TY - JOUR
AU - , Van Breusegem, Frank
AB - Abstract Compartmentation of proteins and processes is a defining feature of eukaryotic cells. The growth and development of organisms is critically dependent on the accurate sorting of proteins within cells. The mechanisms by which cytosol-synthesized proteins are delivered to the membranes and membrane compartments have been extensively characterized. However, the protein complement of any given compartment is not precisely fixed and some proteins can move between compartments in response to metabolic or environmental triggers. The mechanisms and processes that mediate such relocation events are largely uncharacterized. Many proteins can in addition perform multiple functions, catalysing alternative reactions or performing structural, non-enzymatic functions. These alternative functions can be equally important functions in each cellular compartment. Such proteins are generally not dual-targeted proteins in the classic sense of having targeting sequences that direct de novo synthesized proteins to specific cellular locations. We propose that redox post-translational modifications (PTMs) can control the compartmentation of many such proteins, including antioxidant and/or redox-associated enzymes. Catalase, moonlighting proteins, nitric oxide, reactive oxygen species, redox signaling, stromules Introduction Metabolic regulation is shaped by compartmentalization in all cells. Compartmentalization is required to achieve the stable metabolic states that underpin different cell fates (Harrington et al., 2013). Within this context, many proteins perform multiple apparently unrelated functions, often in different locations. These are often called moonlighting proteins, the classic definition of which is proteins with two or more different functions, excluding those arising from gene fusion, homologous non-identical proteins, splice variants, proteins with different post-translational modifications (PTMs), and those with a single function but active in different locations or on different substrates (Jeffery, 1999). However, the number and diversity of proteins that either can have different functions in the same intracellular compartment or that can move from one compartment to another to fulfil different functions has increased enormously in recent years, aided by development and application of bioinformatic (Chapple et al., 2015) proteomic, and cell imaging techniques (Chong et al., 2015; Thul et al., 2017). Such studies have revealed that up to 50% of cellular proteins exist in multiple subcellular localizations, which can change in response to appropriate triggers (Chong et al., 2015), such as disease states like cancers in animals (Min et al., 2016) and stress responses in plants (Sun et al., 2018). Several metabolic enzymes are known to move into the nucleus, affecting epigenetic modifications (Boukouris et al., 2016) and histone expression (He et al., 2013), providing a link between metabolism and gene expression. Plant organellar proteins such as MUTS HOMOLOG1 (MSH) 1, which is a DNA-binding nucleoid protein, function in the creation of epigenetic stress memories in plants that are associated with organellar redox changes (Xu et al., 2012; Virdi et al., 2015). Whilst many studies have been conducted on yeast and mammalian cells, there is also incontrovertible evidence for proteins with multiple, largely unrelated functions in plants (Table 1). For example, recent studies have identified a number of metabolic enzymes as members of the RNA-binding protein repertoire (Marondedze et al., 2016). Table 1. Moonlighting proteins in plants Protein . Function . Location . Moonlighting function . Reference . PUMPKIN Plastid UMP kinase Plastid RNA binding plastid transcript introns Schmid et al. (2019) WHIRLY1 Nuclear-encoded transcription factor involved in pathogen response Nucleus/plastid RNA processing in the plastid Isemer et al. (2012); Foyer et al. (2014) PEX2 Ubiquitin E3 ligase Peroxisome membrane/nucleus? ted3 gain-of-function mutant suppresses photomorphogenesis mutant det1 and evidence for interaction with Hy5 TF in nucleus, but mechanism/function unknown Hu et al. (2002); Desai et al. (2014) Catalase Antioxidant enzyme Peroxisome matrix, cytosol Hijacked to nucleus by plant pathogens to modulate cell death, but