the proteolytic core is controlled by a hexameric ring 110 presumably formed by homooligomers of the Hsp100-type chaperones ClpC1, 111 ClpC2, and ClpD (hence referred to it as ClpCD). Mutants defective in ClpC1 are 112 pale-green but knock-out lines of ClpC2 or ClpD show a wild-type phenotype, 113 supporting a major housekeeping role for the former. These chaperones 114 (particularly ClpC1) can directly recognize substrates and unfold them for 115 continuous translocation into the proteolytic chamber in an ATP-dependent 116 manner (Nishimura et al., 2016; Nishimura and van Wijk, 2015; Peltier et al., 2004). 117 In other cases, substrates are recognized by adaptors and then delivered to the 118 chaperones (Figure 1). Several pathways involving such adaptors are known. One 119 of them involves the recognition of client proteins by a binary adaptor formed by 120 ClpF and ClpS1 (Nishimura et al., 2015). The current model proposes that ClpF, 121 ClpS1, or a ClpF-ClpS1 complex recognizes the protein client and forms a ClpF- 122 ClpS1-substrate complex that then binds to the Hsp100 chaperone ring of the Clp 123 protease complex. A second pathway involves another chaperone type, Hsp70, 124 which can interact with ClpC1 to synergistically promote the unfolding process (Shi 125 and Theg, 2010; Su and Li, 2010). Hsp70 chaperones use J-proteins as adaptors 126 that guide them to protein clients and confer substrate specificity. After recognition 127 of specific proteins by particular J-protein adaptors, the substrates are delivered to 128 Hsp70, which then interacts with ClpC1 for unfolding and subsequent ClpRP- 129 mediated degradation (Pulido et al., 2016; Pulido et al., 2013). The possibility of 130 additional ways of targeting proteins to ClpC1, ClpC2, and ClpD chaperones 131 remains open (Pulido et al., 2017). It is generally assumed that specific signals for 132 degradation (named degrons) might be exposed in protein substrates that need to 133 be removed. The observation that the N-terminus of proteins is a major 134 determinant of protein turnover in prokaryotic and eukaryotic organisms led to 135 propose the N-end rule, which states that particular N-terminal residues act as 136 triggers for whole protein degradation. However, the identification of degrons and 137 the relevance of the N-end rule for the recognition of Clp protease targets by the 138 ClpCD ring chaperones or their adaptors (J-proteins/Hsp70, ClpF/ClpS1) remains 139 an open question (Rowland et al., 2015). 5 Rodriguez-Concepcion et al. 140 141 Several metabolic enzymes are clients of the Clp protease 142 Despite the central relevance of the Clp protease in PQC and plastidial 143 proteostasis, only a few proteins have been conclusively shown to be degraded by 144 this proteolytic complex. As true Clp protease clients, they (a) show increased 145 protein levels (but not transcripts) in mutants defective in Clp protease activity, (b) 146 interact with chaperones of the ClpCD ring or their adaptors, and (c) are degraded 147 at a slower rate in mutants of the chaperone or/and catalytic domains of the Clp 148 complex. Interestingly, most of the proteins experimentally demonstrated to meet 149 these criteria have a role in plastidial metabolism. Besides the thylakoid membrane 150 copper transporter PAA2/HMA8, a protein with stromal domains that is degraded 151 by the Clp protease by mechanisms that are not yet fully understood (Tapken et 152 al., 2015), four enzymes have been found to be Clp substrates in Arabidopsis 153 (Figure 1). The identified enzymes and the pathways in which they participate are 154 described below. We also discuss other plastidial enzymes that accumulate upon 155 interference with Clp protease activity but are not yet confirmed to be direct clients 156 of this proteolytic complex. 157 158 Glutamyl-tRNA reductase and tetrapyrrole biosynthesis. 159 The enzyme glutamyl-tRNA (GluT) reductase (GluTR) catalyzes an early step in 160 the tetrapyrrole biosynthesis pathway (Figure 1). GluT, produced after ligation of 161 Glu to tRNA(Glu), is reduced to Glu-1-semialdehyde by GluTR and then 162 transaminated to 5-aminolevulinic acid (ALA), the first committed precursor of the 163 pathway. Tetrapyrrole products are central for chloroplast metabolism as they are 164 required for photosynthesis (chlorophylls), nitrogen and sulfur assimilation 165 (siroheme), redox reactions (heme), photoperception (phytochromobilin), and 166 even retrograde signaling (Chan et al., 2016; Grimm et al., 2014). Tetrapyrrole 167 biosynthesis is tightly regulated to balance the production of chlorophylls and heme 168 and hence prevent the accumulation of metabolic intermediates that can potentially 169 cause severe photooxidative damage. Thus, chlorophyll biosynthesis needs to be 170 downregulated in darkness to prevent excessive accumulation of 171 protochlorophyllide in the dark (Meskauskiene et al., 2001). 