Protein Maturation and Proteolysis in Plant Plastids, Mitochondria, and Peroxisomes

PP66CH04-VanWijk ARI 30 March 2015 20:57

Klaas J. van Wijk

Department of Plant Biology, Cornell University, Ithaca, New York 14853; email: [email protected]

Annu. Rev. Plant Biol. 2015. 66:75–111 Keywords First published online as a Review in Advance on proteome remodeling, endosymbiotic organelle, degron, proteostasis, January 12, 2015 degradomics, autophagy The Annual Review of Plant Biology is online at plant.annualreviews.org Abstract This article’s doi: Plastids, mitochondria, and peroxisomes are key organelles with dynamic 10.1146/annurev-arplant-043014-115547 proteomes in photosynthetic eukaryotes. Their biogenesis and activity must Copyright c 2015 by Annual Reviews. be coordinated and require intraorganellar protein maturation, degradation, All rights reserved and recycling. The three organelles together are predicted to contain ∼200

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org presequence peptidases, proteases, aminopeptidases, and specific protease chaperones/adaptors, but the substrates and substrate selection mechanisms are poorly understood. Similarly, lifetime determinants of organellar proteins, such as N-end degrons and tagging systems, have not been identified, but the substrate recognition mechanisms likely share similarities between organelles. Novel degradomics tools for systematic analysis of protein lifetime and proteolysis could define such protease-substrate relationships,

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. degrons, and protein lifetime. Intraorganellar proteolysis is complemented by autophagy of whole organelles or selected organellar content, as well as by cytosolic protein ubiquitination and degradation by the proteasome. This review summarizes (putative) plant organellar protease functions and substrate-protease relationships. Examples illustrate key proteolytic events.

75 PP66CH04-VanWijk ARI 30 March 2015 20:57

Contents WHY DO ORGANELLES NEED PROTEOLYSIS? ...... 76 COMMONALITIES BETWEEN PLASTIDIAL, MITOCHONDRIAL, AND PEROXISOMAL PROTEOLYSIS AND ORGANELLE TURNOVER . . . 77 SCOPE OF THIS REVIEW, KEY QUESTIONS, AND NEW FRONTIERS IN ORGANELLAR PROTEOLYSIS...... 78 ORGANELLAR PROTEASE INVENTORY, CLASSIFICATION, ANDCHARACTERIZATION...... 78 Presequence Cleavage, Protein Maturation, Stabilization, and Recycling...... 83 ATP-Dependent Proteases of the AAA+ Family: LON, CLP, and FTSH ...... 85 TheATP-IndependentDEGProteaseFamily...... 90 Intramembrane Cleaving Proteases of the Rhomboid and S2P Families in the Organelles...... 92 Recent Discoveries and Functional Analysis of Other Organellar Proteases ...... 93 PROTEASE NETWORKS, PROTEOSTASIS, AND RETROGRADESIGNALING...... 93 SUBSTRATE SELECTION, PROTEIN LIFETIME DETERMINANTS, DEGRONS, AND POSTTRANSLATIONAL MODIFICATIONS ...... 95 TheN-EndRuleandN-Degrons...... 96 Posttranslational Modiﬁcations Can Affect Proteolysis ...... 96 INVOLVEMENT OF UBIQUITINATION AND THE PROTEASOME IN ORGANELLE BIOGENESIS AND DIFFERENTIATION ...... 97 ORGANELLE PROTEOME TURNOVER THROUGH AUTOPHAGY ANDTARGETEDVESICLETRANSPORT...... 98 Pexophagy...... 98 Mitophagy...... 98 Chlorophagy and Targeted Chloroplast Protein Removal by Rubisco-Containing Bodies and Senescence-Associated Vacuoles...... 99 ORGANELLAR PROTEOLYSIS IN CROP PLANTS, BIOTECHNOLOGY, ANDSYNTHETICBIOLOGY...... 99 DEGRADOMICS: SYSTEMS ANALYSIS OF PROTEOLYSIS ANDPROTEINLIFETIME...... 100 Identiﬁcation of Cleavage Sites and Degradation Products Through

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org Large-ScaleN-orC-TerminalLabelingandEnrichment...... 101 ComparativeProteomicsofProteaseMutants...... 101 ProteinLifetimeAnalysisUsingStableIsotopes...... 101

WHY DO ORGANELLES NEED PROTEOLYSIS?

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Plastids, mitochondria, and peroxisomes each have their own complement of proteins, referred to as their proteomes. Most of these proteins are encoded by nuclear genes, but plastids and mitochondria also contain a set of ∼85 and ∼50 organelle-encoded proteins, respectively. To generate and maintain fully functional organelles, these proteomes require the activity of processing peptidases, proteases, and aminopeptidases for a broad range of functions, including (a) removal

76 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

of presequences of nucleus-encoded organellar proteins; (b) N-terminal methionine cleavage of organelle-encoded proteins; (c) additional N- or C-terminal cleavages for the maturation, stabilization, and possibly activation of proteins; (d ) removal of misfolded, damaged, or aggregated proteins; (e) removal of unwanted proteins in response to environmental or developmental transitions (e.g., from glyoxysome to peroxisome or chloroplast to chromoplast) and during senescence; ( f ) release of membrane-bound proteins (e.g., transcription factors); and ( g) generation of protein degradation products as a respiratory substrate for stressed plants. The objective of this review is not only to summarize information about intraorganellar proteases, but also to identify principles that may govern substrate-protease relationships and protein lifetime as well as commonalities and differences in protease function and operation among the organelles. Over the last decade or so, there has been excellent progress in localizing and identifying organellar proteases. Loss-of-function mutants for many proteases have been identiﬁed in Arabidopsis and recently also in rice and maize. Furthermore, several proteases have been shown to be part of oligomeric structures, and for a handful there is (tentative) evidence for substrates. However, understanding of the role and mechanisms of proteolysis within plant organelles is limited, and many basic questions remain to be answered. Furthermore, intraorganellar proteolysis is complemented in surprising ways by autophagy of chloroplasts, mitochondria, and peroxisomes. Now that it is more widely appreciated that mRNA is an imperfect proxy for protein abundance (136 and references therein), it is essential to turn our attention to plant proteolysis. Proteolysis (and autophagy) must be viewed as part of proteome homeostasis, now referred to as proteostasis. Understanding the contributions of protein preprocessing, proteolysis, and autophagy to organellar proteostasis will allow us to better understand plant development and function, but will also require systems-based approaches.

COMMONALITIES BETWEEN PLASTIDIAL, MITOCHONDRIAL, AND PEROXISOMAL PROTEOLYSIS AND ORGANELLE TURNOVER There are multiple reasons why it is beneﬁcial to review proteolysis in plastids, mitochondria, and peroxisomes together. First of all, both plastids and mitochondria have an endosymbiotic origin, and they are derived respectively from cyanobacteria and α-proteobacteria. Indeed, their gene expression machineries, their protein-folding and maturation factors, and many of their proteases are of prokaryotic origin. Furthermore, more than 100 proteins are dually targeted to both chloroplasts and mitochondria (22), including several proteases, such as LON1 (34), PREP1 (85), FTSH11 (169), α-MPP2 (15), and OOP (84). (Please note that throughout this review,

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org all plant protein names are capitalized, as is the convention for several plant journals; similarly, all gene names are in italics and capitalized.) It is therefore likely that mechanisms for protease substrate recognition also have similarities. Although peroxisomes are unlikely to have an endosymbiotic origin (58), members of the prokaryotic-type protease families found in plastids and mitochondria, such as LON2 and DEG15, are also found in peroxisomes (58). Furthermore, several peroxisomal division proteins are shared with plastids and/or mitochondria, perhaps aiding in the coordination of biogenesis between these three organelles (58, 112). Particularly in leaves,

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. chloroplasts, mitochondria, and peroxisomes are functionally coupled through various metabolic pathways, such as photorespiration (between all three organelles) (16, 58), biotin synthesis (between peroxisomes and mitochondria) (110), and jasmonate biosynthesis (between peroxisomes and plastids). Therefore, coordination of their proteome compositions is needed and requires proteolysis.

www.annualreviews.org • Plant Organelle Proteolysis 77 PP66CH04-VanWijk ARI 30 March 2015 20:57

SCOPE OF THIS REVIEW, KEY QUESTIONS, AND NEW FRONTIERS IN ORGANELLAR PROTEOLYSIS

Degradomics: the Since 2010, reviews have been published on plastid proteases (73), mitochondrial proteases (67, systematic large-scale 90, 113), degradation of photosystem II (29, 194), and specific protease families in plants and or- analysis of protein ganelles, including rhomboids and other intramembrane cleaving proteases (1, 69, 86), LON (144, lifetime and 145), CLP (32, 126), FTSH (104, 182), DEG (150, 151), and processing peptidases (166). Recent proteolysis; specific degradomics tools reviews have also discussed the biogenesis of plastids (68), mitochondria (21, 186), and peroxisomes have been developed (58), as well as autophagy of plant organelles (62, 93, 98, 142). I refer to these and other reviews to help define for more extensive references and discussions, and here consolidate and conceptualize this large protease-substrate amount of information and outline new frontiers in plant organelle proteolysis. I do not discuss relationships, degrons, proteases/proteolysis in other photosynthetic organisms, such as cyanobacteria and algae, mostly and protein lifetime because of space constraints. Naturally, homologs for many of the proteases discussed in this review are also present in bacterial and algal species; although various proteolytic mechanisms are likely shared with higher plants, there will also be differences. Key questions and frontiers in plant organellar proteolysis addressed here include (a) completion of the organelle protease inventory and identification of their cleavage specificities; (b) protein preprocessing, maturation, and fate of cleaved signal peptides; (c) substrate recognition and protein lifetime determinants; (d ) proteolysis in developmental transitions of organelles and other case studies; (e) how autophagy and the proteasome complement intraorganellar proteolysis; and ( f ) systems-based degradomics approaches.

ORGANELLAR PROTEASE INVENTORY, CLASSIFICATION, AND CHARACTERIZATION A universal classification system for proteases, as well as for protease inhibitors, has been developed in which each protein is assigned to a family on the basis of statistically significant similarities in amino acid sequence (141). Protease families that are thought to be homologous are grouped together in a clan. This information is available through the MEROPS database (http://merops.sanger.ac.uk). This database includes proteases for Arabidopsis as well as scattered information for other plant species; however, the information is incomplete even for Arabidopsis and is cumbersome because of the use of various types of Arabidopsis gene identifiers [i.e., not only the Arabidopsis Information Resource (TAIR) AT identifiers, but also older numbers]. Neverthe- less, this classification system provides an excellent starting point for searching and annotating plant proteases and has identified more than 800 putative proteases in Arabidopsis,someofwhich may be esterases or other hydrolases (see also 173).

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org To facilitate systematic analysis and understanding of organellar proteolysis, we have begun to assemble and annotate an inventory of all (potential) organellar proteases and their (most likely) suborganellar localization (K.J. van Wijk & Q. Sun, unpublished work), starting with all Arabidopsis protein sequences listed in MEROPS. Additional candidate proteases were identi- ﬁed based on Arabidopsis genome-wide functional domain prediction, using the Pfam database (http://pfam.xfam.org) for putative protease and protease inhibitor domains. Including direct protease chaperones/adaptors and including all proteins that have protease-like domains, we have

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. so far assembled just over 100 proteins with experimentally determined locations in plastids (74), mitochondria (16), and peroxisomes (7) along with 8 proteins dually targeted between plastids and mitochondria, include a protease inhibitor (Kunitz type). Table 1 provides an overview of these proteins, their assignment to protease families, and their various characteristics. The TargetP subcellular localization predictor (http://www.cbs.dtu.dk/services/TargetP) suggested 200 plastid and/or mitochondrial proteases and protease-related proteins as well as 4 protease inhibitors.

78 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

Table 1 Distributions and characteristics of proteases across plastids, mitochondria, and peroxisomes, with clan and family annotations from the MEROPS database MEROPS Protease clan Plant Substrates and functions in (family) namea (family)b organellesc plants (tentative) Information from nonplant homologs Presequence processing, maturation, and recycling proteases SPP, MPPα and ME (M16) C, M N-terminal cTP and mTP are Family M16 contains metalloendopeptidases -β,PREP1–2 cleaved by SPP and MPP, with varied speciﬁcities, but cleavage is respectively. PREP1 and -2 have seldom far from a terminus of the substrate been suggested to degrade cleaved molecule and is typically upstream of an cTP and mTP (i.e., downstream arginine. The prototype is pitrilysin. of SPP and MPP). MPPα and -β are the noncatalytic and catalytic subunits, respectively. OOP, OCT1 MA (M3) C, M OOP has been suggested to Family M3 contains metallopeptidases with degrade cTPs and mTPs or other varied activities, but most members are peptides. Based on homology, oligopeptidases that typically cleave from OCT1 may aid N-terminal the C terminus. OOP homolog bacterial maturation of mitochondrial oligopeptidase A (OPDA) has been proteins. suggested to act downstream of CLP and LON. OCT1 in yeast mitochondria aids in maturation of the N terminus by cleaving octapeptides N-terminal MG (M24) C, M MAP functions in N-terminal Family M24 contains metalloexopeptidases aminopeptidase methionine removal. ICP55 is that require cocatalytic ions of cobalt or (MAP1B and expected to aid in N-terminal manganese; these are either -C, ICP55, maturation of organellar proteins. aminopeptidases or dipeptidyl peptidases. others) The prototype is methionyl aminopeptidase 1. CCTA, two SK (S41) C The substrate for CTPA is D1. Family S41 contains serine endopeptidases CTPA-liked The substrates for the CTPA-like that recognize degenerate C-terminal proteins are unknown. tripeptides and upstream cleavage sites. The prototype is E. coli C-terminal processing peptidase 1, which degrades incorrectly synthesized proteins, recognized by the C-terminal tripeptide Leu-Ala-Ala as part of the SsrA tag. Active

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org site domains are structurally related to CLP proteases. SPASE I SF (S26) C SPASE I proteases are chloroplast Family S26 contains serine endopeptidases, (PLSP1–3, envelope proteins and thylakoid most of which cleave (in an Ala-Xaa-Ala TPP) proteins with lumenal targeting motif ) N-terminal signal peptides of newly peptides. PLSP1 functions in synthesized secreted proteins with high developmentally controlled ﬁdelity. The prototype is signal peptidase I. intrachloroplast localization and Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. redox regulation. (Continued )

www.annualreviews.org • Plant Organelle Proteolysis 79 PP66CH04-VanWijk ARI 30 March 2015 20:57

Table 1 (Continued) MEROPS Protease clan Plant Substrates and functions in (family) namea (family)b organellesc plants (tentative) Information from nonplant homologs ATP-dependent proteases CLP SK (S14) C, M The CLP system is a general Family S14 contains the serine (CLPP1–P6, household protease but also has endopeptidase CLP and its homologs. CLPR1–R4∗∗, speciﬁc substrates. Examples of Without the interaction of AAA+ CLPT1–T2∗, substrates are GLUTR and the chaperones, CLPP tetradecameric protease ∗ CLPS1 , copper transporter PAA2. can act only on oligopeptides. The adaptor CLPC1–C2∗, CLPS is a substrate selector of protein CLPD∗, aggregates and N-end rule proteins to ∗ CLPX1–X3 ) CLPA. SspB is an adaptor for CLPX (a speciﬁcity-enhancing factor). FTSH1–12, MA (M41) C, M A substrate for the thylakoid Family M41 contains ATP-dependent ∗∗ FTSHi1–5 FTSH1/2/5/8 complex and DEG metalloendopeptidases that carry out proteases (S1 family) is the D1 processive degradation from the C- or reaction center protein. Inactive N-terminal end, mostly of membrane FTSHi is involved in plastid proteins, including unassembled proteins. division and likely complexes with Known bacterial substrates include sigma other FTSH family members. 32, YCCA, and SECY. The prototype is FTSH peptidase. LON1–4, SJ (S16) C, M, P Loss of function in mitochondria Family S16 contains serine endopeptidases, ∗∗ iLON1–4 leads to respiratory and folding most of which are ATP dependent. These stress phenotypes. Peroxisomal proteases degrade proteins that are LON2 is important to ensure unfolded, misfolded, or damaged. They import, and the LON2 chaperone recognize (buried) segments enriched in function is linked to autophagy. aromatic and hydrophobic amino acids, Plastid LON has not been studied. whereas negative amino acids inhibit binding. They also recognize N- and C-terminal tags (C-terminal His-Tyr). The prototype is ONA. ATP-independent proteases DEG1–16 PA (S1) C, M, P DEG proteins act as general Family S1A–F contains serine (DEG6 and -16 household proteases in plastids endopeptidases; it is the largest family and lack a protease and mitochondria and in PST2 includes nonpeptidase homologs. site) cleavage in peroxisomes. The Subfamily S1C contains DEG proteases. Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org substrate of DEG1, -2, -5, and -8 The proteases function in removing is D1. DEG15 functions in PTS2 unfolded or misfolded proteins, apoptosis, cleavage. and general proteome household processes. The prototype is chymotrypsin A (S1A). Intramembrane (RIP) proteases S2P family MM (M50) C Substrates are unknown but Family M50A–B cleaves substrates, mostly (EGY1–3, expected to be membrane-bound transcriptional regulators within or very

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. others) regulatory proteins that are close to membranes. These cleaved released upon cleavage. S2P is proteins are consequently released to take part of the M50A subfamily. on their active role in transcriptional regulation. The prototype is S2P (M50A). (Continued )

80 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

Table 1 (Continued) MEROPS Protease clan Plant Substrates and functions in (family) namea (family)b organellesc plants (tentative) Information from nonplant homologs Rhomboid (19 ST (S54) C, M, P? Substrates are unknown but Family S54 contains membrane-bound rhomboids and expected to be transmembrane serine endopeptidases as well as ∗∗ iRHOMs ) proteins. The iRHOMs are nonpeptidase homologs. Substrate nonpeptidase homologs with recognition can be outside of the unknown functions. membrane plain, but cleavage occurs within transmembrane domains. The prototype is rhomboid 1. Other proteases Alpha/beta SC (S33) C, P Substrates are unknown, but Family S33 contains mainly exopeptidases hydrolases many/most may be nonpeptidases. that act at the N termini of peptides as well as a large number of nonpeptidase homologs, e.g., epoxide hydrolases. The prototype is prolyl aminopeptidase. SPPAe SK (S49) C The substrates are likely thylakoid Family S49A–C contains serine proteins or oligopeptides. endopeptidases and can degrade cleaved signal peptides. Bacterial SPPA prefers hydrophobic amino acids on either side of the scissile bond. The prototype is E. coli signal peptidase A (S49A). M48f MA (M48) C The substrates may be Family M48A–C contains plastoglobule-localized proteins. metalloendopeptidases. Eukaryotic members cleave substrates that are prenylated at a C-terminal motif known as CAAX, in which A is an aliphatic residue, and the lipid is attached to the cysteine residue. This family is related to the M79 family. The prototype is STE24 (M48A). SCO4 M (M79) C Substrates are unknown. Family M79 contains specialized metalloendopeptidases that typically cleave a C-terminal tripeptide (CAAX cleavage) from an isoprenylated protein. This family is related to the M48 family. The prototype is RCE1 peptidase. Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org NANA, CND41 AA (A1) C The substrate for NANA is Family A1 contains aspartic endopeptidases, unknown. The substrate for mostly active at acidic pH. The prototype CND41 is Rubisco. is pepsin A. Cysteine protease CA (C85) P Substrates are unknown. Family C85 contains deubiquitinating cysteine peptidases (OTU domain). The prototype is the OTLD1 deubiquitinating enzyme.