mechanism unknown Zhang et al. (2015) MSH1 Required for organelle genome stability Plastid and mitochondrial targeted Alteration in nuclear DNA methylation Virdi et al. (2015) pdNAD-MDH NAD-dependent malate dehydrogenase Plastid Activity-independent stabiliZation of FtsH12 component of inner envelope membrane protease AAA-ATPase complex. Essential for viability Schreier et al. (2018) AROGENATE DEHYDRATASE2/5 Phenylalanine Biosynthesis Stroma and stromules Interaction with chloroplast division machinery. ADT5 isoform located in nucleus Bross et al. (2017) GAPDH isoforms Glycolysis Calvin cycle Cytosol Redox-sensitive protein accumulating in the nucleus under stress conditions Zaffagnini et al. (2013); Yang and Zhai (2017) LSD1 Forms redox-dependent interaction with a suite of proteins affecting cell division vERSUS cell death Cytosol, nucleus Transcriptional activator Czarnocka et al. (2017) ACO1, aconitase Citrate metabolism, mRNA binding Cytosol TCA cycle enzyme and mRNA-binding protein to promote translation of CSD2 Moeder et al. (2007) Protein . Function . Location . Moonlighting function . Reference . PUMPKIN Plastid UMP kinase Plastid RNA binding plastid transcript introns Schmid et al. (2019) WHIRLY1 Nuclear-encoded transcription factor involved in pathogen response Nucleus/plastid RNA processing in the plastid Isemer et al. (2012); Foyer et al. (2014) PEX2 Ubiquitin E3 ligase Peroxisome membrane/nucleus? ted3 gain-of-function mutant suppresses photomorphogenesis mutant det1 and evidence for interaction with Hy5 TF in nucleus, but mechanism/function unknown Hu et al. (2002); Desai et al. (2014) Catalase Antioxidant enzyme Peroxisome matrix, cytosol Hijacked to nucleus by plant pathogens to modulate cell death, but mechanism unknown Zhang et al. (2015) MSH1 Required for organelle genome stability Plastid and mitochondrial targeted Alteration in nuclear DNA methylation Virdi et al. (2015) pdNAD-MDH NAD-dependent malate dehydrogenase Plastid Activity-independent stabiliZation of FtsH12 component of inner envelope membrane protease AAA-ATPase complex. Essential for viability Schreier et al. (2018) AROGENATE DEHYDRATASE2/5 Phenylalanine Biosynthesis Stroma and stromules Interaction with chloroplast division machinery. ADT5 isoform located in nucleus Bross et al. (2017) GAPDH isoforms Glycolysis Calvin cycle Cytosol Redox-sensitive protein accumulating in the nucleus under stress conditions Zaffagnini et al. (2013); Yang and Zhai (2017) LSD1 Forms redox-dependent interaction with a suite of proteins affecting cell division vERSUS cell death Cytosol, nucleus Transcriptional activator Czarnocka et al. (2017) ACO1, aconitase Citrate metabolism, mRNA binding Cytosol TCA cycle enzyme and mRNA-binding protein to promote translation of CSD2 Moeder et al. (2007) Open in new tab Table 1. Moonlighting proteins in plants Protein . Function . Location . Moonlighting function . Reference . PUMPKIN Plastid UMP kinase Plastid RNA binding plastid transcript introns Schmid et al. (2019) WHIRLY1 Nuclear-encoded transcription factor involved in pathogen response Nucleus/plastid RNA processing in the plastid Isemer et al. (2012); Foyer et al. (2014) PEX2 Ubiquitin E3 ligase Peroxisome membrane/nucleus? ted3 gain-of-function mutant suppresses photomorphogenesis mutant det1 and evidence for interaction with Hy5 TF in nucleus, but mechanism/function unknown Hu et al. (2002); Desai et al. (2014) Catalase Antioxidant enzyme Peroxisome matrix, cytosol Hijacked to nucleus by plant pathogens to modulate cell death, but mechanism unknown Zhang et al. (2015) MSH1 Required for organelle genome stability Plastid and mitochondrial targeted Alteration in nuclear DNA methylation Virdi et al. (2015) pdNAD-MDH NAD-dependent malate dehydrogenase Plastid Activity-independent stabiliZation of FtsH12 component of inner envelope membrane protease AAA-ATPase complex. Essential for viability Schreier et al. (2018) AROGENATE DEHYDRATASE2/5 Phenylalanine Biosynthesis Stroma and stromules Interaction with chloroplast division machinery. ADT5 isoform located in nucleus Bross et al. (2017) GAPDH isoforms Glycolysis Calvin cycle Cytosol Redox-sensitive protein accumulating in the nucleus under stress conditions Zaffagnini et al. (2013); Yang and Zhai (2017) LSD1 Forms redox-dependent interaction