6 Rodriguez-Concepcion et al. 172 As a key regulatory enzyme of the tetrapyrrole pathway, GluTR is controlled at 173 multiple levels. Among them, interaction of the N-terminal region of GluTR with the 174 GluTR binding protein (GBP) results in association of part of the stromal enzyme 175 to the thylakoid membrane (Czarnecki et al., 2011). It is proposed that this 176 mechanism allows two separate pools of GluTR for the separate production of ALA 177 for heme in the thylakoid membrane and for chlorophylls in the stroma. The N- 178 terminal domain of GluTR harbors aggregation-prone motifs, but interaction with 179 the plastidial chaperone SRP43 efficiently prevents aggregation, thus enhancing 180 the stability of the protein and promoting its localization to the thylakoid membrane 181 (Wang et al., 2018). Furthermore, this N-terminal region can also interact with the 182 ClpF and ClpS1 adaptors and the ClpC1 chaperone for Clp protease-mediated 183 degradation of the enzyme (Apitz et al., 2016; Nishimura et al., 2015; Nishimura et 184 al., 2013). A truncated GluTR protein lacking the N-terminal region remains in the 185 stroma and is more stable during prolonged darkness, causing an enhanced 186 accumulation of protochlorophyllide and a necrotic phenotype upon reillumination 187 (Apitz et al., 2016). Consistent with the role of the N-terminal domain as a degron, 188 the rate of GluTR proteolysis in the dark is increased in the absence of GBP (i.e. 189 when the degron is exposed) but decreased in mutants impaired in ClpC1 or the 190 proteolytic core subunit ClpR2 (Apitz et al., 2016). While GluTR accumulates at 191 higher levels in ClpC1 and ClpS1 mutants, the degradation rate of the enzyme in 192 the dark appears not to be altered when ClpS1 is missing. It is therefore concluded 193 that ClpC1 might have a more relevant role than ClpS1 (or ClpF) to recognize and 194 direct GluTR into the Clp proteolytic core for degradation (Apitz et al., 2016). A 195 proposed model (Wang et al., 2018) suggests that the interaction of SRP43 with 196 unfolded GluTR after plastid import might ensure proper enzyme folding and 197 prevent aggregation. Folded GluTR could then interact with GBP for thylakoid 198 membrane targeting. It is likely that competition for the N-terminal domain between 199 GluTR interactors (SPR43 and GBP) and Clp adaptors or/and chaperones, but 200 also structural changes triggered by metabolite binding, misfolding or aggregation, 201 might play a role in determining the degradation rate of this important enzyme. 202 Strikingly, the N-terminus is relevant for the Clp-mediated degradation of GluTR 203 not only in plants but also in bacteria (Wang et al., 1999), suggesting a remarkable 204 evolutionary conservation of the mechanisms controlling GluTR turnover. 7 Rodriguez-Concepcion et al. 205 206 Chlorophyllide a oxygenase and chlorophyll biosynthesis. 207 The final steps of chlorophyll biosynthesis (Figure 1) involve the incorporation of 208 molecular oxygen into the C7-methyl group of the tetrapyrrole ring (porphyrin) of 209 chlorophyllide a to produce chlorophyllide b followed by the esterification of 210 isoprenoid-derived phytyl diphosphate to both chlorophyllides to form chlorophylls 211 a and b, respectively (Kim et al., 2013b). Chlorophyll a occurs in antenna 212 complexes and reaction centers, whereas chlorophyll b is absent from reaction 213 centers. Chlorophyll b levels influence the antenna size of photosystem II and are 214 controlled by the stability of chlorophyllide a oxygenase (CAO), the enzyme that 215 catalyzes the conversion of chlorophyllide a to b (Sakuraba et al., 2007; Yamasato 216 et al., 2005). Degradation of CAO when chlorophyll b is present requires the N- 217 terminal domain (or A domain) of the enzyme (Yamasato et al., 2005) and is 218 impaired in ClpC1-defective mutants (Nakagawara et al., 2007). The current model 219 is that chlorophyll b may modulate the structure of the A domain either by direct 220 interaction or via other factors. The induced change of conformation would then 221 expose a degron for ClpC1-mediated unfolding and eventual ClpRP-mediated 222 degradation of the whole enzyme (Sakuraba et al., 2009). CAO localizes in 223 thylakoid and envelope membranes but lacks obvious membrane‐spanning 224 domain (Eggink et al., 2004; Yamasato et al., 2005). It is therefore possible that 225 interaction of the N-terminal domain of CAO with ClpC1 (or an adaptor) occurs at 226 the periphery of these plastidial membranes. 