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Serine car- SC (S28) P Substrates are unknown. Family S28 contains serine exopeptidases boxypeptidase that hydrolyze prolyl bonds. They are known only in eukaryotes. The prototype is prolyl oligopeptidase. (Continued )

www.annualreviews.org • Plant Organelle Proteolysis 81 PP66CH04-VanWijk ARI 30 March 2015 20:57

Table 1 (Continued) MEROPS Protease clan Plant Substrates and functions in (family) namea (family)b organellesc plants (tentative) Information from nonplant homologs CGEPg SC (S9) C Substrates are unknown. Family S9 contains a varied set of serine-dependent peptidases, mainly of (bioactive) oligopeptides, many of which cleave prolyl bonds. The prototype is prolyl oligopeptidase.

Details of the catalytic sites, structural features, and substrate recognition mechanisms for the (sub)families can be found in the MEROPS database (http://merops.sanger.ac.uk). Abbreviations: CGEP, chloroplast glutamyl peptidase; cTP, chloroplast transit peptide; MPP, mitochondrial processing peptidase; mTP, mitochondrial targeting peptide; RIP, regulated intramembrane proteolysis; SPP, stromal processing peptidase; TPP, thylakoidal processing peptidase. aProteins marked with an asterisk (∗) are accessory proteins rather than proteases; proteins marked with a double asterisk (∗∗) are protease homologs that lack catalytic domains. bThe MEROPS protease naming system is hierarchical and built on the concepts of catalytic type, clan, family, and peptidase (protease). There are seven catalytic types of peptidases: aspartic (A), cysteine (C), glutamic (G), metallo (M), serine (S), threonine (T), and the unusual asparagine peptide lyases (N), along with an unclassiﬁed designation. The name of a clan is formed from the letter for the catalytic type followed by a serial capital letter. A few clans contain more than one catalytic type, and these then start with P. Not all families have yet been assigned to clans. In this table, I list the clan name (from MEROPS) followed by the relevant family name in parentheses. No glutamic (G) or asparagine (N) families have yet been identiﬁed in the three organelles. cC, chloroplast; M, mitochondria; P, peroxisome. dNot detected by proteomics. eLight induced. f Senescence induced. gEnriched in bundle sheath chloroplasts compared with mesophyll chloroplasts.

Given the relatively small size of the peroxisomal proteome—estimated at ∼400 proteins (58), compared with ∼3,000 for plastids and ∼2,000 for mitochondria—we estimate that there are perhaps a dozen peroxisomal proteases. Most of the known chloroplast and mitochondrial proteases and their direct chaperones/ adaptors are ATP-dependent proteases from the AAA+ family; these include the LON, CLP, and FTSH families as well as the ATP-independent DEG family (see also sidebar AAA+ Chaperone). In Arabidopsis, these four families together consist of 61 proteins, 2 of which (LON2 and DEG15) are located in peroxisomes (Table 1). The locations of a handful of these proteins have not been experimentally established but are likely predominantly plastidic or mitochondrial, probably with the exception of a few DEG proteins. Several members of two intramembrane cleaving protease

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org families [rhomboids and site-2 proteases (S2Ps)] have been identiﬁed in the organelles (Table 1).

AAA+ CHAPERONE

The general function of the AAA+ (ATPase associated with various cellular activities) family is to trigger conformational changes in a broad range of protein substrates, thereby aiding in unfolding for degradation, refolding

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. of aggregates, and disassembly of macromolecular protein-protein and protein-DNA complexes. The typical domain architecture of proteins in this superfamily consists of an N-terminal domain that serves as a binding site for adaptor proteins and substrates, followed by one or two characteristic conserved modules (namely AAA domains or nucleotide-binding domains, each of which contains the well-known Walker A and B motifs required for ATP binding and hydrolysis) and the pore loop for substrate binding.

82 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

Second maturation step Trigger for degradation First maturation step N- or C-terminal trimming Removal of N-terminal [removes amino acid(s), • PTM CYTOSOL signal peptide cleaves a second sorting signal, • Damage and/or cleaves a TMD and Active protein • Loss cofactor Protease releases the soluble portion] substrate Preprotein • Change in oligomerization N • Misfolding/unfolding N N PTMs N Folding Import C C C C M16 S41 S26 DEG15 RIP M3, M24 (M50/M54) Proteases (e.g., LON, CLP, ±ATP FTSH, and DEG) cTP, mTP, TMD, lTP, TMD, PTS2 others others

Degradation Oligopeptidases Oligopeptidases products (e.g., M3 and S9) (e.g., M3 and S9) ORGANELLE

Amino acid pool Retrograde signal?

Figure 1 Schematic overview of processing, maturation, activation, proteolysis, and recycling of organellar nucleus-encoded proteins. Except for organellar import and cleavage of the chloroplast transit peptide (cTP) or mitochondrial targeting peptide (mTP), mitochondrion- and plastid-encoded proteins can in principle undergo the same processes. The various peptidase activities are assigned family names (e.g., M16) obtained from the MEROPS database; Table 1 describes these families in more detail and lists speciﬁc proteases for each family. Additional abbreviations: lTP, lumenal transit peptide; PTM, posttranslational modiﬁcation [e.g., (de)phosphorylation, acetylation, and oxidation]; PTS2, peroxisomal targeting signal 2; RIP, regulated intramembrane proteolysis; TMD, transmembrane domain.

At least 20 organellar proteases are (likely) involved in preprotein processing, maturation, and activation as well as recycling (Table 1). Eight organellar proteases that do not belong to any of the categories above have also recently been identified (Table 1). It is striking that, compared with those of nonphotosynthetic eukaryotes and prokaryotes, protease families in plants have greatly expanded, perhaps reflecting specific adaptation to the autotrophic and sessile nature of plants. In the following sections, I discuss these proteases and Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org families. Figure 1 and Table 1 together provide a conceptual overview of the organellar protein life cycle and illustrate the points at which the various proteases play a functional role.

Presequence Cleavage, Protein Maturation, Stabilization, and Recycling Most plastid and mitochondrial proteins are nucleus encoded and targeted through an N-terminal

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. transit peptide [chloroplast transit peptide (cTP) or mitochondrial targeting peptide (mTP)] (Figure 1). After import, cTPs are cleaved by the essential, single-gene stromal processing peptidase (SPP), whereas mTPs are cleaved by the heterodimeric mitochondrial processing peptidase (MPPα and MPPβ) (reviewed in 166); both SPP and MPP are members of the M16 metalloprotease family (Table 1). In the case of proteins with bipartite cleavable presequences (e.g., thylakoid lumenal proteins and mitochondrial intermembrane space proteins), the initial presequence

www.annualreviews.org • Plant Organelle Proteolysis 83 PP66CH04-VanWijk ARI 30 March 2015 20:57

removal is followed by additional processing steps performed by more specialized peptidases located in the relevant compartments. Examples of these peptidases are the type-I signal peptidase family (SPASE I) located in the mitochondrial inner membranes (Imp) or the chloroplast inner Degron: a degradation signal for envelope and thylakoid membranes [PLSP1, PLSP2, and thylakoidal processing peptidase (TPP)] a protein that has been (57). These peptidases are anchored at the membrane with their catalytic site on the trans side of marked for the membrane (trans, per convention, here means the opposite side of the membrane from where degradation; an the newly synthesized proteins are coming from), facilitating cleavage after protein translocation. N-degron is a degron The chloroplast-encoded D1 protein represents a classic but perhaps unusual case (discovered located at the N ∼ terminus of a protein 20 years ago) in which the C terminus needs to be cleaved by a dedicated thylakoid peptidase (CTPA) for full maturation and activation (to create a suitable ligand for cofactors in the water- splitting complex). Just recently, a loss-of-function ctpa mutant in Arabidopsis confirmed the in vivo importance of C-terminal D1 protein processing (25). Two CTPA homologs with a conserved catalytic site exist (with predicted cTPs), but unlike CTPA, they are not detected in proteomics studies; furthermore, they appear to be either not essential or not involved in D1 processing, and substrates remain to be identified (25). Peroxisomal proteins are targeted to the matrix by a non- cleavable tripeptide targeting signal at the extreme C terminus (PTS1) or a cleavable nanopeptide signal at the N terminus (PTS2) (58). PTS2 is cleaved off by DEG15, a separate clade within the DEG family, thus acting as the glyoxysomal processing peptidase. Following the removal of these organellar presequences, proteins can undergo additional trimming events, likely to avoid the presence of destabilizing amino acids at the N terminus that may form unwanted N-terminal degradation signals (N-degrons) (Figure 1). This has been elegantly demonstrated for several yeast mitochondrial proteins that are stabilized by the removal of one or eight amino acids from their mature sequences by either the ICP55 or OCT1 peptidase, respectively (177, 180; reviewed in 113). Indeed, the consensus sites for cTP/mTP cleavages are still only loosely defined, most likely because identification of in vivo organellar N termini includes additional processing steps (as in yeast mitochondria), which would mean that the in vivo N termini of such proteins do not directly reflect SPP or MPP cleavage sites. Extensive analysis of chloroplast N termini using labeling and N-terminal peptide enrichment supports the idea of additional N-terminal processing steps in chloroplasts (E. Rowland & K.J. van Wijk, manuscript in preparation). The chloroplast- and mitochondrion-encoded proteins typically undergo cotranslational N- terminal deformylation and N-terminal methionine excision carried out by two aminopeptidases (MAP1 and -2) (47, 197), followed by intraorganellar protein sorting, folding, and assembly. Both deformylation and N-terminal methionine excision are essential posttranslational modifications (PTMs) required for normal leaf development, and the absence of these PTMs can result in protein

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org degradation (111). The cleaved cTP, mTP, and PTS2 represent considerable protein mass, and they must be degraded to prevent potentially toxic accumulation of nonfunctional peptides and to help recycle amino acids. Work by the Glaser laboratory (reviewed in 83) suggested that the PREP protease can (partially) degrade cleaved cTP and mTPs. PREP is represented by two paralogs, PREP1 and -2, both of which are dually targeted to chloroplasts and mitochondria. The evidence for degradation is based mainly on recombinant PREP activity and a limited substrate set. Additionally, incubation

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. of the mTP of the ATPase β subunit with stroma or matrix proteins from wild-type and prep1 prep2 double-mutant plants showed that all mitochondrial activity but not all chloroplast activity toward synthetic mTP was lost in the double mutant, suggesting that PREP can indeed degrade cleaved mTPs. However, there is still no evidence for in vivo degradation of endogenous cleaved cTPs and mTPs following the activity of SPP and MPP, respectively. Complementation of the prep1 prep2 double knockout with a form of PREP1 that targets only to the mitochondria elegantly

84 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

Table 2 Arabidopsis protein accession numbers of members of the CLP, DEG, LON, and FTSH families Gene Name Organelle Gene Name Organelle Gene Name Organelle ATCG00670 CLPP1 C AT3G27925 DEG1 C (TL) AT1G50250 FTSH1 C (TI) AT5G23140 CLPP2 M AT2G47940 DEG2 C AT2G30950 FTSH2 C (TI) AT1G66670 CLPP3 C AT1G65630 DEG3 M∗ AT2G29080 FTSH3 M ∗ AT5G45390 CLPP4 C AT1G65640 DEG4 M AT2G26140 FTSH4 M AT1G02560 CLPP5 C AT4G18370 DEG5 C (TL) AT5G42270 FTSH5 C (TI) AT1G11750 CLPP6 C AT1G51150 DEG6 C∗ AT5G15250 FTSH6 C (TI) AT1G49970 CLPR1 C AT3G03380 DEG7 C AT3G47060 FTSH7 C(IE) AT1G12410 CLPR2 C AT5G39830 DEG8 C (TL) AT1G06430 FTSH8 C (TI) AT1G09130 CLPR3 C AT5G40200 DEG9 N? AT5G58870 FTSH9 C(IE) AT4G17040 CLPR4 C AT5G36950 DEG10 M AT1G07510 FTSH10 M AT1G68660 CLPS1 C AT3G16540 DEG11 M, C∗ AT5G53170 FTSH11 M, C (IE) ∗ AT5G50920 CLPC1 C AT3G16550 DEG12 M AT1G79560 FTSH12 C(IE) ∗ AT3G48870 CLPC2 C AT5G40560 DEG13 S AT4G23940 FTSHi1 C(IE) AT5G51070 CLPD C AT5G27660 DEG14 M AT3G16290 FTSHi2 C(IE) AT5G53350 CLPX1 M AT1G28320 DEG15 P AT3G02450 FTSHi3 C(IE) AT5G49840 CLPX2 M∗ AT5G54745 DEG16 S∗ AT5G64580 FTSHi4 C(IE) AT1G33360 CLPX3 M∗ AT5G26860 LON1 M, C AT3G04340 FTSHi5 C AT5G47040 LON2 P AT3G05780 LON3 M AT3G05790 LON4 M, C

All subcellular localizations are based on experimental results except those marked with an asterisk (∗), which are based only on prediction. Abbreviations: C, chloroplast/plastid; IE, inner envelope of chloroplast; M, mitochondria; N, nucleus; P, peroxisome; TI, integral membrane thylakoid protein; TL, thylakoid lumen protein; S, predicted signal peptide.

showed that the strong growth phenotype of the double mutant was due to loss of PREP1 activity in chloroplasts rather than in mitochondria (85). A recent study showed that the protease OOP, also dually targeted to chloroplasts and mitochondria, can degrade peptides with a length between 8 and 23 amino acids. Kmiec et al. (84) suggested that OOP operates downstream of PREP1 and -2 and/or the CLP protease. Similarly, an earlier study of the OOP homolog in Escherichia coli (oligopeptidase A) also suggested that it functions downstream of LON and CLP proteases (65). It is interesting that SPP, MPP1 and -2, and PREP1 and -2 together with several other proteins form the Arabidopsis family of M16 metalloproteases; whether these additional M16 proteases are Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org also involved in cTP or mTP removal is unknown.

ATP-Dependent Proteases of the AAA+ Family: LON, CLP, and FTSH A signiﬁcant number of organellar proteases are ATP-dependent members of the AAA+ superfamily. These proteases bind and hydrolyze ATP to facilitate the unfolding of protease substrates prior to cleavage. Many of these proteins belong to the LON, CLP or FTSH family (Table 2). Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Figure 2 shows a schematic outline of the organization of these protease systems as well as the ATP-independent DEG protease system.

The LON family. The ATP-dependent LON proteases are widely conserved serine peptidases of the S16 family. LON is a tandem fusion of an AAA+ molecular chaperone and a protease with a serine-lysine catalytic dyad, which together form soluble single-ring hexameric complexes in

www.annualreviews.org • Plant Organelle Proteolysis 85 PP66CH04-VanWijk ARI 30 March 2015 20:57

LON Mitochondrial CLP FTSH

Adaptors? ATP ADP + Pi Hexamer Hexamer ATP ADP + Pi CLPX hexamer ADP + P i ATP Heptamer (CLPP2)6

Heptamer (CLPP2)6

DEG

Trimer

Trimer 12-mers Cage 24-mers

Trimer

Figure 2 + Highly simpliﬁed models of the three major AAA protease systems (LON, CLP, and FTSH) and the ATP-independent DEG protease system in the organelles. Substrates must be recognized by N-terminal, C-terminal, or internal amino acid residues within the substrates. In the case of the AAA+ proteases, recognition involves auxiliary domains of the AAA+ proteins and possible adaptor proteins (e.g., CLPS1 in the case of chloroplast CLP). In the LON system and the membrane-bound FTSH system, the protease and ATPase domains are within the same proteins, whereas in the CLP system, separate AAA+ proteins associate with CLP proteases. The AAA+ proteins typically form hexameric hetero- or homo-oligomers. DEG protease are soluble but can associate with membranes. DEG proteases in prokaryotes can function as trimers, hexamers, or even higher-order oligomers (12- and 24-mers); oligomerization is dynamic and involves PDZ domains.

bacteria and human mitochondria (Figure 2) but, surprisingly, heptameric rings in yeast (176). In nonplant systems, LON proteases preferentially degrade unfolded and misfolded/damaged

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org proteins by recognizing unstructured and exposed protein segments that are enriched in aromatic and hydrophobic residues but lack negative residues. LON also recognizes N- and C-terminal degradation tags; for example, the LON substrate SulA in E. coli has a moderately hydrophobic C- terminal tag that ends with a histidine preceded by a tyrosine. Fusion of this tag to other proteins is sufﬁcient to direct them for degradation by LON (reviewed in 52). Plant organellar LON substrate recognition mechanisms or degrons have not been determined. The Arabidopsis genome encodes four soluble (type-A) LONs (LON1–4) (144). Experimentation showed that LON2 is

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. peroxisomal with a PTS1-targeting signal, LON1 and -3 are mitochondrial, and LON4 is dually targeted to plastids and mitochondria (128). The oligomeric state of plant LON proteins has not been determined (but for discussion, see 144). Two independent studies have described the Arabidopsis lon1 loss-of-function mutant (143, 160). The lon1 mutant alleles result in reduced growth and respiration, an aberrant mitochondrial morphology, increased levels of mitochondrial chaperones, a decrease of several mitochondrial

86 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

inner-membrane complexes, and differential changes within the tricarboxylic acid cycle proteome. LON2 localizes to peroxisomes in four-day-old cotyledons but to the cytosol in older seedlings (101). The lon2 Arabidopsis phenotype suggests that LON2 facilitates continued matrix protein mAAA and iAAA import in mature peroxisomes, but LON2 substrates are not yet known. Interestingly, a deﬁciency proteases: in peroxisomal matrix protease LON2 causes enhanced autophagy of peroxisomes as well as mitochondrial FTSH peroxisomal protein accumulation in the cytosol due to a decrease in the number of peroxisomes. proteins with the This is because LON2 plays dual roles as an ATP-dependent protease and as a chaperone essential active site facing the for suppressing autophagy. Thus, the intermolecular modulation between protease and chaperone matrix (mAAA) or the intermembrane space function of LON2 regulates autophagy (13, 41, 49, 193) (see also Organelle Proteome Turnover (iAAA) Through Autophagy and Targeted Vesicle Transport, below). In addition to these four conserved LON proteins, the Arabidopsis genome encodes four proteins with sequence similarity to the LON proteins; however, they lack the conserved catalytic LON domain (K. Nishimura & K.J. van Wijk, unpublished results). We suggest naming these iLON proteins, by analogy with iRHOMs, the inactive forms of the rhomboids. Proteomics data suggest that three of these proteins are localized to chloroplasts; the localization of the fourth is unclear, but it lacks a predicted N-terminal signal sequence for the chloroplast, mitochondria, or endoplasmic reticulum.

The FTSH and FTSHi family. FTSH proteases are membrane-bound ATP proteases of the AAA+ family (M41) with one or two transmembrane domains (TMDs) in the N-terminal portion, followed by the ATPase domain and a C-terminal catalytic protease domain and zinc-binding motif (His-Glu-X-X-His) (Figure 2). The number of TMDs affects the topology, in particular whether the catalytic site is on the cis or trans side of the membrane. FTSH proteases form homo- or heterohexameric complexes with a chamber that contains the internal catalytic sites and enables controlled access through an adaxial pore facing away from the membrane. FTSH proteins have a prokaryotic origin and are also found in plastids and mitochondria; they have been extensively studied in a wide range of species (for recent reviews, see 66, 124, 182) (Table 2). The Arabidopsis genome encodes 12 FTSH proteins. FTSH3 and -10 (each with two TMDs; also called mAAA-type proteins) form homo- or hetero-oligomers in the mitochondrial inner membrane with the active site facing the matrix, whereas FTSH4 and -11 (each with one TMD; also called iAAA-type proteins) locate as homo-oligomers in the mitochondrial inner membrane with the active site facing the intermembrane space; FTSH11 is also targeted to the inner envelope membrane of chloroplasts. FTSH7, -9, and -12 are located in the chloroplast inner envelope membrane. FTSH1, -2, -5, and -8 (each with one TMD) form a 400–450-kDa heterohexamer in the thylakoid membrane with the active site facing the stroma. FTSH6 is also localized in plastids,

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org but the intraplastid location is unclear. Additionally, there are ﬁve FTSH proteins that lack their catalytic protease domain but retain their chaperone domain (FTSHi1–5); these proteins are likely located in the inner chloroplast envelope (182) and coexpress as a group together with FTSH12 (71). In the mitochondria of various yeast species, FTSH complexes associate with prohibitin proteins. Such an interaction also occurs for Arabidopsis mitochondrial FTSH3 and -10, forming a 2-MDa complex (134). Prohibitins have been proposed to function as scaffold proteins in a variety of processes in mitochondria and in nonplant species outside of mitochondria; the function of

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. their interaction with plant FTSH3 and -10 is perhaps to stabilize FTSH complexes (134, 172). In particular, the function and organization of the thylakoid heteromeric FTSH complex (FTSH1, -2, -5, and -8) have been extensively characterized through loss-of-function mutants, suppressor screens, structural analysis, and detailed biochemical characterization of chloroplasts from such loss-of-function mutants (for reviews, see 66, 124, 182). Single-mutant phenotypes range from variegated phenotypes with reduced biomass (loss of FTSH2 or -5) to no visible

www.annualreviews.org • Plant Organelle Proteolysis 87 PP66CH04-VanWijk ARI 30 March 2015 20:57

phenotype (loss of FTSH1 or -8). Interestingly, the variegated phenotype of the ftsh2 mutant was suppressed by complementation with an FTSH2 gene lacking the catalytic protease site but not lacking the chaperone activity (147). A suppressor screen of the ftsh2 mutant identified nearly N-end rule: a rule for the degradation of a dozen suppressors of the variegated phenotype (reviewed in 138). Thylakoid FTSH, together N-degron substrates with DEG proteases, is most likely involved in general household proteolysis of the thylakoid that relates the proteome, including degradation of the D1 protein of photosystem II as part of the repair cycle regulation of the in (for reviews and discussion, see 29, 116, 193). vivo half-life of a The function of FTSH6 is not clear, and earlier reports stating that FTSH6 is involved in de- protein to the identity of its N-terminal grading specific light-harvesting complex II proteins appear to be incorrect (183). Dually targeted residue FTSH11 loss-of-function alleles in Arabidopsis were identified in a screen for thermosensitive mutants, but the mutants were not sensitive to light stress (28). The mutants were arrested in growth and development through their life cycle at temperatures above 30◦C, which is atypical for Ara- bidopsis. This function of FTSH11 resembles the function of yeast mitochondrial FTSH YME1, and heat sensitivity was not found in other Arabidopsis ftsh mutants. Whether this thermointoler- ance results from mitochondrial or chloroplast activity remains to be determined. The functions of mitochondrial FTSH members FTSH3, -4, and -10 have not been studied. FTSHi1 (also named ARC1) is essential for embryogenesis because of its role in chloroplast division; underexpression of FTSHi results in premature plastid division but has no direct effect on FTSZ division protein levels, whereas the functions of the other (coexpressing) FTSHi members are not known (71). Very little information is available on how FTSH proteases select their targets for degradation, even when considering nonplant species. A recent paper generated an FTSH-trap mutant for E. coli in which substrates were trapped inside the partially inactivated complex (187). Several substrates were identified, some of which were also trapped in bacterial LON or CLPPX complexes, but so far this has not helped define substrate recognition principles.

The CLP family. The serine-type, ATP-dependent CLP protease system (S14) is the most abun- dant and complex stromal protease in the plastid and is also present in mitochondria (reviewed in 126) (Figures 2 and 3, Table 2). CLP systems are extensively studied in prokaryotes, have a general household function, and regulate speciﬁc pathways or processes (reviewed in, e.g., 149). Because of their similarities, the E. coli CLP system provides an excellent template for studying the plant CLP system. The E. coli CLP protease is a barrel-like core complex with two stacked heptameric rings of CLPP protease subunits. This core complex associates with a hexamer of CLPA/X ATP-dependent chaperones. The catalytic sites of the 14 identical CLPP protease subunits are enclosed within the barrel-shaped structure. Generally, only peptides and proteins in an unfolded state can enter the cavity, and CLPA/X chaperones and ATP hydrolysis are required to

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org unfold and feed proteins into the CLP protease cavity. Recognition of substrates occurs via N- or C-terminal tags or via internal domains and is further regulated by adaptor proteins, such as E. coli SspB (for CLPX) and CLPS (for CLPA). Binding of CLPS to CLPA shifts the afﬁnity of CLPA toward protein aggregates and so-called N-end rule proteins. (For an up-to-date comprehensive review on the plant CLP system, see 122.) Plants contain a chloroplast (plastid) CLPAP-type system, and mitochondria contain a CLPXP system (126). A CLPS homolog named CLPS1 (117) is present in plant plastids, but there is (so

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. far) no evidence for SsrA or SsrB in plants. Figure 3 shows a summarizing model and substrate delivery system of plant CLP. Arabidopsis plastids and chloroplasts contain ﬁve serine-type CLP proteases (CLPP1 and CLPP3–P6) and four nonproteolytic CLPR subunits (CLPR1–R4), which together comprise the tetradecameric and asymmetric ∼350-kDa CLP protease core formed by two heptameric rings. Furthermore, three CLP AAA+ chaperones (CLPC1, -C2, and -D), similar to E. coli CLPA and the adaptor CLPS1, deliver protein substrates to the chaperone complex.

88 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

N-end rule substrates N-degron Other CLPS1 substrates

Aggregates ATP ADP

600 kDa Hexamer CLPC1–C2

Heptamer CLPP1, CLPR1–R4 (R ring) Degradation CLPPR core products Heptamer CLPP3–P6 (P ring) 350 kDa Oligopeptidase [e.g., OOP (M3)] CLPT1–T2

Amino acids ? ? Unstructured substrates (cTPs)

Figure 3 Organization of the chloroplast CLP protease and its substrate delivery and downstream recycling. This schematic view of the chloroplast CLP system is based on experimental information for Arabidopsis and functional homology to the Escherichia coli CLPAPS system. The CLP system exempliﬁes many of the regulatory and structural features of proteases: Substrates are selected through various signals intrinsic to the substrate, explaining the wide variety of likely substrates; an adaptor protein, CLPS1, enhances the delivery of proteins that possess speciﬁc features (e.g., an N-degron) at the expense of other substrates; ATP- dependent chaperone activity helps to progressively unfold selected (folded or aggregates) substrates, which is immediately followed by transfer into the proteolytic chamber of the barrel-like CLPPR core complex; and the collective activity of the catalytic sites (facing into the chamber) of the CLPP subunits cleaves the trapped substrates, which is followed by the release of small peptides (probably approximately 8–10 amino acids long). Downstream oligopeptidases degrade these peptides into individual amino acids. CLPT1 and -T2 are found only in chloroplasts of higher plants and play a role in stabilizing the CLPPR core complex and perhaps facilitating delivery of unstructured substrates, such as chloroplast transit peptides (cTPs).

Attached to the CLPPR core are CLPT1 and -T2, which have high sequence similarity to the N- terminal domain of the CLPC/D chaperones (126, 132). These have been hypothesized to regulate chaperone binding and/or substrate selection (126) or to aid in the assembly or stabilization of the CLPPR core complex (157; J. Kim & K.J. van Wijk, manuscript in preparation). By contrast, plant

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org mitochondria contain a homotetradecameric CLPP2 core complex and three CLPX chaperones. Beyond the identification of this CLPP2 core by native gels, no experimental analysis has been carried out for this mitochondrial CLPPX system. Olinares et al. (125) recently determined the quantitative subunit compositions for the intact CLPPR core and each heptameric ring. The chloroplast CLPPR protease was affinity purified from Arabidopsis clpr4 and clpp3 null mutants complemented with C-terminal Strep II–tagged versions of CLPR4 and -P3, respectively. The subunit stoichiometry was determined by a mass spectrometry–

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. based multiplexed absolute quantiﬁcation strategy using a concatemer of stable isotope–labeled proteotypic peptides generated from a synthetic gene, also known as the QconCAT approach (137). The results showed that the CLPPR core consisted of one heptameric ring (called the P ring) containing CLPP3, -P4, -P5, and -P6 in a 1:2:3:1 ratio and another ring (called the R ring) containing CLPP1, -R1, -R2, -R3, and -R4 in a 3:1:1:1:1 ratio. Moreover, based on biochemical and phylogenetic analysis, it was suggested that CLPT1 and -T2 bind to the adaxial side of the

www.annualreviews.org • Plant Organelle Proteolysis 89 PP66CH04-VanWijk ARI 30 March 2015 20:57

P ring (125, 157). This CLP core complexity is puzzling and suggests a specific adaptation of the CLPPRT protease to the chloroplast proteome. Genetic and phenotypic analyses of various clpp and clpr mutants in Arabidopsis showed that Virescent: displaying a reduced greenness of each of the tested genes affects embryogenesis or seedling development and chloroplast biogene- leaves; many sis. Interestingly, the severity of the phenotypes differs greatly among the various clpp and clpr null loss-of-function mutants (77). In the case of CLPP3, -P4, and -P5 and perhaps CLPP6, the severity of the pheno- mutants for type follows the copy number in the CLP core. Complete loss of gene expression for CLPP5 (three chloroplast proteases copies) and CLPP4 (two copies) is embryo lethal, whereas null mutants for CLPP3 can germinate, and other chloroplast proteins have such a develop seedlings under heterotrophic (but not autotrophic) conditions, and even produce viable phenotype seeds (77). Complete loss of CLPR2 or CLPR4 delayed embryogenesis and resulted in developmental arrest in the cotyledon stage (78). This arrest could be broken by growth on sucrose, but seedlings remained sterile (78, 126). By contrast, the Arabidopsis clpr1-1 null mutant has only a weak virescent phenotype because CLPR1 is partially redundant with CLPR3. In addition, CLPP1, the only plastid-encoded CLP subunit (three copies per core), is essential for leaf development in tobacco. Comparative quantitative proteomics studies of virescent clpr2, clpr4,andclpp3 mutants showed very similar metabolic and protein homeostasis defects, indicative of chloroplast protein- folding stress (all chloroplast chaperone systems are strongly upregulated), strong downregulation of photosynthesis, and compensatory upregulation of the ATP import pathway (77). This is consistent with the idea that chloroplasts contain a single CLPPR core complex. Similarly to CLPR1 and -C1 (138), CLPR4 suppresses the variegated phenotype of loss-of-function alleles for FTSH2 (190). CLP core mutants were also identified in maize (for CLP5) and rice (for CLPP6), and their pale green and growth phenotypes are consistent with an important role of the CLP protease in monocotyledons and with previous proteomics observations in developing maize leaves (107) (see Figure 4, below). The Arabidopsis clps1 null mutant lacks a visible phenotype, and no genetic interactions with the CLPC/D chaperone or CLPPR core mutants were observed. However, clps1 (but not clpc1-1) has increased sensitivity to the translational elongation inhibitor chloramphenicol, suggesting a link between translational capacity and CLPS1. Moreover, CLPS1 was upregulated in the clpc1-1 mutant, and quantitative proteomics of clps1, clpc1,andclps1 clpc1 mutants showed specific molecular phenotypes attributed to loss of CLPS1 or CLPC1. Affinity purification identified candidate CLPS1 substrates, including glutamic acid tRNA reductase, a control point for tetrapyrrole synthesis. CLPS1 interaction with five substrates strictly depended on two conserved CLPS1 residues involved in N-degron recognition (121). The N-terminal domain of CLPC is necessary for interaction with the chloroplast inner envelope and for functional complementation of the virescent phenotype (30). Furthermore, both CLPC2 and CLPD can interact with cTPs, presumably soon

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org after they cross the inner membrane through the TIC complex and perhaps before they are cleaved off by SPP (18). The main challenge for understanding the plant CLP system is to identify more direct in vivo substrates, perhaps through substrate trapping, as was so successful for E. coli CLPP (61) and Staphylococcus CLPP (42). Additionally, reconstitution of plant CLP protease activity will be needed to determine the molecular mechanisms underlying substrate delivery and proteolysis and the signiﬁcance of the complexity of the plastid CLPPR protease core. The in vivo signiﬁcance

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. of mitochondrial CLP remains to be determined.