227 228 Deoxyxylulose 5-phosphate synthase and isoprenoid biosynthesis. 229 The isoprenoid phytol chain of chlorophylls is synthesized from precursors derived 230 from the stromal methylerythritol 4-phosphate (MEP) pathway. In fact, a strict 231 coordination between the tetrapyrrole and the MEP pathways is essential to 232 prevent the formation of oxidative metabolic intermediates (Kim et al., 2013b). 233 MEP-derived prenyl diphosphates isopentenyl diphosphate (IPP) and dimethylallyl 234 diphosphate (DMAPP) are also used for the production of other plastidial 235 isoprenoids involved in photosynthesis (tocopherols and carotenoids as 236 photoprotectants, and prenylquinones such as plastoquinone and phylloquinones 8 Rodriguez-Concepcion et al. 237 for electron transport) and signaling (including several types of retrograde signals, 238 hormones, and volatiles for environmental interactions) (Rodriguez-Concepcion 239 and Boronat, 2015). The control of protein turnover has a major impact on MEP 240 pathway enzymes including deoxyxylulose 5-phosphate (DXP) synthase (DXS), 241 DXP reductoisomerase (DXR), hydroxymethylbutenyl 4-diphosphate (HMBPP) 242 synthase (HDS), and HMBPP reductase (HDR) (Figure 1). All these enzymes 243 accumulate at higher levels in Arabidopsis plants with decreased Clp proteolytic 244 activity such as mutants defective in either ClpC1 or ClpRP subunits (Table 1) 245 (Flores-Pérez et al., 2008; Kim et al., 2013a; Kim et al., 2009; Nishimura et al., 246 2013; Rodriguez-Villalon et al., 2009; Rudella et al., 2006; Sauret-Güeto et al., 247 2006; Welsch et al., 2018; Zybailov et al., 2009). However, only DXS has been 248 experimentally shown to be a direct Clp substrate based on the three criteria 249 described above (Figure 1). DXS can either directly interact with ClpC1 or be 250 delivered to this chaperone via J20, a J-protein adaptor of Hsp70 (Pulido et al., 251 2016), as detailed below. DXR is more stable and does not rely on J20 for 252 degradation (Perello et al., 2016; Pulido et al., 2016). Instead, removal of excess 253 DXR takes place by packaging the enzyme in vesicles formed by engulfing stromal 254 fractions that likely retain Clp protease activity (Perello et al., 2016). The 255 accumulation of MEP pathway enzymes does not change in mutants defective in 256 ClpF or ClpS1 adaptors with only one exception: HDS (Nishimura et al., 2015; 257 Pulido et al., 2016). However, HDS levels do not increase but decrease in the 258 mutants (Table 1) (Nishimura et al., 2015), i.e. the opposite that it would be 259 expected if the ClpF/ClpS1 pathway delivers this enzyme to the Clp complex. 260 Beyond Arabidopsis, work in other systems has shown that the accumulation of 261 DXS and other MEP pathway enzymes is directly or indirectly controlled by the Clp 262 protease in tobacco (Nicotiana tabacum) chloroplasts (Moreno et al., 2018), 263 tomato (Solanum lycopersicum) chromoplasts (D'Andrea et al., 2018), the plastid- 264 like apicoplasts of the malaria parasite Plasmodium falciparum (Florentin et al., 265 2017) and even bacterial cells (Ninnis et al., 2009). 266 Consistent with the major contribution of DXS to the control of the MEP pathway 267 flux (Pokhilko et al., 2015; Wright et al., 2014), its activity is regulated by several 268 post-translational mechanisms (Rodriguez-Concepcion and Boronat, 2015), 269 including Clp-mediated degradation (Flores-Perez et al., 2008; Llamas et al., 2017; 9 Rodriguez-Concepcion et al. 270 Perello et al., 2016; Pulido et al., 2016; Welsch et al., 2018). A shortage of MEP 271 pathway products upregulates DXS enzymatic activity and promotes the 272 accumulation of higher protein levels (Ghirardo et al., 2014; Han et al., 2013; 273 Pokhilko et al., 2015). It is possible that binding of IPP or DMAPP to DXS causes 274 a conformational change that inactivates the enzyme and renders it more 275 susceptible to degradation (Pokhilko et al., 2015). Stress episodes can also disrupt 276 the native structure of DXS, resulting in protein misfolding, and eventually 277 aggregation in chloroplasts. The high aggregation propensity of DXS was also 278 observed in cyanobacteria, the evolutionary ancestors of chloroplasts (Kudoh et 279 al., 2017). Work in Arabidopsis has unveiled a PQC mechanism to deal with the 280 unwanted accumulation of inactive DXS proteins (Figure 1). The J-protein adaptor 281 J20 specifically binds misfolded forms of DXS and delivers them to the Hsp70 282 chaperone for either proper folding or degradation (Pulido et al., 2013). The fate of 283 Hsp70-bound enzyme appears to depend on the relative abundance of Hsp100 284 chaperones that synergistically contribute to the unfolding process. Under normal 285 growth conditions, inactive Hsp70-bound DXS proteins are delivered to ClpC1 and 286 unfolded prior entrance into the ClpRP proteolytic chamber to be degraded. 