The ATP-Independent DEG Protease Family DEG proteases [also called high-temperature requirement A (HtrA) proteases] are ATP- independent serine endopeptidases in the S1 family (53). They typically comprise an N-terminal

90 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

proteolytic domain (with a serine–histidine–aspartic acid catalytic triad) and one or two C-terminal PDZ domains. They are often peripherally attached to membranes but can also be soluble. Their basic native structure is a trimeric ring mediated by interactions between the protease domains. Catalytic triad: three These trimers can form hexamers through the stacking of two trimers, mediated by PDZ domains, amino acid residues thus forming a catalytic cage; however, even 12- and 24-mers can form. Protease activity and chap- that function together erone activities are influenced by temperature and oligomeric state. Described DEG functions at the center of the include counteracting protein-folding stress as well as proteome remodeling, apoptosis, and gen- active site of proteases eral proteome quality control. Substrate recognition is regulated by (a) the PDZ domains, which (or other enzymes) influence the substrate specificity, as well as by (b) the processivity of the protease domain (53). The Arabidopsis genome encodes 16 DEG proteins, which are related to bacterial DEGP rather than DEGS or DEGQ (Table 2). The 16 family members vary in the number of PDZ domains (zero, one, or two), and DEG6 and -16 lack the conserved catalytic protease site. DEG5 and -15 lack PDZ domains, and DEG7 is a tandem duplication with two protease and four PDZ domains. (For the DEG phylogeny and nomenclature for Arabidopsis, rice, and poplar and a general review of plant DEG proteases, see 150, 151.) Six Arabidopsis DEG proteins have been shown to locate in chloroplasts, namely DEG1, -5, and -8 on the lumenal side of the thylakoid membrane and DEG2, -7, and -11 in the stroma and/or peripherally associated to the thylakoid. DEG15 is located in peroxisomes (35, 55), DEG10 and -14 are located in mitochondria (14), and DEG9 is located in the nucleus (150, 151). The locations of the other DEG proteins (DEG3, -4, -6, -12, -13, and -16; Table 2) have not been experimentally studied, and their accumulation levels appear to be low because most are not detected by proteomics techniques (see, e.g., the Plant Proteome Database; http://ppdb.tc.cornell.edu). Except for DEG13, these other DEGs are predicted to localize to mitochondria and/or plastids. Functions for Arabidopsis mitochondrial DEG14 (14) and peroxisomal DEG15 (35, 55, 152) have been studied in vitro and in vivo. Accumulation of DEG14-encoding transcripts is induced by heat stress but not by other stress conditions; consistently, overexpression of DEG14 confers thermotolerance. DEG14 degrades misfolded proteins in particular. Thus, the main function of mitochondrial DEG14 appears to confer thermotolerance by degrading misfolded proteins. DEG15 functions as the glyoxysomal processing peptidase for proteins with a PTS2 signal sequence (55, 152). Interestingly, DEG15 can change its function from a processing peptidase (as a dimer) to a general protease (as a monomer, cleaving downstream of arginine/lysine) through a calcium-concentration-dependent and calmodulin (CLM3)–mediated switch in oligomeric state (35). Phylogenetic analyses indicate that the calmodulin-binding site is conserved in peroxisomal processing proteases of higher plants but not in nonphotosynthetic eukaryotes. Neither DEG15 (152) nor CLM3 (35) knockout plants have a growth phenotype, but both showed increased

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org resistance to the herbicide 4-(2,4-dichlorophenoxy)butyric acid (2,4-DB). Great progress has been made in structural analysis for chloroplast DEG proteases, and the regulation of oligomerization and consequences for DEG activity show fascinating variation. X- ray structures have been determined for the three lumenal DEG proteins (DEG1, -5, and -8) (81, 163) and the stromal protein DEG2 (162). The lumen is an unusual compartment in that acidiﬁcation occurs in the light. DEG1 accumulates as an inactive monomer and transforms under (even slightly) acidic conditions into the proteolytically active but rigid hexamer, formed by two

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. stacked trimers with the active sites sequestered inside the cage (81). This is a dynamic equilibrium in the dependence of pH and protein concentration. This system ensures selective activation of DEG1 during daylight, when the lumen acidiﬁes and photosynthetic proteins are damaged, which is consistent with the observed in vivo degradation function (72) and perhaps also the chaperone function (164). Crystallization of recombinant stromal DEG2 showed a hexameric structure (162), but a more active form induced by the presence of substrate is a larger oligomer (12- or 24-mer)

www.annualreviews.org • Plant Organelle Proteolysis 91 PP66CH04-VanWijk ARI 30 March 2015 20:57

similar to bacterial DEGP/Q when incubated with substrates. DEG5 and -8 were crystallized as shallow, funnel-shaped trimers with the catalytic triad on the concave side of the funnel, but these represent proteolytically inactive states (163). The DEG5 trimer contains two calcium ions in a central channel, suggesting a link between its activity and calcium ions in chloroplasts. These three structural studies show a variety of DEG activation mechanisms, all of which involve modiﬁcation of the oligomeric state; thus, unlike CLP proteases, which employ chaperones and their unfolding activity to control proteolysis, DEG proteins use structural changes, induced by external factors (pH, substrates, and perhaps calcium) to regulate substrate access to the proteolytic chamber. Functional analyses of lumenal DEG proteases and DEG2 have identiﬁed cleavages in soluble loops of the D1 protein as the key function of these chloroplast proteases. The activity of these DEG proteases is complemented by the activity of the thylakoid FTSH complex (reviewed in 29, 116, 193). Other lumenal proteins (e.g., OEC33 and plastocyanin) likely also serve as substrates for lumenal DEG proteases.

Intramembrane Cleaving Proteases of the Rhomboid and S2P Families in the Organelles All protease families discussed above cleave proteins in loops and domains exposed to the aqueous environment of the cell. However, some proteases can cleave proteins within or very close to the lipid bilayer, a process called regulated intramembrane proteolysis (RIP). These RIP proteases are members of the rhomboid, S2P, and SPP-like families and are present in most species (171), including plants (Table 1). The catalytic site residues of RIP proteases project into a cavity that is buried within the membrane lipid bilayer. The realization that RIP plays an important role either in the completion of protein maturation or in cell signaling—e.g., by release of membrane- bound transcription factors (∼10% of all transcription factors are membrane anchored) and other effectors—has triggered many studies (170, 171). RIP proteases are well represented in plants, including in plastids and mitochondria (recently reviewed in 1, 69). I do not discuss the plant SPP- like family here because none of the seven members in Arabidopsis appear to localize to plastids, mitochondria, or peroxisomes.

The rhomboid family. The Arabidopsis genome encodes 19 rhomboid-related proteins (S54 family), but there is no consistent naming for Arabidopsis and other plant species. Some of these may lack proteolytic activity and are named iRHOM proteins; however, in nonplant systems, these iRHOMs have important functions, e.g., in endoplasmic reticulum–associated degradation (2). Fewer than half of the Arabidopsis rhomboid-related proteins are predicted to locate to mitochon-

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org dria or plastids, and only one has a predicted endoplasmic reticulum signal peptide. Based on various experimental data, two rhomboids are located in the Golgi apparatus (AtRBL1 and -2), two are located in the plastid inner envelope membrane, and one is located in the mitochondria (PARL). The functions and substrates of these rhomboids are unknown, but a loss-of-function mutant for plastid-localized AT1G25290 resulted in an aberrant pollen phenotype and various abnormalities in the ﬂowers, leading to reduced fertility and a root phenotype (167). The role of this rhomboid in nonphotosynthetic plastids and chloroplasts may relate to jasmonate metabolism,

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. the ﬁrst step that occurs in plastids (87, 167). Jeyaraju et al. (69) described a general model for the recruitment and cleavage of substrates by rhomboids.

The S2P family. The S2P family of RIP proteases consists of ﬁve plastid proteins—EGY1 (26, 96), EGY2 (27), EGY3, AT2G32480 (also named AraSP; 17), and AT1G05140—as well as AT4G20310, for which the subcellular location is unknown. All six S2P proteins have multiple

92 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

predicted TMDs. The three EGY proteins are integral thylakoid proteins, whereas AT2G32480 and AT1G05140 are likely located in the inner chloroplast envelope. Loss-of-function Arabidopsis mutants for EGY1 and AT2G32480 have reduced growth and chloroplast defects, whereas the mutant for EGY2 has a more subtle phenotype. Consistently, loss-of-function mutations in the EGY1 homolog in tomato result in chlorophyll-deﬁcient phenotypes, including reduced rates of chlorophyll synthesis during de-etiolation and enhanced rates of chlorophyll loss in leaves and fruits as they age. EGY2 is coexpressed with a plastid rhomboid, AT1G25290, and they may work in sequence (167).

Recent Discoveries and Functional Analysis of Other Organellar Proteases In this section, I briefly summarize recent identification and characterization of organellar proteases that do not belong to the families discussed above and are not discussed in the context of protein processing and maturation (Table 1). The SPPA protease was identified as a light-induced thylakoid protease more than a decade ago (95), and a subsequent study showed that it plays a role in light acclimation, consistent with its light- induced accumulation (188). The snowy cotyledon 4 (sco4) mutant was identified as one of several mutants with white cotyledons (4). SCO4 is distantly related to CAAX proteases, and loss-of-function mutants are sensitive to high light. A recent study also identified a chloroplast-localized aspartic acid protease that the authors named NANA (“small” in Italian) because the loss-of-function mutant has a dwarf phenotype along with an altered leaf morphology and a delayed flowering time (131). NANA is related to the aspartic acid protease CND41, which was characterized in both tobacco and Arabidopsis as being involved in Rubisco degradation during senescence (see 131 and references therein). It was suggested that NANA has a regulatory function in chloroplasts that influences not only photosynthetic carbon metabolism but also plastid and nuclear gene expression. A comparative proteomics study of peroxisomes from etiolated seedlings and green leaves also identified a cysteine protease that is a member of the C1 family containing deubiquitinating peptidases (139); the protease was named RESPONSE TO DROUGHT21A-LIKE1, and mutant analysis showed that it affects functions including β-oxidation, seed germination, and growth. Comparative proteomics and mass spectrometry studies have identified two other chloroplast proteases, M48 and chloroplast glutamyl peptidase (cGEP). The M48 protease was identified as a low-abundance protease specifically located in the thylakoid-associated lipoprotein particles called plastoglobules (106). Coexpression analysis based on public microarray data and experimental follow-up (N.H. Bhuiyan & K.J. van Wijk, unpublished data) confirmed its protease activity and clearly placed its function in the context of senescence. The M48 family contains proteases that cleave prenylated proteins and is related to the M79 family, which includes the SCO4 protein. Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org A comparative proteomics study of maize bundle sheath and mesophyll chloroplasts showed that

CGEP preferentially accumulates in bundle sheath chloroplasts, suggesting involvement in C4 chloroplast differentiation (107). A proteomics study also identified a serine carboxypeptidase (S28) in peroxisomes (102). Finally, one peroxisomal and several plastid-localized members of the S33 family have been identified. This family contains mainly exopeptidases that act at the N termini of peptides, along with a large number of nonpeptidase homologs (e.g., epoxide hydrolases). Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. PROTEASE NETWORKS, PROTEOSTASIS, AND RETROGRADE SIGNALING Plastids and mitochondria contain many proteases and aminopeptidases. These enzymes can differ in their substrate and cleavage site specificity, and some proteases (such as OOP) have a maximum length or mass limit for their substrates. Protein accumulation levels also differ and

www.annualreviews.org • Plant Organelle Proteolysis 93 PP66CH04-VanWijk ARI 30 March 2015 20:57

Leaf and chloroplast development 0–1 cm 4–5 cm 8–9 cm 13–14 cm

3 μm 3 μm 3 μm 3 μm

3.5

3.0

2.5

2.0

(NSAF ×1,000) 1.5 CLPPRT core 1.0 CLPC1–2 (2×) Relative protein abundance FTSH1, -2, -5, and -8 0.5 PREP1 (10×) DEGP2 (20×) 0.0 02468101214

Distance from base (cm) Figure 4 The dependence of chloroplast protease accumulation on maize leaf and chloroplast development, showing that that the protease composition changes based on the developmental state and cellular functions. The relative protein concentrations for the chloroplast CLPPRT core, CLPC1 and -C2 chaperones, stromal PREP1 and DEGP2, and thylakoid FTSH complex were determined by label-free spectral counting of different sections along the developing leaf. As illustrated by the electron microscopy images (top), the base of the leaf contains nonphotosynthetic proplastids that gradually develop into full photosynthetic chloroplasts at the tip of the leaf. Abbreviation: NSAF, normalized spectral abundance factor. Adapted from Reference 107.

can be inﬂuenced by developmental and metabolic state or by cell and tissue type, as shown for developing maize leaves (Figure 4). It is likely that proteins can be cleaved (in parallel) by different proteases, perhaps depending on the speciﬁc PTMs, and/or that proteases are degraded

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org by the sequential activity of proteases and aminopeptidases. There are two cases in chloroplasts where parallel and/or sequential activities of different proteases have been explored. In the case of the D1 protein, both the FTSH and DEG proteases and perhaps even the CLP protease are involved in its degradation, as discussed above. Because FTSH proteases typically processively degrade proteins, FTSH most likely acts downstream of the lumenal and stromal DEG proteases. The OOP protease has been suggested to degrade cleavage products generated by PREP1/2 and/or the CLP protease system. However, oligopep-

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. tidase activity is needed downstream of several other proteolytic activities as well, as outlined in Figure 1. Suppression mutant screens for the variegated chloroplast FTSH2 mutant var2 yielded multiple alleles, including CLP protease family members (138); this suggests that either the CLP protease can cover for FTSH2 activity or, more likely, mutations in CLP genes lead to delayed chloroplast biogenesis and therefore alleviate the requirement for FTSH activity during a critical phase in chloroplast biogenesis. It is interesting that yeast mitochondria lack a CLP protease

94 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

PTM (e.g., N-degron) Delivery proteins (adaptors) PTM Conformation Unfoldases Conformation Oligomerization ATP Oligomerization Stable Active Inactive protein Substrate protease protease

Figure 5 Conceptualized view of protease activation and the generation of protease substrates. Proteins can become protease substrates after undergoing one or more modiﬁcations as indicated. Some proteases require activation, which could involve a posttranslational modiﬁcation (PTM), oligomerization, or a conformational change.

system, and it is believed that the LON protease carries out most CLP functions. By contrast, plant mitochondria do contain a homomeric CLPP2 core and three CLPX chaperones (Figure 2)that function in parallel to LON1 and -4; whether and (if so) to what extent the mitochondrial CLP and LON protease functions are intertwined remain to be determined. Retrograde signaling from organelles to the nucleus is important to instruct the nuclear genome about the physiological state of the organelles. Multiple retrograde pathways have been proposed for plastids and mitochondria (11, 40, 120). Loss of protease activity in the organelles likely results in organellar signaling (e.g., in clp protease mutants resulting in multifold upregulation of all plastid chaperone systems; 77, 78, 196), perhaps directly involving proteolysis, as was demonstrated in the nematode Caenorhabditis elegans (115).

SUBSTRATE SELECTION, PROTEIN LIFETIME DETERMINANTS, DEGRONS, AND POSTTRANSLATIONAL MODIFICATIONS It is critically important that proteases only cleave proteins specifically selected for degradation (Figure 5). One way that this is achieved is by shielding the protease active site within an oligomeric structure (sequestration of catalytic sites), such as the barrel-like structures of the CLP protease core complex. Proteins can be actively degraded by such proteases only if they are linearized and threaded into the protease cavity. For folded proteins, this threading requires ATP-dependent chaperone activity, either by specific chaperones (e.g., CLPC or CLPX) or by domains in the actual protease (e.g., FTSH). Selectivity can also be triggered by (a) conformational changes or changes in oligomeric state, leading to exposure of (often hydrophobic) domains to the surface, and (b) specific degrons through the addition of a specific tag, such as ubiquitin in the case of the proteasome and an N-terminal tag in the case of N-degrons in bacteria. Surprisingly, no tagging system has yet been discovered in any of the three organelles, and ubiquitination is not known to take place within the organelles (although it does occur at the outward surface facing Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org the cytoplasm). These substrate recognition sites can be collectively called degrons. Degrons can be functionally identified by mutations that prevent degradation of known substrates or by sequences that confer susceptibility to a protease when appended to a protein that is not normally degraded. The degron accessibility for one protease can also be controlled by endo- proteolytic processing by a different protease [for an extensive discussion of types and recognition of degrons of AAA+ proteases (CLP, LON, and FTSH), see 149]. Substrate delivery or adaptor proteins have been identified that recognize and select substrates Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. and deliver them directly or indirectly to specific proteases. A clear example for the photosynthetic bacterium Synechocystis is the NblA1/A2 heterodimer protein, which delivers phycobilisome proteins to the CLP protease (10). Other, more general adaptor proteins for the CLP system have been found in various prokaryotes, such as CLPS, which recognizes and delivers N-degron- containing proteins as well as protein aggregates (37, 39, 146, 184). As discussed above, CLPS is present in plant chloroplasts and likely also functions in substrate selection and delivery (121);

www.annualreviews.org • Plant Organelle Proteolysis 95 PP66CH04-VanWijk ARI 30 March 2015 20:57

PROTEIN STABILITY AND LIFETIME

Protein stability is often used synonymously with protein lifetime and refers to how long a protein is present in a cell. Protein lifetime is often calculated as protein half-life (i.e., the time it takes for 50% of the existing protein population to be degraded). Alternatively, protein stability can refer to the protein’s capacity to remain in a stable and functional form (i.e., not modiﬁed with negative consequences for function, damaged, or aggregated).

multiple additional adaptors could be present in the organelles. A key objective for organellar protease research should thus be to identify degrons, speciﬁc adaptors, and underlying molecular recognition mechanisms.

The N-End Rule and N-Degrons Surprisingly little is known about what determines plant organelle protein lifetime and stability, even though this is of critical importance in proteostasis (see also sidebar Protein Stability and Life- time). The N termini of proteins are major determinants of protein stability/half-life in bacteria (175), eukaryotes (50), mitochondria, and likely plastids/chloroplasts (5, 121). Early observations led to the formulation of the N-end rule, which states that certain amino acids, when exposed at the N terminus of a protein, act as triggers for degradation (9). The N-end rule was initially based on observations of proteins in yeast and other eukaryotes, in particular for proteins degraded by the proteasome following polyubiquitination. The N-end rule in prokaryotes is different from that in eukaryotes in part because prokaryotes do not have a proteasome and lack ubiquitination (168). Recent reviews have summarized some of the history of these discoveries and the present understanding of the N-end rule pathway in prokaryotes and eukaryotes (37, 45, 165). Mitochondria and chloroplasts/plastids originally derived from prokaryotic endosymbionts, suggesting that prokaryotic N-end rules might exist in these organelles. But because so many of the original bacterial genes were lost or transferred to the nuclear genome (51, 75) and many proteins in mitochondria and plastids lack clear bacterial homologs, it is highly likely that these original prokaryotic N-end rules changed during the evolution of plants. Although an N-end rule for chloroplasts/plastids in plants is not known, overexpression studies of plastid-targeted proteins have shown that the amino acids at the N terminus can greatly affect protein half-life (5; for discussion, see 44). Moreover, the presence of chloroplast CLPS suggests the presence of an N-degron (121). Because of the lack of experimental observations, an N-end rule has not been determined for plant mitochondria, but one is quite well deﬁned for mitochondrial protein stability in yeast (113, 177, 179, 180). Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org

Posttranslational Modiﬁcations Can Affect Proteolysis Proteins can undergo many different PTMs that can potentially inﬂuence protein stability, either by leading to conformational changes that expose internal regions acting as degrons or by functioning as recognition signals for substrate selection and degradation. Oxidative stress can result in several types of PTMs, such as carbonylation and methionine sulfoxide formation. Ox-

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. idative carbonylation has attracted much attention because of its irreversible nature; carbonylated proteins are marked for proteolysis by the proteasome (outside organelles) and the LON protease (within mitochondria) or form aggregates that can accumulate with age but can be removed by the combined action of chaperones and AAA+ proteases (103, 123, 158). Carbonylation has been studied mainly in nonplant mitochondria but also occurs on a large scale in Arabidopsis,including in chloroplasts (70) and other organelles (140). Little information is available about the

96 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

proteolytic removal of carbonylated proteins in plant organelles, and this warrants future attention, in particular because oxidative stress is so common on plants. Phosphorylation is a highly dynamic, reversible PTM that has been extensively studied in plants; a meta-analysis for Arabidopsis identiﬁed more than 60,000 phosphopeptides matching to >8,000 proteins (174). The stability of several proteins (DELLA, PIF3, and UPS1) in Arabidopsis is regulated by (de)phosphorylation, but these are present outside of the organelles, and their degradation likely involves ubiquitination and the proteasome. Within the three organelles, the phosphorylation state of the D1 protein of photosystem II indirectly affects its degradation through the photosystem II disassembly process and thus the exposure of the D1 proteins to thylakoid proteases (74). Molecular support for protein degradation directly induced by phosphorylation remains to be demonstrated within the organelles. Proteins also undergo other PTMs, such as N-terminal or lysine acetylation, which may serve to stabilize or activate mature proteins. For instance, Hoshiyasu et al. (56) suggested that N- terminal acetylation of the ε subunit of the thylakoid ATP synthase stabilizes this protein during drought stress. Extensive (reversible) lysine acetylation was recently described for plant mitochondrial proteins (88, 159), but unlike in chloroplasts (197), irreversible N-terminal acetylation is absent in mitochondria (59). Postulated functions of N-terminal acetylation include protein localization and protein stability but are generally not very clear. In contrast, lysine acetylation in mitochondria and peroxisomes is driven by acetyl–coenzyme A (acetyl-CoA) produced within these organelles and likely functions in enzymatic regulation (89). Gibellini et al. (46) observed an interesting connection between lysine (de)acetylation and LON proteases in human mitochondria: A mitochondrial lysine deacetylase (sirtuin 3) could deacetylate LON, thereby perhaps enabling LON activity. However, direct evidence for a role for protein acetylation in protein degradation is lacking for plants. N-terminal enrichment and degradomics techniques may resolve this question (see Degradomics: Systems Analysis of Proteolysis and Protein Lifetime, below).

INVOLVEMENT OF UBIQUITINATION AND THE PROTEASOME IN ORGANELLE BIOGENESIS AND DIFFERENTIATION Several studies have demonstrated that the proteasome can degrade protein precursor proteins (including chloroplast proteases CLPP4 and FTSH1) en route to plastids following ubiquitination in the cytosol (94, 148, 154, 155). This involves the cytosolic E3 ubiquitin ligase AtCHIP recognizing cTPs and the 26S proteasome, and perhaps also involves N-terminal acetylation of the precursor protein (for discussion, see 92). Unlike nuclear and peroxisomal proteins, mitochondrial and chloroplast precursor proteins must be unfolded before or during organellar

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org import, which makes them more susceptible to aggregation and hence suggests the need for degradation (92). Such extraorganellar ubiquitination and proteasome-dependent degradation could also provide a mechanism to avoid overloading of the organellar protein import apparatus and subsequent accumulation of superfluous proteins. Additionally, ubiquitination of specific outer-membrane components of the TOC chloroplast protein import apparatus alters the composition of the TOC machinery, thereby facilitating plastid differentiation. Specifically, Ling et al. (100) identified the RING-type ubiquitin E3 ligase SP1 in the plastid outer envelope

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. membrane, where it mediates ubiquitination of TOC components and their degradation by the cytosolic ubiquitin-proteasome system (UPS). The mitochondrial outer-membrane-bound ubiquitin protease UBP27, which cleaves ubiquitins from proteins (a deubiquitination enzyme), plays a role in mitochondrial morphology, possibly by modulating the function of mitochondrial division proteins (130). Thus, although there is no ubiquitination within the three organelles, proteasome activity does directly impact their proteome composition.

www.annualreviews.org • Plant Organelle Proteolysis 97 PP66CH04-VanWijk ARI 30 March 2015 20:57

ORGANELLE PROTEOME TURNOVER THROUGH AUTOPHAGY AND TARGETED VESICLE TRANSPORT Autophagy has recently emerged as a crucial homeostatic system in plants that represents an alternative to the UPS for removing unwanted intracellular proteins, protein complexes, or whole organelles (7, 98, 105), including glyoxysomes/peroxisomes (pexophagy) (93), mitochondria (mitophagy) (97), and chloroplasts (chlorophagy) (62, 64, 181). During autophagy, portions of the cytoplasm, including whole organelles, are enclosed by de novo–formed vesicles composed of a double membrane, termed autophagosomes. After transport to the vacuole, the outer membrane of the autophagosome fuses with the tonoplast to release a single-membrane autophagic body that is subsequently degraded in the vacuole. The genes required for the autophagic process are named ATG and are largely conserved between plants and nonphotosynthetic eukaryotes (98). Re- cruitment proteins (ATG8-interacting receptors) have been discovered that tether speciﬁc cargo to enveloping autophagy vesicles, allowing autophagy to be a highly selective process, similar to the UPS. Below, I summarize key ﬁndings for autophagy of the three organelles and functional relationships to intraorganellar proteolysis.

Pexophagy Just over the last two years, ﬁve studies have clearly demonstrated that autophagy of plant peroxisomes is a key process in removing damaged/oxidized peroxisomes (156, 192) and removing glyoxysomes (specialized peroxisomes) in cotyledons once stored lipids are metabolized and seedlings have become photosynthetic (41, 49, 76; reviewed in 8, 93). One of the major func-

tions of peroxisomes is the detoxiﬁcation of hydrogen peroxide (H2O2) through catalase activity as part of the photorespiratory pathway in photosynthetic leaves as well as during β-oxidation in

young seedlings. However catalase itself can be inactivated by H2O2 and can result in accumulation of protein aggregates inside peroxisomes, which require autophagy for removal (156, 192). Importantly, the autophagic removal of glyoxysomes is complemented by additional degradation processes, namely interglyoxysomal proteolysis by LON2 (see ATP-Dependent Proteases of the AAA+ Family: LON, CLP, and FTSH, above) and the UPS at the outward-facing periphery site of the peroxisome (19, 41, 101). The UPS requires the activity of the ubiquitin-conjugating enzyme PEX4 and other PEX proteins involved in peroxisomal protein export and degradation by the proteasome (101). Interestingly, because the peroxisomal matrix protease LON2 plays dual roles as an ATP-dependent protease and a chaperone essential for suppressing autophagy, LON2 deﬁciency causes enhanced pexophagy as well as an accumulation of peroxisomal proteins in the

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org cytosol resulting from a decrease in the number of peroxisomes. Thus, the intermolecular modulation between the protease and chaperone functions of LON2 regulates autophagy (49, 193). Furthermore, during senescence, leaf peroxisomes are converted to glyoxysomes (54), but whether functional transitions also involve both LON2 and pexophagy is unclear.

Mitophagy

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Whereas mitophagy has been extensively demonstrated in nonphotosynthetic eukaryotes (20, 48), it was not until 2014 that compelling genetic evidence for mitophagy was available for plants. Vier- stra and colleagues (97) showed that whole Arabidopsis mitochondria or mitochondrial fragments created by ﬁssion are degraded during senescence via an autophagic route that requires ATG11 and other ATG components. However, how the autophagic machinery recognizes mitochondria (e.g., by interaction of mitochondrial surface proteins with ATG11) and how this recognition is

98 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

regulated are not known. Furthermore, the relation between mitophagy and intramitochondrial proteases has not been explored.

Protein turnover: the Chlorophagy and Targeted Chloroplast Protein Removal by net balance between protein synthesis and Rubisco-Containing Bodies and Senescence-Associated Vacuoles protein degradation; Leaves make enormous investments in the photosynthetic machinery located in chloroplasts, an understanding of with a high percentage just in stromal-localized Rubisco. It is therefore not surprising that the protein turnover will include determination chloroplast is a target for nutrient recycling during nutrient or carbon starvation as well as during of the rates of protein leaf senescence. Recently, genetic evidence has shown that autophagy plays a role in the turnover of synthesis and protein whole chloroplasts (ATG4 dependent) as well as in the selective removal of Rubisco (and a few other degradation stromal proteins) through vesicles called Rubisco-containing bodies (ATG4 and -5 dependent). Most of the work demonstrating the involvement of ATG genes was carried out by Ishida and colleagues (e.g., 63, 127, 181; for details and an excellent review, see 62). In addition, during leaf senescence, chloroplast proteins, in particular Rubisco and glutamate synthase, relocalize to senescence-associated vacuoles, which are small, single-membrane acidic vesicles (or vacuoles) with high cysteine peptidase activity, likely from SAG12 (23, 109, 129). This latter pathway appears to be independent of direct involvement of autophagy genes, and the functional relation to Rubisco-containing bodies is unclear. In conclusion, most evidence for chloroplast protein turnover during senescence points at chlorophagy, Rubisco-containing bodies, and senescence-associated vacuoles. However, particularly during the onset of senescence, intrachloroplast proteases likely play important roles in controlled dismantling of the chloroplast photosynthetic apparatus and thylakoid proteome. Sur- prisingly, little is known about these intrachloroplast proteolysis events (see 6). Whereas the chlorophagy described above is most likely macroautophagy, microautophagy appears to con- tribute to diurnal starch degradation (185). However, whether this contributes to chloroplast proteome dynamics is not clear. Degradation of amyloplasts in columella cells, GFP-ATG8a accumulation patterns, and observation of autophagosome-like structures in root tips and root expression patterns of ATG18a in response to water stress or hydrotropic stimulation have suggested that nonphotosynthetic plastids can also undergo autophagy (114).

ORGANELLAR PROTEOLYSIS IN CROP PLANTS, BIOTECHNOLOGY, AND SYNTHETIC BIOLOGY So far, Arabidopsis has been the most exploited species for the discovery and characterization of

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org plant proteases and proteostasis as well as the study of autophagy. However, there is a growing recognition of the importance of these processes in crop plants in terms of maximizing biomass and yield. Indeed, in the last few years, studies of organellar proteolysis using loss-of-function mutants have been published for the CLP system in maize (191) and rice (36, 99), and map-based cloning of a tomato fruit-ripening mutant identiﬁed a homolog of Arabidopsis chloroplast EGY1 (12). Other studies have shown that the chloroplast CGEP protease differentially accumulates between

bundle sheath and mesophyll chloroplasts, suggesting a role in C4 chloroplast differentiation and

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. proteostasis (43, 107; N.H. Bhuiyan & K.J. van Wijk, unpublished data). Plants are an excellent system for producing proteins with nutritional (38) or pharmaceuti- cal value, in particular vaccines and antisera (133) such as the experimental vaccine against the Ebola virus, which is generated in tobacco leaves. These applications are grouped under the term molecular farming, and chloroplasts are the favored subcellular compartment for overexpressing such products (44, 108). However, some of these proteins (such as rotavirus VP6 protein) are

www.annualreviews.org • Plant Organelle Proteolysis 99 PP66CH04-VanWijk ARI 30 March 2015 20:57

insufﬁciently stable, thereby severely limiting protein accumulation, and high accumulation levels of other proteins can be toxic for the plants (24). Understanding the molecular details of protease substrate selection and degrons is thus essential to maximize the success of molecular farming. Chloroplast biotechnology and synthetic biology could also aid in the accumulation of high levels of a variety of cell wall–degrading enzymes in plastids, which may help to overcome bottlenecks in the production of biofuels from lignocellulosic biomass (178, 195). Understanding proteolysis and protease recognition signals in plastids will be critical for optimizing transgenic protein accumulation levels.

DEGRADOMICS: SYSTEMS ANALYSIS OF PROTEOLYSIS AND PROTEIN LIFETIME To understand the role and substrates of speciﬁc proteases and to determine protein lifetimes, large-scale techniques are required that can track hundreds of proteins and potential degradation products in vivo. Indeed, there is great interest in developing such large-scale systems biology techniques (3, 60, 82) (Table 3). It is now possible to identify the N termini of hundreds of proteins in a single sample using mass spectrometry combined with speciﬁc protein enrichment strategies (e.g., 91, 135, 161, 189). It is also now possible to make large-scale estimates of protein degradation rates or net protein lifetime (the difference between protein synthesis and degradation) using (pulsed) metabolic labeling with stable isotopes supplied as amino acids [in particular for single-cell systems (33, 60, 79)] or with 15N/14N for whole-plant hydroponic labeling followed by mass spectrometry and mathematical modeling. However, these methods are far from accurate, in part because of the complexities of plant amino acid metabolism (117, 119, 153). In the following sections, I explain the different large-scale techniques (see Table 3).

Table 3 Overview of degradomics techniques and their applications (for more details and discussion, see 91, 135) Technique Application Examples Positional Identifying neo–N termini, neo–C Selection of N-terminal peptides (e.g., TAILS, COFRADIC) proteomics termini, and cleavage sites Selection of C-terminal peptides (e.g., C-TAILS) Metabolic Measuring differentials between Measurement of net changes in a stable isotope label by SILAC (13C-Arg, protein protein synthesis and degradation 13C-Lys, other 13C-labeled amino acid); a variant is pulsed SILAC labeling Measurement of net changes in a stable isotope label by 14N/15N labeling through inorganic nitrogen sources (ammonium and nitrate salts) Comparative Comparing (steady-state) protein 2DE gel followed by image-based quantification of stained gel and MS-based Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org quantitative levels and observing protein protein identification proteomics fragments MS-based 1DE gel profiling (looking for protein fragments) (PROTOMAP) MS-based label-free proteomics (spectral counting, peak intensity) MS-based stable isotope–labeled proteomics (metabolic label or postharvest label) (e.g., iTRAQ, ICAT) MS-based absolute quantification (QconCat, AQUA) MS-based targeted proteomics using MRM or SRM (quantification of a

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. preselected set of proteins)

Abbreviations: 1DE, one-dimensional electrophoresis; 2DE, two-dimensional electrophoresis; AQUA, absolute quantitation; COFRADIC, combined fractional diagonal chromatography; C-TAILS, C-terminal amino isotopic labeling of substrates; ICAT, isotope-coded afﬁnity tag; iTRAQ, isobaric tags for relative and absolute quantitation; MRM, multiple reaction monitoring; MS, mass spectrometry; PROTOMAP, protein topography and migration analysis platform; SILAC, stable isotope labeling by amino acids in cell culture; SRM, selected reaction monitoring; TAILS, terminal amino isotopic labeling of substrates.

100 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

Identification of Cleavage Sites and Degradation Products Through Large-Scale N- or C-Terminal Labeling and Enrichment To identify protein degradation products in vivo or in vitro, it is highly beneficial to identify the N or C termini of such fragments with high certainty in complex proteomes. Typical proteomics workflows generally yield only partial coverage of protein sequences, and it is often difficult to know which peptides represent the true N or C termini. Therefore, systematic identification of N or C termini requires specific labeling and enrichment strategies. Identification and recognition of N termini of nucleus-encoded proteins accumulating within organelles such as chloroplasts and mitochondria are challenging because they undergo N-terminal processing (Figure 1). To directly identify and recognize N termini in complex protein mixtures, two strategies in particular have been applied: combined fractional diagonal chromatography (COFRADIC), used by Gevaert and colleagues (161), and terminal amino isotopic labeling of substrates (TAILS), used by Overall and colleagues (80, 91). Both techniques can also be applied to C termini, but most studies have focused on N termini to identify degradation products (Table 3).