287 Environmental episodes causing protein folding stress (e.g. heat) do not cause 288 changes in the levels of ClpC1 but induce the accumulation of ClpB3, another 289 plastidial member of the Hsp100 family. Unlike ClpC1 and the other Hsp100 290 chaperones associated to the ClpCD complex, ClpB3 lacks the IG(F/L) tripeptide 291 (also named P-loop motif) required for interaction with the ClpRP core subunits but 292 harbors a domain that allows efficient interaction with Hsp70 chaperones (Flores- 293 Perez et al., 2016; Kim et al., 2001; Peltier et al., 2004; Pulido et al., 2016). 294 Therefore, stress episodes activate the ClpB3-dependent pathway to unfold 295 Hsp70-bound DXS into the stroma for spontaneous refolding and reactivation 296 (Pulido et al., 2016). This model implies that although the refolding strategy (via 297 ClpB3) is likely favored, Clp-mediated degradation (via ClpC1) is the housekeeping 298 pathway to remove inactive DXS. Most likely, both share the same goal, i.e. to 299 prevent protein aggregation in the plastid and maintain the enzymes in a 300 catalytically active form. The observation that double mutants defective in ClpB3 301 and ClpR2 were unable to survive under autotrophic conditions (Zybailov et al., 302 2009) illustrates the key relevance of the seemingly antagonistic refolding and 303 degradation pathways for plant life. The common regulation of both the MEP and 10 Rodriguez-Concepcion et al. 304 the tetrapyrrole pathways by the Clp protease (Figure 1) likely contributes to finely 305 balance the supply of metabolic precursors for the production of chlorophylls (Kim 306 et al., 2013b). 307 308 Phytoene synthase and carotenoid biosynthesis. 309 The activity of the MEP pathway enzymes generates prenyl diphosphates used by 310 the enzyme phytoene synthase (PSY) to produce phytoene, the first committed 311 intermediate of the carotenoid pathway. Carotenoids are essential for 312 photoprotection of the photosynthetic apparatus against excess light, act as 313 precursors of hormones and retrograde signals, and function as natural pigments 314 in non-photosynthetic organs such as flowers and fruits (Rodriguez-Concepcion et 315 al., 2018). Yeast two-hybrid experiments followed by bimolecular fluorescence 316 complementation assays in plant cells showed that PSY, the main rate-determining 317 enzyme of the pathway (Fraser et al., 2002), can physically interact with ClpS1 and 318 several Clp protease subunits, including ClpC1 and ClpD (Welsch et al., 2018). 319 However, only the absence of ClpC1 results in increased enzyme levels as a 320 consequence of reduced PSY turnover rate, suggesting a dispensable contribution 321 of ClpS1 or ClpD in the delivery of the enzyme to ClpRP-mediated degradation 322 (Figure 1). As expected, PSY enzyme levels are also increased in mutants 323 defective in individual subunits of the ClpRP proteolytic core such as ClpP4 and 324 ClpR1. PSY stability further increases in the presence of ORANGE (OR), a J-like- 325 protein with chaperone activity that also promotes PSY enzymatic activity (Park et 326 al., 2016; Pulido and Leister, 2018; Welsch et al., 2018; Zhou et al., 2015). OR 327 might play a similar role as ClpB3 in the case of DXS, i.e. promote correct PSY 328 folding (and hence enzymatic activity) and prevent its misfolding and eventual 329 aggregation by the Clp protease (Figure 1). 330 Similar to PSY, other carotenoid biosynthetic enzymes overaccumulate in mutants 331 defective in ClpC1 or ClpRP core subunits (Figure 1) (Welsch et al., 2018). While 332 further experiments will be required to conclude whether these enzymes are true 333 Clp protease targets, the available evidence suggests that Clp-dependent 334 mechanisms contribute to coordinate the MEP pathway and the carotenoid 335 pathway by influencing the accumulation of multiple enzymes of both pathways. 336 Besides possible secondary effects eventually impacting enzyme levels, this 11 Rodriguez-Concepcion et al. 337 control mechanism involves the direct degradation of some enzymes, including 338 those catalyzing the main rate-determining steps (DXS and PSY). Because the Clp 339 protease also regulates the production of chlorophylls via direct control of GluTR 340 and CAO turnover (Figure 1), this proteolytic complex might further coordinate 341 chlorophyll and carotenoid biosynthesis for an efficient photosynthetic function in 342 chloroplasts. Recent results further suggest that the Clp protease might ensure a 343 proper supply of MEP-derived precursors for carotenoid biosynthesis in different 344 plastid types. Downregulation