Comparative Proteomics of Protease Mutants Changes in the steady-state proteome in Arabidopsis protease loss-of-function mutants can reflect accumulation of nondegraded protease substrates if these substrates are specifically degraded by this protease. Such changes also reflect downstream or pleiotropic effects, for instance, when loss of protease activity blocks specific essential functions. Analysis of proteome samples at different time points in the life cycles of protease mutants can further narrow down when, where, and in which pathways or processes a protease is functionally most relevant. Two techniques have been employed for comparative proteomics of organellar protease mutants (LON1 and DEG14): two-dimensional electrophoresis (2DE) (14, 160) and label-free quantification using a high-resolution mass spectrometer (LTQ Orbitrap) and spectral counting (SPC) (77, 78, 121, 196). SPC has been applied to loss-of-function mutants for different components of the chloroplast CLP protease system, namely the core subunits CLPR2 (196), CLPR4 (78), and CLPP3 (77); CLPS; and CLPC1 (121), as well as to a double mutant for CLPT1 and CLPT2 ( J. Kim, G. Friso & K.J. van Wijk, manuscript in preparation). This technique allowed the identification of 2,000–2,500 proteins and the quantification (with statistical significance analysis) of several hundred proteins. I should emphasize that although these SPC-based studies allowed large-scale quantification of changes in proteome composition, likely only a subset of the upregulated proteins are protease substrates. Indeed, affinity purification identified some of these

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org upregulated proteins as direct interactors with CLPS1 (121). Together, the above studies show that comparative methods can provide insight into protease functions, but complementary methods are required to truly identify substrates. Combining comparative proteomics with the use of pharmacological reagents (e.g., to block translation, speciﬁc PTMs, or proteases) or inducible repression of speciﬁc proteases in vivo (e.g., by inducible RNA interference) appears to be a promising approach for identifying substrates. Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Protein Lifetime Analysis Using Stable Isotopes Whereas comparative proteomics tracks steady-state proteomes, an alternative and complementary method is to determine (or rather estimate) protein lifetime (calculated as half-life) rates (119). Application of such measurements to protease mutants could identify potential substrates (recognized by extended lifetime) but can still be confounded by pleiotropic effects, similar to the

www.annualreviews.org • Plant Organelle Proteolysis 101 PP66CH04-VanWijk ARI 30 March 2015 20:57

steady-state proteome analysis described above. Before the development of protein mass spectrometry, protein lifetime measurements were typically done by pulse and chase radioactive isotope labeling (e.g., 35S-methione), often in combination with translation inhibitors. The protein lifetime is measured by quantification of the loss of radiolabeled protein (e.g., by immunoprecipitation). Although this is still a useful technique, it is limited to analysis of smaller numbers of predefined proteins and cannot be applied at the (untargeted) whole-proteome scale. Furthermore, these ra- dioisotopes are not compatible with mass spectrometry measurements because of safety concerns, and they therefore require specific antisera for detection/identification. For a global analysis of possible protein substrates, mass spectrometry in combination with stable isotopes can be applied. The most commonly used stable isotope for whole plants is 15N, supplied as inorganic nitrogen (ammonium and nitrate salts) (for discussion, see 119). To determine protein lifetimes, the complete proteome (from seed) should first be labeled with an isotope label, aiming for >98% labeling of the relevant amino acid(s). The cells or organisms are then transferred to a medium containing a different isotope label, and the change in ratio between the two labeled forms (peptide) for each protein is determined. Unfortunately, rather than directly measuring protein degradation, this method measures the net differences between protein synthesis and degradation, and assumptions have to be made regarding synthesis rates. These rates differ among proteins and can also greatly vary for each protein depending on the developmental state and external signals. Furthermore, this method suffers from a delayed response (a lag of many hours) because of the sizable amino acid pools (reviewed in 118). This approach could be suitable for identifying candidate protease substrates by differential protein lifetimes when comparing wild-type and protease mutants.

SUMMARY POINTS 1. Chloroplasts (plastids), mitochondria, and peroxisomes have commonalties in their proteolytic systems that should be further explored to improve understanding of protease- substrate relationships and the coordination of organellar function. 2. DEG14 (located in the mitochondria) and FTSH11 (located in mitochondria and chloroplasts) confer thermotolerance in Arabidopsis, likely by degrading misfolded proteins. No other organellar proteases are (yet) known to confer thermotolerance. 3. Different protease systems have developed unique ways to prevent unwanted proteolysis. These occur through the sequestration of active sites within oligomeric structures, selective activation by changes in oligomeric state, and speciﬁc substrate delivery systems

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org and adaptors that direct substrates to the proteolytic sites. 4. There is a fascinating interplay between intraorganellar proteolysis, the proteasome, and autophagy. These three processes are known to act in a complementary fashion, but much remains to be discovered about their functions and interactions. 5. Organellar proteolysis and autophagy have been studied mainly in Arabidopsis, but several recent studies have illustrated the importance of organellar proteases and autophagy in

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. crop plants. These ﬁndings could potentially have a large impact on agriculture. 6. Recent development in proteomics and degradomics techniques can and need to be applied to identify protease substrate and cleavage sites.

102 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

DISCLOSURE STATEMENT The author is not aware of any afﬁliations, memberships, funding, or ﬁnancial holdings that might be perceived as affecting the objectivity of this review.

ACKNOWLEDGMENTS Research on chloroplasts and proteases in my lab has been generously supported by the National Science Foundation through grants MCB-1021963, IOS-1127017, and IOS-0922560. I thank members of my lab for critically reading this review and for valuable suggestions.

LITERATURE CITED 1. Adam Z. 2013 . Emerging roles for diverse intramembrane proteases in plant biology. Biochim. Biophys. Acta 1828:2933–36 2. Adrain C, Freeman M. 2012. New lives for old: evolution of pseudoenzyme function illustrated by iRhoms. Nat. Rev. Mol. Cell Biol. 13:489–98 3. Agard NJ, Wells JA. 2009. Methods for the proteomic identification of protease substrates. Curr. Opin. Chem. Biol. 13:503–9 4. Albrecht-Borth V, Kauss D, Fan D, Hu Y, Collinge D, et al. 2013. A novel proteinase, SNOWY COTYLEDON4, is required for photosynthetic acclimation to higher light intensities in Arabidop- sis. Plant Physiol. 163:732–45 5. Apel W, Schulze WX, Bock R. 2010. Identification of protein stability determinants in chloroplasts. Plant J. 63:636–50 6. Avice JC, Etienne P. 2014. Leaf senescence and nitrogen remobilization efficiency in oilseed rape (Brassica napus L.). J. Exp. Bot. 65:3813–24 7. Avila-Ospina L, Moison M, Yoshimoto K, Masclaux-Daubresse C. 2014. Autophagy, plant senescence, and nutrient recycling. J. Exp. Bot. 65:3799–811 8. Avin-Wittenberg T, Fernie AR. 2014. At long last: evidence for pexophagy in plants. Mol. Plant 7:1257– 60 9. Bachmair A, Finley D, Varshavsky A. 1986. In vivo half-life of a protein is a function of its amino-terminal residue. Science 234:179–86 10. Baier A, Winkler W, Korte T, Lockau W, Karradt A. 2014. Degradation of phycobilisomes in Synechocystis sp. PCC6803: evidence for essential formation of an NblA1/NblA2 heterodimer and its codegradation by a Clp protease complex. J. Biol. Chem. 289:11755–66 11. Barajas-Lopez´ JDD, Blanco NE, Strand A. 2013. Plastid-to-nucleus communication, signals controlling the running of the plant cell. Biochim. Biophys. Acta 1833:425–37 12. Barry CS, Aldridge GM, Herzog G, Ma Q, McQuinn RP, et al. 2012. Altered chloroplast development and delayed fruit ripening caused by mutations in a zinc metalloprotease at the lutescent2 locus of tomato. Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org Plant Physiol. 159:1086–98 13. Bartel B, Farmer LM, Rinaldi MA, Young PG, Danan CH, Burkhart SE. 2014. Mutation of the Arabidopsis LON2 peroxisomal protease enhances pexophagy. Autophagy 10:518–19 14. Basak I, Pal R, Patil KS, Dunne A, Ho HP, et al. 2014. Arabidopsis AtPARK13, which confers thermotolerance, targets misfolded proteins. J. Biol. Chem. 289:14458–69 15. Baudisch B, Klosgen RB. 2012. Dual targeting of a processing peptidase into both endosymbiotic organelles mediated by a transport signal of unusual architecture. Mol. Plant 5:494–503

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. 16. Bauwe H, Hagemann M, Fernie AR. 2010. Photorespiration: players, partners and origin. Trends Plant Sci. 15:330–36 17. Bolter B, Nada A, Fulgosi H, Soll J. 2006. A chloroplastic inner envelope membrane protease is essential for plant development. FEBS Lett. 580:789–94 18. Bruch EM, Rosano GL, Ceccarelli EA. 2012. Chloroplastic Hsp100 chaperones ClpC2 and ClpD interact in vitro with a transit peptide only when it is located at the N-terminus of a protein. BMC Plant Biol. 12:57

www.annualreviews.org • Plant Organelle Proteolysis 103 PP66CH04-VanWijk ARI 30 March 2015 20:57

19. Burkhart SE, Lingard MJ, Bartel B. 2013. Genetic dissection of peroxisome-associated matrix protein degradation in Arabidopsis thaliana. Genetics 193:125–41 20. Campello S, Strappazzon F, Cecconi F. 2014. Mitochondrial dismissal in mammals, from protein degradation to mitophagy. Biochim. Biophys. Acta 1837:451–60 21. Carrie C, Murcha MW, Giraud E, Ng S, Zhang MF, et al. 2013. How do plants make mitochondria? Planta 237:429–39 22. Carrie C, Small I. 2013. A reevaluation of dual-targeting of proteins to mitochondria and chloroplasts. Biochim. Biophys. Acta 1833:253–59 23. Carrion CA, Costa ML, Martinez DE, Mohr C, Humbeck K, Guiamet JJ. 2013. In vivo inhibition of cysteine proteases provides evidence for the involvement of “senescence-associated vacuoles” in chloroplast protein degradation during dark-induced senescence of tobacco leaves. J. Exp. Bot. 64:4967–80 24. Chakrabarti SK, Lutz KA, Lertwiriyawong B, Svab Z, Maliga P. 2006. Expression of the cry9Aa2 B.t. gene in tobacco chloroplasts confers resistance to potato tuber moth. Transgenic Res. 15:481–88 25. Che Y, Fu A, Hou X, McDonald K, Buchanan BB, et al. 2013. C-terminal processing of reaction center protein D1 is essential for the function and assembly of photosystem II in Arabidopsis. PNAS 110:16247– 52 26. Chen G, Bi YR, Li N. 2005. EGY1 encodes a membrane-associated and ATP-independent metalloprotease that is required for chloroplast development. Plant J. 41:364–75 27. Chen G, Law K, Ho P, Zhang X, Li N. 2012. EGY2, a chloroplast membrane metalloprotease, plays a role in hypocotyl elongation in Arabidopsis. Mol. Biol. Rep. 39:2147–55 28. Chen J, Burke JJ, Velten J, Xin Z. 2006. FtsH11 protease plays a critical role in Arabidopsis thermotolerance. Plant J. 48:73–84 29. Chi W, Sun X, Zhang L. 2012. The roles of chloroplast proteases in the biogenesis and maintenance of photosystem II. Biochim. Biophys. Acta 1817:239–46 30. Chu CC, Li HM. 2012. The amino-terminal domain of chloroplast Hsp93 is important for its membrane association and functions in vivo. Plant Physiol. 158:1656–65 31. Chung T, Suttangkakul A, Vierstra RD. 2009. The ATG autophagic conjugation system in maize: ATG transcripts and abundance of the ATG8-lipid adduct are regulated by development and nutrient availability. Plant Physiol. 149:220–34 32. Clarke AK. 2012. The chloroplast ATP-dependent Clp protease in vascular plants—new dimensions and future challenges. Physiol. Plant. 145:235–44 33. Claydon AJ, Beynon R. 2012. Proteome dynamics: revisiting turnover with a global perspective. Mol. Cell. Proteomics 11:1551–65 34. Daras G, Rigas S, Tsitsekian D, Zur H, Tuller T, Hatzopoulos P. 2014. Alternative transcription initi- ation and the AUG context conﬁguration control dual-organellar targeting and functional competence of Arabidopsis Lon1 protease. Mol. Plant 7:989–1005 35. Dolze E, Chigri F, Howing T, Hierl G, Isono E, et al. 2013. Calmodulin-like protein AtCML3 medi-

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org ates dimerization of peroxisomal processing protease AtDEG15 and contributes to normal peroxisome metabolism. Plant Mol. Biol. 83:607–24 36. Dong H, Fei GL, Wu CY, Wu FQ, Sun YY, et al. 2013. A rice virescent-yellow leaf mutant reveals new insights into the role and assembly of plastid caseinolytic protease in higher plants. Plant Physiol. 162:1867–80 37. Dougan DA, Micevski D, Truscott KN. 2012. The N-end rule pathway: from recognition by N- recognins, to destruction by AAA+ proteases. Biochim. Biophys. Acta 1823:83–91 38. Enﬁssi EM, Fraser PD, Lois LM, Boronat A, Schuch W, Bramley PM. 2005. Metabolic engineering of Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. the mevalonate and non-mevalonate isopentenyl diphosphate-forming pathways for the production of health-promoting isoprenoids in tomato. Plant Biotechnol. J. 3:17–27 39. Erbse A, Schmidt R, Bornemann T, Schneider-Mergener J, Mogk A, et al. 2006. ClpS is an essential component of the N-end rule pathway in Escherichia coli. Nature 439:753–56 40. Estavillo GM, Chan KX, Phua SY, Pogson BJ. 2012. Reconsidering the nature and mode of action of metabolite retrograde signals from the chloroplast. Front. Plant Sci. 3:300

104 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

41. Farmer LM, Rinaldi MA, Young PG, Danan CH, Burkhart SE, Bartel B. 2013. Disrupting autophagy restores peroxisome function to an Arabidopsis lon2 mutant and reveals a role for the LON2 protease in peroxisomal matrix protein degradation. Plant Cell 25:4085–100 42. Feng J, Michalik S, Varming AN, Andersen JH, Albrecht D, et al. 2013. Trapping and proteomic identification of cellular substrates of the ClpP protease in Staphylococcus aureus. J. Proteome Res. 12:547– 58 43. Friso G, Majeran W, Huang M, Sun Q, van Wijk KJ. 2010. Reconstruction of metabolic pathways, protein expression, and homeostasis machineries across maize bundle sheath and mesophyll chloroplasts: large-scale quantitative proteomics using the first maize genome assembly. Plant Physiol. 152:1219–50 44. Gao M, Li Y, Xue X, Wang X, Long J. 2012. Stable plastid transformation for high-level recombinant protein expression: promises and challenges. J. Biomed. Biotechnol. 2012:158232 45. Gibbs DJ, Bacardit J, Bachmair A, Holdsworth MJ. 2014. The eukaryotic N-end rule pathway: conserved mechanisms and diverse functions. Trends Cell Biol. 24:603–11 46. Gibellini L, Pinti M, Beretti F, Pierri CL, Onofrio A, et al. 2014. Sirtuin 3 interacts with Lon protease and regulates its acetylation status. Mitochondrion 18:76–81 47. Giglione C, Fieulaine S, Meinnel T. 2009. Cotranslational processing mechanisms: towards a dynamic 3D model. Trends Biochem. Sci. 34:417–26 48. Gomes LC, Scorrano L. 2013. Mitochondrial morphology in mitophagy and macroautophagy. Biochim. Biophys. Acta 1833:205–12 49. Goto-Yamada S, Mano S, Nakamori C, Kondo M, Yamawaki R, et al. 2014. Chaperone and protease functions of LON protease 2 modulate the peroxisomal transition and degradation with autophagy. Plant Cell Physiol. 55:482–96 50. Graciet E, Walter F, Maoileidigh DO, Pollmann S, Meyerowitz EM, et al. 2009. The N-end rule pathway controls multiple functions during Arabidopsis shoot and leaf development. PNAS 106:13618–23 51. Greiner S, Bock R. 2013. Tuning a menage´ a` trois: co-evolution and co-adaptation of nuclear and organellar genomes in plants. BioEssays 35:354–65 52. Gur E. 2013. The Lon AAA+ protease. Subcell. Biochem. 66:35–51 53. Hansen G, Hilgenfeld R. 2013. Architecture and regulation of HtrA-family proteins involved in protein quality control and stress response. Cell. Mol. Life Sci. 70:761–75 54. Hayashi M, Toriyama K, Kondo M, Kato A, Mano S, et al. 2000. Functional transformation of plant peroxisomes. Cell Biochem. Biophys. 32:295–304 55. Helm M, Luck C, Prestele J, Hierl G, Huesgen PF, et al. 2007. Dual specificities of the glyoxysomal/ peroxisomal processing protease Deg15 in higher plants. PNAS 104:11501–6 56. Hoshiyasu S, Kohzuma K, Yoshida K, Fujiwara M, Fukao Y, et al. 2013. Potential involvement of N- terminal acetylation in the quantitative regulation of the epsilon subunit of chloroplast ATP synthase under drought stress. Biosci. Biotechnol. Biochem. 77:998–1007 57. Hsu SC, Endow JK, Ruppel NJ, Roston RL, Baldwin AJ, Inoue K. 2011. Functional diversification of thylakoidal processing peptidases in Arabidopsis thaliana. PLOS ONE 6:e27258

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org 58. Hu J, Baker A, Bartel B, Linka N, Mullen RT, et al. 2012. Plant peroxisomes: biogenesis and function. Plant Cell 24:2279–303 59. Huang S, Taylor NL, Whelan J, Millar AH. 2009. Refining the definition of plant mitochondrial presequences through analysis of sorting signals, N-terminal modifications, and cleavage motifs. Plant Physiol. 150:1272–85 60. Hughes C, Krijgsveld J. 2012. Developments in quantitative mass spectrometry for the analysis of proteome dynamics. Trends Biotechnol. 30:668–76 61. Humbard MA, Surkov S, De Donatis GM, Jenkins LM, Maurizi MR. 2013. The N-degradome of Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Escherichia coli: Limited proteolysis in vivo generates a large pool of proteins bearing N-degrons. J. Biol. Chem. 288:28913–24 62. Ishida H, Izumi M, Wada S, Makino A. 2014. Roles of autophagy in chloroplast recycling. Biochim. Biophys. Acta 1837:512–21 63. Ishida H, Yoshimoto K. 2008. Chloroplasts are partially mobilized to the vacuole by autophagy. Autophagy 4:961–62

www.annualreviews.org • Plant Organelle Proteolysis 105 PP66CH04-VanWijk ARI 30 March 2015 20:57

64. Ishida H, Yoshimoto K, Izumi M, Reisen D, Yano Y, et al. 2008. Mobilization of Rubisco and stroma- localized fluorescent proteins of chloroplasts to the vacuole by an ATG gene-dependent autophagic process. Plant Physiol. 148:142–55 65. Jain R, Chan MK. 2007. Support for a potential role of E. coli oligopeptidase A in protein degradation. Biochem. Biophys. Res. Commun. 359:486–90 66. Janska H, Kwasniak M, Szczepanowska J. 2012. Protein quality control in organelles—AAA/FtsH story. Biochim. Biophys. Acta 1833:381–87 67. Janska H, Piechota J, Kwasniak M. 2012. ATP-dependent proteases in biogenesis and maintenance of plant mitochondria. Biochim. Biophys. Acta 1797:1071–75 68. Jarvis P, Lopez-Juez E. 2013. Biogenesis and homeostasis of chloroplasts and other plastids. Nat. Rev. Mol. Cell Biol. 14:787–802 69. Jeyaraju DV, Sood A, Laforce-Lavoie A, Pellegrini L. 2013. Rhomboid proteases in mitochondria and plastids: keeping organelles in shape. Biochim. Biophys. Acta 1833:371–80 70. Johansson E, Olsson O, Nystrom T. 2004. Progression and specificity of protein oxidation in the life cycle of Arabidopsis thaliana. J. Biol. Chem. 279:22204–8 71. Kadirjan-Kalbach DK, Yoder DW, Ruckle ME, Larkin RM, Osteryoung KW. 2012. FtsHi1/ARC1 is an essential gene in Arabidopsis that links chloroplast biogenesis and division. Plant J. 72:856–67 72. Kapri-Pardes E, Naveh L, Adam Z. 2007. The thylakoid lumen protease Deg1 is involved in the repair of photosystem II from photoinhibition in Arabidopsis. Plant Cell 19:1039–47 73. Kato Y, Sakamoto W. 2010. New insights into the types and function of proteases in plastids. Int. Rev. CellMol.Biol. 280:185–218 74. Kato Y, Sakamoto W. 2014. Phosphorylation of photosystem II core proteins prevents undesirable cleavage of D1 and contributes to the fine-tuned repair of photosystem II. Plant J. 79:312–21 75. Keeling PJ. 2013. The number, speed, and impact of plastid endosymbioses in eukaryotic evolution. Annu. Rev. Plant Biol. 64:583–607 76. Kim J, Lee H, Lee HN, Kim SH, Shin KD, Chung T. 2013. Autophagy-related proteins are required for degradation of peroxisomes in Arabidopsis hypocotyls during seedling growth. Plant Cell 25:4956–66 77. Kim J, Olinares PD, Oh SH, Ghisaura S, Poliakov A, et al. 2013. Modified Clp protease complex in the ClpP3 null mutant and consequences for chloroplast development and function in Arabidopsis. Plant Physiol. 162:157–79 78. Kim J, Rudella A, Ramirez Rodriguez V, Zybailov B, Olinares PD, van Wijk KJ. 2009. Subunits of the plastid ClpPR protease complex have differential contributions to embryogenesis, plastid biogenesis, and plant development in Arabidopsis. Plant Cell 21:1669–92 79. Kirchner M, Selbach M. 2012. In vivo quantitative proteome profiling: planning and evaluation of SILAC experiments. Methods Mol. Biol. 893:175–99 80. Kleifeld O, Doucet A, Prudova A, auf dem Keller U, Gioia M, et al. 2011. Identifying and quantifying proteolytic events and the natural N terminome by terminal amine isotopic labeling of substrates. Nat. Protoc. 6:1578–611

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org 81. Kley J, Schmidt B, Boyanov B, Stolt-Bergner PC, Kirk R, et al. 2011. Structural adaptation of the plant protease Deg1 to repair photosystem II during light exposure. Nat. Struct. Mol. Biol. 18:728–31 82. Klingler D, Hardt M. 2012. Proﬁling protease activities by dynamic proteomics workﬂows. Proteomics 12:587–96 83. Kmiec B, Glaser E. 2012. A novel mitochondrial and chloroplast peptidasome, PreP. Physiol. Plant. 145:180–86 84. Kmiec B, Teixeira PF, Berntsson RP, Murcha MW, Branca RM, et al. 2013. Organellar oligopeptidase (OOP) provides a complementary pathway for targeting peptide degradation in mitochondria and Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. chloroplasts. PNAS 110:E3761–69 85. Kmiec B, Teixeira PF, Glaser E. 2014. Phenotypical consequences of expressing the dually targeted Presequence Protease, AtPreP, exclusively in mitochondria. Biochimie 100:167–70 86. Knopf RR, Adam Z. 2012. Rhomboid proteases in plants—still in square one? Physiol. Plant. 145:41–51 87. Knopf RR, Feder A, Mayer K, Lin A, Rozenberg M, et al. 2012. Rhomboid proteins in the chloroplast envelope affect the level of allene oxide synthase in Arabidopsis thaliana. Plant J. 72:559–71

106 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

88. Konig¨ AC, Hartl M, Boersema PJ, Mann M, Finkemeier I. 2014. The mitochondrial lysine acetylome of Arabidopsis. Mitochondrion 19:252–60 89. Konig¨ AC, Hartl M, Pham PA, Laxa M, Boersema PJ, et al. 2014. The Arabidopsis class II sirtuin is a lysine deacetylase and interacts with mitochondrial energy metabolism. Plant Physiol. 164:1401–14 90. Kwasniak M, Pogorzelec L, Migdal I, Smakowska E, Janska H. 2012. Proteolytic system of plant mitochondria. Physiol. Plant. 145:187–95 91. Lange PF, Overall CM. 2013. Protein TAILS: when termini tell tales of proteolysis and function. Curr. Opin. Chem. Biol. 17:73–82 92. Lee DW, Jung C, Hwang I. 2013. Cytosolic events involved in chloroplast protein targeting. Biochim. Biophys. Acta 1833:245–52 93. Lee HN, Kim J, Chung T. 2014. Degradation of plant peroxisomes by autophagy. Front. Plant Sci. 5:139 94. Lee S, Lee DW, Lee Y, Mayer U, Stierhof YD, et al. 2009. Heat shock protein cognate 70-4 and an E3 ubiquitin ligase, CHIP, mediate plastid-destined precursor degradation through the ubiquitin-26S proteasome system in Arabidopsis. Plant Cell 21:3984–4001 95. Lensch M, Herrmann RG, Sokolenko A. 2001. Identification and characterization of SppA, a novel light- inducible chloroplast protease complex associated with thylakoid membranes. J. Biol. Chem. 276:33645– 51 96. Li B, Li Q, Xiong L, Kronzucker HJ, Kramer U, Shi W. 2012. Arabidopsis plastid AMOS1/EGY1 integrates abscisic acid signaling to regulate global gene expression response to ammonium stress. Plant Physiol. 160:2040–51 97. Li F, Chung T, Vierstra RD. 2014. AUTOPHAGY-RELATED11 plays a critical role in general autophagy- and senescence-induced mitophagy in Arabidopsis. Plant Cell 26:788–807 98. Li F, Vierstra RD. 2012. Autophagy: a multifaceted intracellular system for bulk and selective recycling. Trends Plant Sci. 17:526–37 99. Li W, Wu C, Hu G, Xing L, Qian W, et al. 2013. Characterization and fine mapping of a novel rice narrow leaf mutant nal9. J. Integr. Plant Biol. 55:1016–25 100. Ling Q, Huang W, Baldwin A, Jarvis P. 2012. Chloroplast biogenesis is regulated by direct action of the ubiquitin-proteasome system. Science 338:655–59 101. Lingard MJ, Bartel B. 2009. Arabidopsis LON2 is necessary for peroxisomal function and sustained matrix protein import. Plant Physiol. 151:1354–65 102. Lingner T, Kataya AR, Antonicelli GE, Benichou A, Nilssen K, et al. 2011. Identification of novel plant peroxisomal targeting signals by a combination of machine learning methods and in vivo subcellular targeting analyses. Plant Cell 23:1556–72 103. Lionaki E, Tavernarakis N. 2013. Oxidative stress and mitochondrial protein quality control in aging. J. Proteomics 92:181–94 104. Liu X, Yu F, Rodermel S. 2010. Arabidopsis chloroplast FtsH, var2 and suppressors of var2 leaf variegation: a review. J. Integr. Plant Biol. 52:750–61 105. Liu Y, Bassham DC. 2012. Autophagy: pathways for self-eating in plant cells. Annu. Rev. Plant Biol. 63:215–37 Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org 106. Lundquist P, Poliakov A, Bhuiyan NH, Zybailov B, Sun Q, van Wijk KJ. 2012. The functional network of the Arabidopsis thaliana plastoglobule proteome based on quantitative proteomics and genome-wide co-expression analysis. Plant Physiol. 58:1172–92 107. Majeran M, Friso G, Ponnala L, Connolly B, Huang M, et al. 2010. Structural and metabolic transitions of C4 leaf development and differentiation defined by microscopy and quantitative proteomics. Plant Cell 22:3509–42 108. Maliga P, Bock R. 2011. Plastid biotechnology: food, fuel, and medicine for the 21st century. Plant Physiol. 155:1501–10 Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. 109. Martinez DE, Costa ML, Gomez FM, Otegui MS, Guiamet JJ. 2008. “Senescence-associated vacuoles” are involved in the degradation of chloroplast proteins in tobacco leaves. Plant J. 56:196–206 110. Maruyama J, Yamaoka S, Matsuo I, Tsutsumi N, Kitamoto K. 2012. A newly discovered function of peroxisomes: involvement in biotin biosynthesis. Plant Signal. Behav. 7:1589–93 111. Meinnel T, Serero A, Giglione C. 2006. Impact of the N-terminal amino acid on targeted protein degradation. Biol. Chem. 387:839–51

www.annualreviews.org • Plant Organelle Proteolysis 107 PP66CH04-VanWijk ARI 30 March 2015 20:57

112. Mohanty A, McBride HM. 2013. Emerging roles of mitochondria in the evolution, biogenesis, and function of peroxisomes. Front. Physiol. 4:268 113. Mossmann D, Meisinger C, Vogtle FN. 2012. Processing of mitochondrial presequences. Biochim. Bio- phys. Acta 1819:1098–106 114. Nakamura M, Toyota M, Tasaka M, Morita MT. 2011. An Arabidopsis E3 ligase, SHOOT GRAVITROPISM9, modulates the interaction between statoliths and F-actin in gravity sensing. Plant Cell 23:1830–48 115. Nargund AM, Pellegrino MW, Fiorese CJ, Baker BM, Haynes CM. 2012. Mitochondrial import efﬁ- ciency of ATFS-1 regulates mitochondrial UPR activation. Science 337:587–90 116. Nath K, Jajoo A, Poudyal RS, Timilsina R, Park YS, et al. 2013. Towards a critical understanding of the photosystem II repair mechanism and its regulation during stress conditions. FEBS Lett. 587:3372–81 117. Nelson CJ, Alexova R, Jacoby RP, Millar AH. 2014. Proteins with high turnover rate in barley leaves estimated by proteome analysis combined with in planta isotope labelling. Plant Physiol. 166:91–108 118. Nelson CJ, Li L, Jacoby RP, Millar AH. 2013. Degradation rate of mitochondrial proteins in Arabidopsis thaliana cells. J. Proteome Res. 12:3449–59 119. Nelson CJ, Li L, Millar AH. 2013. Quantitative analysis of protein turnover in plants. Proteomics 14:579– 92 120. Ng S, De Clercq I, Van Aken O, Law SR, Ivanova A, et al. 2014. Anterograde and retrograde regulation of nuclear genes encoding mitochondrial proteins during growth, development, and stress. Mol. Plant 7:1075–93 121. Nishimura K, Asakura Y, Friso G, Kim J, Oh SH, et al. 2013. ClpS1 is a conserved substrate selector for the chloroplast Clp protease system in Arabidopsis. Plant Cell 25:2276–301 122. Nishimura K, van Wijk KJ. 2014. Organization, function and substrates of the essential Clp protease system in plastids. Biochim. Biophys. Acta. In press. doi: 10.1016/j.bbabio.2014.11.012 123. Nystrom T. 2005. Role of oxidative carbonylation in protein quality control and senescence. EMBO J. 24:1311–17 124. Okuno T, Ogura T. 2013. FtsH protease-mediated regulation of various cellular functions. Subcell. Biochem. 66:53–69 125. Olinares PD, Kim J, Davis JI, van Wijk KJ. 2011. Subunit stoichiometry, evolution, and functional implications of an asymmetric plant plastid ClpP/R protease complex in Arabidopsis. Plant Cell 23:2348– 61 126. Olinares PD, Kim J, van Wijk KJ. 2011. The Clp protease system; a central component of the chloroplast protease network. Biochim. Biophys. Acta 1807:999–1011 127. Ono Y, Wada S, Izumi M, Makino A, Ishida H. 2013. Evidence for contribution of autophagy to Rubisco degradation during leaf senescence in Arabidopsis thaliana. Plant Cell Environ. 36:1147–59 128. Ostersetzer O, Kato Y, Adam Z, Sakamoto W. 2007. Multiple intracellular locations of Lon protease in Arabidopsis: evidence for the localization of AtLon4 to chloroplasts. Plant Cell Physiol. 48:881–85 129. Otegui MS, Noh YS, Martinez DE, Vila Petroff MG, Staehelin LA, et al. 2005. Senescence-associated

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org vacuoles with intense proteolytic activity develop in leaves of Arabidopsis and soybean. Plant J. 41:831–44 130. Pan R, Kaur N, Hu J. 2014. The Arabidopsis mitochondrial membrane-bound ubiquitin protease UBP27 contributes to mitochondrial morphogenesis. Plant J. 78:1047–59 131. Paparelli E, Gonzali S, Parlanti S, Novi G, Giorgi FM, et al. 2012. Misexpression of a chloroplast aspartyl protease leads to severe growth defects and alters carbohydrate metabolism in Arabidopsis. Plant Physiol. 160:1237–50 132. Peltier JB, Ripoll DR, Friso G, Rudella A, Cai Y, et al. 2004. Clp protease complexes from photosynthetic and non-photosynthetic plastids and mitochondria of plants, their predicted three-dimensional Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. structures, and functional implications. J. Biol. Chem. 279:4768–81 133. Peters J, Stoger E. 2011. Transgenic crops for the production of recombinant vaccines and anti-microbial antibodies. Hum. Vaccines 7:367–74 134. Piechota J, Kolodziejczak M, Juszczak I, Sakamoto W, Janska H. 2010. Identiﬁcation and characterization of high molecular weight complexes formed by matrix AAA proteases and prohibitins in mitochondria of Arabidopsis thaliana. J. Biol. Chem. 285:12512–21