of Clp protease activity during tomato ripening led to 345 increased levels of both DXS and PSY enzymes in the carotenoid-accumulating 346 chromoplasts of ripe fruit (D'Andrea et al., 2018). Importantly, Clp-defective fruits 347 developed aberrant chromoplasts with disorganized membranes and displayed an 348 orange color (instead of red) because they accumulated enhanced levels of the 349 pro-vitamin A carotenoid β-carotene (D'Andrea et al., 2018). These data indicate 350 that Clp protease activity controls not only carotenoid biosynthesis (via DXS and 351 PSY stability) but also storage (via plastidial ultrastructure). It is conceivable that 352 during normal fruit ripening Clp protease function might facilitate the turnover and 353 removal of many other enzymes and structural proteins that lose their functionality 354 or that are not further required as chloroplasts differentiate into chromoplasts. 355 356 Other enzymes and metabolic pathways. 357 Genetic interference with the accumulation of Clp subunits perturbs the 358 stoichiometry in the complex and negatively impacts its protein degradation 359 activity. Based on the premise that decreased Clp proteolytic activity should lead 360 to increased accumulation of its protein targets, quantitative proteomic approaches 361 have been employed for the identification of potential Clp protease clients. Initially, 362 the steady state proteome of wild-type Arabidopsis plants was compared with that 363 of mutants defective in particular subunits of the complex (Kim et al., 2013a; Kim 364 et al., 2009; Rudella et al., 2006; Sjogren et al., 2006; Stanne et al., 2009; Zybailov 365 et al., 2009). While components of metabolic pathways impacting the production 366 of starch, fatty acids, amino acids, isoprenoids, and thiamine were found to be 367 upregulated in the mutants, this strategy per se could not distinguish between 368 primary and downstream effects (e.g. loss of photosynthetic activity and 369 downregulation of Calvin cycle enzymes) associated with the reduction of Clp 12 Rodriguez-Concepcion et al. 370 protease activity (Nishimura and van Wijk, 2015). Further biochemical approaches 371 using mutants defective in the adaptors ClpF and ClpS1 went a step forward and 372 identified GluTR as a target of the complex (Nishimura et al., 2015; Nishimura et 373 al., 2013). Other potential clients identified in these works are the enzymes 374 deoxyarabinoheptulosonate 7-phosphate synthase (DHS) and chorismate 375 synthase (CS), which catalyze the first and last reaction of the shikimate pathway, 376 respectively. While DHS isoforms were consistently found to be upregulated in 377 mutants defective in ClpC1 and ClpRP core subunits, the results for CS are much 378 less clear (Table 1). The product of the shikimate pathway, chorismate, is 379 converted into aromatic amino acids such as Phe or Tyr, which are also precursors 380 for the biosynthesis of phenylpropanoids (e.g. lignins, flavonols, and anthocyanins) 381 and isoprenoids such as tocopherols, plastoquinone, and phylloquinones (Figure 382 1). 383 More recently, induced downregulation of Clp complex subunits in tobacco led to 384 propose additional enzymes as potential Clp protease clients based on their 385 overaccumulation at early time points after triggering repression of Clp proteolytic 386 activity (Moreno et al., 2018). Among them, two enzymes involved in thiamine 387 biosynthesis appear as good candidates: THI1 and THIC (Figure 1). These two 388 enzymes catalyze the first steps in the biosynthesis of the two moieties required 389 for the production of thiamine, thiazole (THI1) and pyrimidine (THIC) (Goyer, 390 2010). Supporting the conclusion that they might be true Clp protease targets, the 391 Arabidopsis THI1 and THIC proteins are accumulated at higher levels in mutants 392 defective in ClpC1 or ClpR2 (Table 1) (Nishimura et al., 2013; Zybailov et al., 393 2009). In the case of THI1, ClpS1 mutants contained increased protein levels in 394 one study (Nishimura et al., 2013) but decreased in another (Nishimura et al., 395 2015), preventing to reach a conclusion on the possible role of ClpS1 in delivering 396 this enzyme to degradation. In any case, it is interesting to note that thiamine 397 diphosphate (vitamin B1) serves as a cofactor for many enzymes (including DXS; 398 Figure 1) and hence represents an important node in the control of the plant cell 399 metabolism. 400 401 Plastidial proteostasis requires communication with the nucleus 13 Rodriguez-Concepcion et al. 402 Highly reproducible patterns deduced from quantitative proteomics analyses of 403 Clp-defective mutants include a strong reduction of photosynthetic capacity (in part 404 due to loss of thylakoid membrane homeostasis) and a systematic upregulation of 405 all stromal chaperone systems, with most consistent increases typically observed 406 for the stromal Hsp100 chaperone ClpB3 (Table 1). It was proposed that 407 accumulation of ClpB3 and stromal protein folding machineries in the mutants was 408 a response to deal with protein aggregation likely occurring when Clp protease 409 activity was defective (Nishimura and van Wijk, 2015; Zybailov et al., 2009). By 410 downregulating the expression of the plastome-encoded ClpP1 subunit of the 411 ClpRP core in the unicellular alga Chlamydomonas reinhardtii, it was observed that 412 these chaperones and small heat shock proteins such as Hsp21 increased at both 413 transcript and protein levels (Ramundo et al., 2014). A role for retrograde signaling 414 was later suggested in plants to increase the supply of nuclear-encoded plastid- 415 targeted chaperones when loss of Clp protease activity caused protein folding 416 stress in the plastid (Nishimura and van Wijk, 2015). More recently, the existence 417 of a mechanism upregulating the expression of nuclear genes encoding plastid- 418 targeted chaperones such as Hsp21, Hsp70, and ClpB3 was demonstrated in 419 Arabidopsis (Llamas et al., 2017). The mechanism was designated chloroplast 420 unfolded protein response (cpUPR), based on its conceptual similarity to those 421 previously reported in mitochondria and endoplasmic reticulum (Hwang and Qi, 422 2018; Lin and Haynes, 2016). Although the term “unfolded” might be misleading 423 and it would be most appropriate to name it protein folding stress response, we will 424 keep using the term cpUPR to be consistent with the literature. 425 Studies using DXS as a model showed that plastid proteostasis involves a dynamic 426 balance between degradation and repair of structurally compromised proteins. In 427 non-stressed conditions, the Clp protease (via ClpC1) removes DXS proteins that 428 become non-functional (e.g. when losing their native conformation). However, 429 protein folding stress can occur when Clp protease activity becomes compromised 430 and cannot deal with the accumulation of misfolded proteins. For example, 431 malfunctioning of the plastome gene expression system can lead to altered levels 432 of the catalytic ClpP1 subunit (Flores-Pérez et al., 2008), causing reductions in Clp 433 protease activity (because of distortions in the stoichiometry of the complex) that 434 eventually lead to protein aggregation (Llamas et al., 2017). Protein folding stress 14 Rodriguez-Concepcion et al. 435 in the plastid somehow sends an unknown retrograde signal to upregulate the 436 expression of the nuclear gene encoding the transcription factor HsfA2 (Llamas et 437 al., 2017). HsfA2 directly binds the promoters of genes encoding ClpB3 and other 438 plastidial chaperones, inducing their expression. As a result, more of these 439 chaperones are made and sent to plastids to alleviate protein folding stress in this 440 organelle. Higher levels of plastidial ClpB3 then promote the disaggregation of 441 DXS, i.e. the recovery pathway. It is expected that upregulating the supply of 442 plastidial chaperones would also increase the global folding capacity of the plastid, 443 which should contribute to effectively remove protein aggregates, promote correct 444 folding, and restore proteostasis. 445 Consistent with the conclusion that the basic PQC mechanisms are conserved 446 among different plastid types, the downregulation of Clp protease activity during 447 tomato fruit ripening also resulted in increased accumulation of chaperones in the 448 chromoplasts of ripe fruit (D'Andrea et al., 2018). Specifically, genes encoding 449 tomato homologues of ClpB3 and OR were upregulated in Clp-defective fruit, 450 leading to increased levels of the corresponding proteins and subsequent 451 protection of DXS and PSY enzymes, respectively, against degradation. 452 Interestingly, OR chaperones not only promote PSY stability and enzymatic activity 453 but also act as positive regulators of chromoplast differentiation and repressors of 454 carotenoid degradation (Chayut et al., 2017; Park et al., 2016; Zhou et al., 2015). 