108 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

135. Plasman K, Van Damme P, Gevaert K. 2013. Contemporary positional proteomics strategies to study protein processing. Curr. Opin. Chem. Biol. 17:66–72 136. Ponnala L, Wang Y, Sun Q, van Wijk KJ. 2014. Correlation of mRNA and protein abundance in the developing maize leaf. Plant J. 78:424–40 137. Pratt JM, Simpson DM, Doherty MK, Rivers J, Gaskell SJ, Beynon RJ. 2006. Multiplexed absolute quantification for proteomics using concatenated signature peptides encoded by QconCAT genes. Nat. Protoc. 1:1029–43 138. Putarjunan A, Liu X, Nolan T, Yu F, Rodermel S. 2013. Understanding chloroplast biogenesis using second-site suppressors of immutans and var2. Photosynth. Res. 116:437–53 139. Quan S, Yang P, Cassin-Ross G, Kaur N, Switzenberg R, et al. 2013. Proteome analysis of peroxisomes from etiolated Arabidopsis seedlings identifies a peroxisomal protease involved in β-oxidation and development. Plant Physiol. 163:1518–38 140. Rajjou L, Lovigny Y, Groot SP, Belghazi M, Job C, Job D. 2008. Proteome-wide characterization of seed aging in Arabidopsis. A comparison between artificial and natural aging protocols. Plant Physiol. 148:620–41 141. Rawlings ND, Waller M, Barrett AJ, Bateman A. 2014. MEROPS: the database of proteolytic enzymes, their substrates and inhibitors. Nucleic Acids Res. 42:D503–9 142. Ren C, Liu J, Gong Q. 2014. Functions of autophagy in plant carbon and nitrogen metabolism. Front. Plant Sci. 5:301 143. Rigas S, Daras G, Laxa M, Marathias N, Fasseas C, et al. 2009. Role of Lon1 protease in post-germinative growth and maintenance of mitochondrial function in Arabidopsis thaliana. New Phytol. 181:588–600 144. Rigas S, Daras G, Tsitsekian D, Alatzas A, Hatzopoulos P. 2014. Evolution and significance of the Lon gene family in Arabidopsis organelle biogenesis and energy metabolism. Front. Plant Sci. 5:145 145. Rigas S, Daras G, Tsitsekian D, Hatzopoulos P. 2012. The multifaceted role of Lon proteolysis in seedling establishment and maintenance of plant organelle function: living from protein destruction. Physiol. Plant. 145:215–23 146. Roman-Hernandez G, Hou JY, Grant RA, Sauer RT, Baker TA. 2011. The ClpS adaptor mediates staged delivery of N-end rule substrates to the AAA+ ClpAP protease. Mol. Cell 43:217–28 147. Sakamoto W, Miura E, Kaji Y, Okuno T, Nishizono M, Ogura T. 2004. Allelic characterization of the leaf-variegated mutation var2 identifies the conserved amino acid residues of FtsH that are important for ATP hydrolysis and proteolysis. Plant Mol. Biol. 56:705–16 148. Sako K, Yanagawa Y, Kanai T, Sato T, Seki M, et al. 2014. Proteomic analysis of the 26S proteasome reveals its direct interaction with transit peptides of plastid protein precursors for their degradation. J. Proteome Res. 13:3223–30 149. Sauer RT, Baker TA. 2011. AAA+ proteases: ATP-fueled machines of protein destruction. Annu. Rev. Biochem. 80:587–612 150. Schuhmann H, Adamska I. 2012. Deg proteases and their role in protein quality control and processing in different subcellular compartments of the plant cell. Physiol. Plant. 145:224–34

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org 151. Schuhmann H, Huesgen PF, Adamska I. 2012. The family of Deg/HtrA proteases in plants. BMC Plant Biol. 12:52 152. Schuhmann H, Huesgen PF, Gietl C, Adamska I. 2008. The DEG15 serine protease cleaves peroxisomal targeting signal 2-containing proteins in Arabidopsis. Plant Physiol. 148:1847–56 153. Schwanhausser B, Gossen M, Dittmar G, Selbach M. 2009. Global analysis of cellular protein translation by pulsed SILAC. Proteomics 9:205–9 154. Shen G, Adam Z, Zhang H. 2007. The E3 ligase AtCHIP ubiquitylates FtsH1, a component of the chloroplast FtsH protease, and affects protein degradation in chloroplasts. Plant J. 52:309–21 Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. 155. Shen G, Yan J, Pasapula V, Luo J, He C, et al. 2007. The chloroplast protease subunit ClpP4 is a substrate of the E3 ligase AtCHIP and plays an important role in chloroplast function. Plant J. 49:228–37 156. Shibata M, Oikawa K, Yoshimoto K, Kondo M, Mano S, et al. 2013. Highly oxidized peroxisomes are selectively degraded via autophagy in Arabidopsis. Plant Cell 25:4967–83 157. Sjogren LL, Clarke AK. 2011. Assembly of the chloroplast ATP-dependent Clp protease in Arabidopsis is regulated by the ClpT accessory proteins. Plant Cell 23:322–32

www.annualreviews.org • Plant Organelle Proteolysis 109 PP66CH04-VanWijk ARI 30 March 2015 20:57

158. Smakowska E, Czarna M, Janska H. 2014. Mitochondrial ATP-dependent proteases in protection against accumulation of carbonylated proteins. Mitochondrion 19:245–51 ε 159. Smith-Hammond CL, Hoyos E, Miernyk JA. 2014. The pea seedling mitochondrial N -lysine acetylome. Mitochondrion 19:154–65 160. Solheim C, Li L, Hatzopoulos P, Millar AH. 2012. Loss of Lon1 in Arabidopsis changes the mitochondrial proteome leading to altered metabolite profiles and growth retardation without an accumulation of oxidative damage. Plant Physiol. 160:1187–203 161. Staes A, Impens F, Van Damme P, Ruttens B, Goethals M, et al. 2011. Selecting protein N-terminal peptides by combined fractional diagonal chromatography. Nat. Protoc. 6:1130–41 162. Sun R, Fan H, Gao F, Lin Y, Zhang L, et al. 2012. Crystal structure of Arabidopsis Deg2 protein reveals an internal PDZ ligand locking the hexameric resting state. J. Biol. Chem. 287:37564–69 163. Sun W, Gao F, Fan H, Shan X, Sun R, et al. 2013. The structures of Arabidopsis Deg5 and Deg8 reveal new insights into HtrA proteases. Acta Crystallogr. D 69:830–37 164. Sun X, Ouyang M, Guo J, Ma J, Lu C, et al. 2010. The thylakoid protease Deg1 is involved in photosystem-II assembly in Arabidopsis thaliana. Plant J. 62:240–49 165. Tasaki T, Sriram SM, Park KS, Kwon YT. 2012. The N-end rule pathway. Annu. Rev. Biochem. 81:261–89 166. Teixeira PF, Glaser E. 2013. Processing peptidases in mitochondria and chloroplasts. Biochim. Biophys. Acta 1833:360–70 167. Thompson EP, Smith SG, Glover BJ. 2012. An Arabidopsis rhomboid protease has roles in the chloroplast and in flower development. J. Exp. Bot. 63:3559–70 168. Tobias JW, Shrader TE, Rocap G, Varshavsky A. 1991. The N-end rule in bacteria. Science 254:1374–77 169. Urantowka A, Knorpp C, Olczak T, Kolodziejczak M, Janska H. 2005. Plant mitochondria contain at least two i-AAA-like complexes. Plant Mol. Biol. 59:239–52 170. Urban S, ed. 2013. Intramembrane proteases. Biochim Biophys. Acta 1828(12, Spec. Issue). Amsterdam: Elsevier 171. Urban S. 2013. Mechanisms and cellular functions of intramembrane proteases. Biochim. Biophys. Acta 1828:2797–800 172. Van Aken O, Whelan J, Van Breusegem F. 2010. Prohibitins: mitochondrial partners in development and stress response. Trends Plant Sci. 15:275–82 173. van der Hoorn RA. 2008. Plant proteases: from phenotypes to molecular mechanisms. Annu. Rev. Plant Biol. 59:191–223 174. van Wijk KJ, Friso G, Walther D, Schulze WX. 2014. Meta-analysis of Arabidopsis thaliana phospho- proteomics data reveals compartmentalization of phosphorylation motifs. Plant Cell 26:2367–89 175. Varshavsky A. 2011. The N-end rule pathway and regulation by proteolysis. Protein Sci. 20:1298–345 176. Venkatesh S, Lee J, Singh K, Lee I, Suzuki CK. 2012. Multitasking in the mitochondrion by the ATP- dependent Lon protease. Biochim. Biophys. Acta 1823:56–66 177. Venne AS, Vogtle FN, Meisinger C, Sickmann A, Zahedi RP. 2013. Novel highly sensitive, specific, and

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org straightforward strategy for comprehensive N-terminal proteomics reveals unknown substrates of the mitochondrial peptidase Icp55. J. Proteome Res. 12:3823–30 178. Verma D, Kanagaraj A, Jin S, Singh ND, Kolattukudy PE, Daniell H. 2010. Chloroplast-derived enzyme cocktails hydrolyse lignocellulosic biomass and release fermentable sugars. Plant Biotechnol. J. 8:332–50 179. Vogtle FN, Prinz C, Kellermann J, Lottspeich F, Pfanner N, Meisinger C. 2011. Mitochondrial protein turnover: role of the precursor intermediate peptidase Oct1 in protein stabilization. Mol. Biol. Cell 22:2135–43 180. Vogtle FN, Wortelkamp S, Zahedi RP, Becker D, Leidhold C, et al. 2009. Global analysis of the Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. mitochondrial N-proteome identiﬁes a processing peptidase critical for protein stability. Cell 139:428– 39 181. Wada S, Ishida H. 2009. Chloroplasts autophagy during senescence of individually darkened leaves. Plant Signal. Behav. 4:565–67 182. Wagner R, Aigner H, Funk C. 2012. FtsH proteases located in the plant chloroplast. Physiol. Plant. 145:203–14

110 van Wijk PP66CH04-VanWijk ARI 30 March 2015 20:57

183. Wagner R, Aigner H, Pruzinska A, Jankanpaa HJ, Jansson S, Funk C. 2011. Fitness analyses of Arabidopsis thaliana mutants depleted of FtsH metalloproteases and characterization of three FtsH6 deletion mutants exposed to high light stress, senescence and chilling. New Phytol. 191:449–58 184. Wang KH, Sauer RT, Baker TA. 2007. ClpS modulates but is not essential for bacterial N-end rule degradation. Genes Dev. 21:403–8 185. Wang Y, Yu B, Zhao J, Guo J, Li Y, et al. 2013. Autophagy contributes to leaf starch degradation. Plant Cell 25:1383–99 186. Welchen E, Garcia L, Mansilla N, Gonzalez DH. 2014. Coordination of plant mitochondrial biogenesis: keeping pace with cellular requirements. Front. Plant Sci. 4:551 187. Westphal K, Langklotz S, Thomanek N, Narberhaus F. 2012. A trapping approach reveals novel substrates and physiological functions of the essential protease FtsH in Escherichia coli. J. Biol. Chem. 287:42962–71 188. Wetzel CM, Harmacek LD, Yuan LH, Wopereis JL, Chubb R, Turini P. 2009. Loss of chloroplast protease SPPA function alters high light acclimation processes in Arabidopsis thaliana L. (Heynh.). J. Exp. Bot. 60:1715–27 189. Wiita AP, Seaman JE, Wells JA. 2014. Global analysis of cellular proteolysis by selective enzymatic labeling of protein N-termini. Methods Enzymol. 544:327–58 190. Wu W, Zhu Y, Ma Z, Sun Y, Quan Q, et al. 2013. Proteomic evidence for genetic epistasis: ClpR4 mutations switch leaf variegation to virescence in Arabidopsis. Plant J. 76:943–56 191. Xing A, Williams ME, Bourett TM, Hu W, Hou Z, et al. 2014. A pair of homoeolog ClpP5 genes underlies a virescent yellow-like mutant and its modifier in maize. Plant J. 79 192–205 192. Yoshimoto K, Shibata M, Kondo M, Oikawa K, Sato M, et al. 2014. Organ-specific quality control of plant peroxisomes is mediated by autophagy. J. Cell Sci. 127:1161–68 193. Yoshioka-Nishimura M, Nanba D, Takaki T, Ohba C, Tsumura N, et al. 2014. Quality control of photosystem II: direct imaging of the changes in the thylakoid structure and distribution of FtsH proteases in spinach chloroplasts under light stress. Plant Cell Physiol. 55:1255–65 194. Yoshioka-Nishimura M, Yamamoto Y. 2014. Quality control of photosystem II: the molecular basis for the action of FtsH protease and the dynamics of the thylakoid membranes. J. Photochem. Photobiol. B 137:100–6 195. Yu LX, Gray BN, Rutzke CJ, Walker LP, Wilson DB, Hanson MR. 2007. Expression of thermostable microbial cellulases in the chloroplasts of nicotine-free tobacco. J. Biotechnol. 131:362–69 196. Zybailov B, Friso G, Kim J, Rudella A, Rodriguez VR, et al. 2009. Large scale comparative proteomics of a chloroplast Clp protease mutant reveals folding stress, altered protein homeostasis, and feedback regulation of metabolism. Mol. Cell. Proteomics 8:1789–810 197. Zybailov B, Rutschow H, Friso G, Rudella A, Emanuelsson O, et al. 2008. Sorting signals, N-terminal modifications and abundance of the chloroplast proteome. PLOS ONE 3:e1994 Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only.

www.annualreviews.org • Plant Organelle Proteolysis 111 PP66-FrontMatter ARI 23 January 2015 17:42

Annual Review of Plant Biology Contents Volume 66, 2015

From the Concept of Totipotency to Biofortiﬁed Cereals Ingo Potrykus ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp1 The Structure of Photosystem II and the Mechanism of Water Oxidation in Photosynthesis Jian-Ren Shen ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp23 The Plastid Terminal Oxidase: Its Elusive Function Points to Multiple Contributions to Plastid Physiology Wojciech J. Nawrocki, Nicolas J. Tourasse, Antoine Taly, Fabrice Rappaport, and Francis-Andr´e Wollman pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp49 Protein Maturation and Proteolysis in Plant Plastids, Mitochondria, and Peroxisomes Klaas J. van Wijk ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp75 United in Diversity: Mechanosensitive Ion Channels in Plants Eric S. Hamilton, Angela M. Schlegel, and Elizabeth S. Haswell pppppppppppppppppppppppp113 The Evolution of Plant Secretory Structures and Emergence of Terpenoid Chemical Diversity Bernd Markus Lange pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp139 Strigolactones, a Novel Carotenoid-Derived Plant Hormone Salim Al-Babili and Harro J. Bouwmeester ppppppppppppppppppppppppppppppppppppppppppppppp161 Moving Toward a Comprehensive Map of Central Plant Metabolism ppppppppppppppppppppppppppppppppppppppppppppppppppppp

Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org Ronan Sulpice and Peter C. McKeown 187 Engineering Plastid Genomes: Methods, Tools, and Applications in Basic Research and Biotechnology Ralph Bock ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp211 RNA-Directed DNA Methylation: The Evolution of a Complex Epigenetic Pathway in Flowering Plants Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Marjori A. Matzke, Tatsuo Kanno, and Antonius J.M. Matzke ppppppppppppppppppppppppp243 The Polycomb Group Protein Regulatory Network Iva Mozgova and Lars Hennig ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp269

v PP66-FrontMatter ARI 23 January 2015 17:42

The Molecular Biology of Meiosis in Plants Raphaël Mercier, Christine Mézard, Eric Jenczewski, Nicolas Macaisne, and Mathilde Grelon ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp297 Genome Evolution in Maize: From Genomes Back to Genes James C. Schnable ppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp329 Oxygen Sensing and Signaling Joost T. van Dongen and Francesco Licausi pppppppppppppppppppppppppppppppppppppppppppppppp345 Diverse Stomatal Signaling and the Signal Integration Mechanism Yoshiyuki Murata, Izumi C. Mori, and Shintaro Munemasa pppppppppppppppppppppppppppp369 The Mechanism and Key Molecules Involved in Pollen Tube Guidance Tetsuya Higashiyama and Hidenori Takeuchi ppppppppppppppppppppppppppppppppppppppppppppp393 Signaling to Actin Stochastic Dynamics Jiejie Li, Laurent Blanchoin, and Christopher J. Staiger ppppppppppppppppppppppppppppppppp415 Photoperiodic Flowering: Time Measurement Mechanisms in Leaves Young Hun Song, Jae Sung Shim, Hannah A. Kinmonth-Schultz, and Takato Imaizumi pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp441 Brachypodium distachyon and Setaria viridis: Model Genetic Systems for the Grasses Thomas P. Brutnell, Jeffrey L. Bennetzen, and John P. Vogel ppppppppppppppppppppppppppp465 Effector-Triggered Immunity: From Pathogen Perception to Robust Defense Haitao Cui, Kenichi Tsuda, and Jane E. Parker pppppppppppppppppppppppppppppppppppppppppp487 Fungal Effectors and Plant Susceptibility Libera Lo Presti, Daniel Lanver, Gabriel Schweizer, Shigeyuki Tanaka, Liang Liang, Marie Tollot, Alga Zuccaro, Stefanie Reissmann, and Regine Kahmann pppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppppp513 Responses of Temperate Forest Productivity to Insect and Pathogen Disturbances Annu. Rev. Plant Biol. 2015.66:75-111. Downloaded from www.annualreviews.org Charles E. Flower and Miquel A. Gonzalez-Meler ppppppppppppppppppppppppppppppppppppppp547 Plant Adaptation to Acid Soils: The Molecular Basis for Crop Aluminum Resistance Leon V. Kochian, Miguel A. Piñeros, Jiping Liu, and Jurandir V. Magalhaes ppppppppp571 Terrestrial Ecosystems in a Changing Environment: A Dominant

Access provided by Chinese Academy of Agricultural Sciences (Agricultural Information Institute) on 05/10/16. For personal use only. Role for Water Carl J. Bernacchi and Andy VanLoocke pppppppppppppppppppppppppppppppppppppppppppppppppppp599

vi Contents