455 It is proposed that Clp protease-regulated expression of nuclear genes encoding 456 chaperones such as OR and ClpB3 might be a compensatory mechanism 457 triggered when Clp protease activity is compromised during fruit ripening 458 (D'Andrea et al., 2018). When Clp protease activity becomes limiting, the resulting 459 defective protein turnover might impair chromoplast differentiation. The 460 accumulation of non-functional proteins, however, would trigger a cpUPR to 461 eventually mitigate protein folding stress (e.g. via ClpB3) and restore chromoplast 462 development (e.g. via OR). The observation that, similar to that described in 463 Arabidopsis (Llamas et al., 2017), tomato HsfA2 regulates the expression of 464 plastidial chaperones to increase the protection against protein folding stress 465 (Fragkostefanakis et al., 2015), suggests that several components of the organelle 466 UPR might be conserved among different species. A better understanding of how 467 the Clp protease coordinates with chaperones such as ClpB3 and OR in different 15 Rodriguez-Concepcion et al. 468 plastid types should contribute to more rational engineering approaches for 469 improved production of carotenoids and other plastidial metabolites. 470 471 Open questions 472 While significant progress has been achieved in the last few years on the 473 identification of Clp protease protein substrates, we still have an incomplete map 474 of the plastidial pathways impacted by this major proteolytic machinery. In addition, 475 substrates involved in the storage of plastidial metabolites remain to be identified. 476 For example, Clp-defective mutants typically show increased levels of proteins 477 associated to plastoglobules, which are thylakoid-derived compartments that store 478 isoprenoids and other lipids (van Wijk and Kessler, 2017). However, it is unclear 479 whether this is a primary effect of reducing the turnover of Clp protein substrates 480 or a secondary consequence derived from interferences with thylakoid membrane 481 homeostasis or stress responses (Nishimura and van Wijk, 2015). Recent 482 approaches using inducible lines have been useful to distinguish early from late 483 accumulating proteins (Moreno et al., 2018; Ramundo et al., 2014). Clp-trapping 484 strategies similar to those used in bacteria (Flynn et al., 2003; Trentini et al., 2016) 485 were expected to also be useful to identify bona fide plant targets, despite the 486 challenges derived from the much higher complexity and essential nature of the 487 plastidial Clp protease. Recent experiments using in vivo tagged core complexes 488 containing inactive ClpP3 and ClpP5 variants in Arabidopsis could not identify new 489 Clp protease clients, proposedly because the bottleneck for degradation might not 490 be the catalytic activity of the ClpRP core but substrate recognition and unfolding 491 (Liao et al., 2018). In vivo substrate trapping through partial inactivation of ClpC1, 492 ClpC2 or ClpD might therefore provide an alternative, more efficient strategy to 493 identify protein clients. 494 To date, only ClpC1-dependent substrates have been described, suggesting that 495 the other chaperones of the ClpCD ring (ClpC2 and ClpD) are partially redundant 496 (Kovacheva et al., 2007; Nishimura et al., 2015). Strikingly, ClpD is found 497 exclusively in stroma, whereas ClpC1 and ClpC2 are also present in association 498 with envelope membranes for PQC of imported proteins (Sjogren et al., 2014). A 499 related question is how the substrates of the Clp protease are selected for 500 degradation, while preventing the removal of functional proteins and off-targets. 16 Rodriguez-Concepcion et al. 501 While the relevance of the N-end rule for chloroplast protein turnover remains 502 unclear (Rowland et al., 2015), it is likely that the first step in the degradation 503 pathway could be exposure of a degron after a conformational change caused by 504 metabolite binding (as in the case of CAO) or by misfolding (as in the case of 505 GluTR, DXS and PSY). Degrons could also be exposed after removal of proteins 506 bound to the motifs required for degradation (as in the case of GluTR). Then, the 507 degron could be recognized by the ClpF/ClpS1 tandem or the J-protein/Hsp70 508 system and delivered to ClpC1 or any other Hsp100 component of the Clp 509 chaperone ring for unfolding and ClpRP-mediated proteolysis. Rather than a 510 particular amino acid sequence, degrons might comprise a secondary structure 511 and/or a post-translational modification (phosphorylation, acetylation, etc), which 512 enormously complicate their identification. For instance, arginine phosphorylation 513 was found to direct substrates for degradation by the Clp protease in bacteria 514 (Trentini et al., 2016). Furthermore, it is likely that some enzymes could be 515 degraded by different proteases in the plastid (Nishimura et al., 2016). Considering 516 the protease network as a whole (Majsec et al., 2017) would also be required to 517 fully understand how plastidial metabolism is regulated inside plastids in response 518 to developmental and environmental cues. 519 Finally, the molecular pathway connecting Clp protease defects with enhanced 520 expression of genes for plastid-targeted chaperones remains unknown. Recent 521 results have shown that GUN1, a central regulator of plastid to nucleus 522 communication, is degraded by the Clp protease via ClpC1 (Wu et al., 2018). 523 Another protein involved in tetrapyrrole-dependent retrograde signaling, GUN5, is 524 also upregulated in Arabidopsis mutants defective in ClpC1 and Clp protease 525 activity (Table 1). A role for the Clp protease in controlling the turnover of 526 retrograde signaling mediators such as GUN1 and GUN5 is expected to represent 527 a powerful tool in the communication of plastidial protein folding stress to the 528 nucleus. However, neither GUN1 nor GUN5 appear to be required for the 529 upregulation of genes encoding plastidial chaperones such as Hsp21, Hsp70, and 530 ClpB3 that occurs soon after triggering protein aggregation in chloroplasts by 531 interfering with Clp protease activity (Llamas et al., 2017). Alternative good 532 candidates to participate in this process are the DNA-interacting factors WHIRLY 533 (pTAC1/WHY1 and pTAC11/WHY3) and pTAC12/HEMERA, which have been 17 Rodriguez-Concepcion et al. 534 located in plastidial transcriptionally active chromosomes (pTACs) or nucleoids but 535 also in the nucleus (Melonek et al., 2016). These proteins are more abundant in 536 Clp-defective plants (Table 1) (Moreno et al., 2018) and display the same size in 537 chloroplasts and nuclei, suggesting that their accumulation in plastids with a 538 compromised Clp proteolytic activity might cause their translocation to the nucleus 539 to regulate nuclear gene expression. Future work should contribute to confirm this 540 hypothesis and identify other dual-localized factors potentially involved in the 541 cpUPR-associated retrograde signaling pathway that allows plants to overcome 542 protein folding stress in the plastid. 543 544 ACKNOWLEDGEMENTS 545 We thank Juan Camilo Moreno (Max-Planck-Institut für Molekulare 546 Pflanzenphysiologie) for comments on the manuscript. Work in our lab is funded 547 by grants BIO2015-71703-REDT, BIO2017-90877-REDT and BIO2017-84041-P 548 from the Spanish Ministry of Economy and Competitiveness (MINECO), 549 2017SGR-710 from Generalitat de Catalunya, and CA15136 (COST action 550 EuroCaroten) from the European Commission H2020 program. 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Large scale comparative proteomics of a chloroplast Clp protease 812 mutant reveals folding stress, altered protein homeostasis, and feedback 813 regulation of metabolism. Molecular Cell Proteomics 8, 1789-1810. 814 815 24 Rodriguez-Concepcion et al. 816 TABLES 817 Table 1. Levels of plastidial proteins in Clp-defective Arabidopsis mutants. Zybailov Kim et Kim et Nishimura et al., et al., al., al., Nishimura et al., 2013 2015 2009 2009 2013 clps1 clpr4- clpp3- clpc1- clpr2- clpf1- Accession Name clpc1- clps1 clps1 1 1 1 1 1 1 At4g15560 DXS nd nd higher 22,88 nd nd nd nd At5g62790 DXR 3,53 2,06 2,05 1,73 0,78 1,09 1,63 1,06 At2g02500 MCT nd 3,48 nd nd nd 0,90 higher 1,06 MEP At2g26930 CMK nd 0,71 nd nd nd 0,70 nd 0,25 pathway At1g63970 MDS nd 1,61 nd nd nd 0,78 nd 0,81 At5g60600 HDS 5,10 3,54 2,05 2,58 0,71 0,63 9,25 0,50 At4g34350 HDR nd nd 3,38 higher 1,12 1,02 nd 0,50 At4g33510 DHS2 nd 3,30 higher higher nd 0,37 3,01 0,74 Shikimate At4g39980 DHS1 nd 2,65 nd nd nd nd 8,16 nd pathway At1g48850 CS nd nd 2,33 1,06 1,24 1,03 nd 1,27 Thiamine At5g54770 THI1 2,37 1,63 17,02 4,26 2,25 0,69 5,35 0,54 pathway At2g29630 THIC nd 0,42 6,98 2,41 0,57 0,68 5,01 0,74 At1g14410 WHY1 5,30 nd nd nd nd 1,76 2,67 nd Transcription At2g02740 WHY3 nd 7,37 nd higher nd 0,57 nd 0,51 factors At2g34640 HEMERA nd 1,55 nd nd nd nd nd nd At3g59400 GUN4 nd 0,90 nd nd nd 0,88 nd 1,01 Other At5g13630 GUN5 6,71 1,35 3,46 3,52 0,32 1,53 7,04 0,77 proteins At5g15450 ClpB3 4,46 4,84 3,39 2,62 1,62 0,60 3,67 0,55 818 819 Numbers indicate the ratio of protein levels in the indicated Arabidopsis mutant relative to 820 the wild-type (fold-change) 821 Colors indicate when protein levels in the mutant are statistically higher (blue) or lower 822 (red) compared to the wild-type 823 higher, proteins detected only or preferentially in the mutant 824 nd, not detected 825 826 25 Rodriguez-Concepcion et al. 827 FIGURE LEGENDS 828 829 830 Figure 1. Degradation of plastidial enzymes by the Clp protease complex. 831 Enzymes are indicated in green, adaptor systems are represented in purple, and 832 proteins that prevent degradation are shown in black. Green arrows represent 833 metabolic pathways. Solid black arrows mark confirmed interactions and dashed 834 black arrows indicate potential interactions. The structure of the Clp protease 835 complex and its components is also shown. 26