with a suite of proteins affecting cell division vERSUS cell death Cytosol, nucleus Transcriptional activator Czarnocka et al. (2017) ACO1, aconitase Citrate metabolism, mRNA binding Cytosol TCA cycle enzyme and mRNA-binding protein to promote translation of CSD2 Moeder et al. (2007) Protein . Function . Location . Moonlighting function . Reference . PUMPKIN Plastid UMP kinase Plastid RNA binding plastid transcript introns Schmid et al. (2019) WHIRLY1 Nuclear-encoded transcription factor involved in pathogen response Nucleus/plastid RNA processing in the plastid Isemer et al. (2012); Foyer et al. (2014) PEX2 Ubiquitin E3 ligase Peroxisome membrane/nucleus? ted3 gain-of-function mutant suppresses photomorphogenesis mutant det1 and evidence for interaction with Hy5 TF in nucleus, but mechanism/function unknown Hu et al. (2002); Desai et al. (2014) Catalase Antioxidant enzyme Peroxisome matrix, cytosol Hijacked to nucleus by plant pathogens to modulate cell death, but mechanism unknown Zhang et al. (2015) MSH1 Required for organelle genome stability Plastid and mitochondrial targeted Alteration in nuclear DNA methylation Virdi et al. (2015) pdNAD-MDH NAD-dependent malate dehydrogenase Plastid Activity-independent stabiliZation of FtsH12 component of inner envelope membrane protease AAA-ATPase complex. Essential for viability Schreier et al. (2018) AROGENATE DEHYDRATASE2/5 Phenylalanine Biosynthesis Stroma and stromules Interaction with chloroplast division machinery. ADT5 isoform located in nucleus Bross et al. (2017) GAPDH isoforms Glycolysis Calvin cycle Cytosol Redox-sensitive protein accumulating in the nucleus under stress conditions Zaffagnini et al. (2013); Yang and Zhai (2017) LSD1 Forms redox-dependent interaction with a suite of proteins affecting cell division vERSUS cell death Cytosol, nucleus Transcriptional activator Czarnocka et al. (2017) ACO1, aconitase Citrate metabolism, mRNA binding Cytosol TCA cycle enzyme and mRNA-binding protein to promote translation of CSD2 Moeder et al. (2007) Open in new tab It is important to note that not all proteins that move compartments exhibit moonlighting functions, and not all proteins with such properties move compartments. For example, l-galactono-1,4-lactone dehydrogenase has dual functions in plant mitochondria. As an enzyme it is responsible for the synthesis of ascorbic acid, and as a chaperone it is essential for the assembly of respiratory complex I (Schimmeyer et al., 2016). Similarly, plastid NAD-dependent malate dehydrogenase has a non-enzymatic function in stabilizing the FtsH12 component of the inner envelope AAA ATPase (Schreier et al., 2018). Proteins such as peroxiredoxins (PRXs) that readily undergo redox PTMs in their roles as reactive oxygen species (ROS) scavengers and oxidases have evolved to support multiple functions (acting as peroxidases, signalling proteins, and chaperones) under optimal and stress conditions (Chen et al., 2018). Like other redox proteins, whose functions are supported by thiol-based biochemistry, PRX can interact with multiple cellular partners in animals and plants, from thioredoxins (TRXs) to transcription factors (TFs) (Liebthal et al., 2018). Regulated protein relocation between the different compartments of the cell provides a robust and flexible mechanism for metabolic, genetic, and epigenetic regulation in response to metabolic stimuli and environmental cues. Such responses often entail shifts in cellular redox homeostasis that lead to both oxidative and reductive events that shift protein functions and compartmentation. One important paradigm for such redox-mediated nucleocytoplasmic shuttling in plants is Non-expresser of pathogenesis related proteins (NPR)1, which is a master regulator of salicylic acid (SA)-mediated systemic acquired resistance (SAR) leading to broad-spectrum disease resistance in plants (Mou et al., 2003). NPR1 is similar to the immune cofactor IκB and the TF NF-κB in mammals, suggesting that there is conservation of immune responses (Sun et al., 2018). In the cytosol, NPR1 exists in a large disulfide-bonded oligomeric complex. Stress-induced SA accumulation leads to reduction of the intermolecular disulfide bonds within the complex by TRXs (Tada et al., 2008) and release of the NPR1 monomers. These are then phosphorylated in the cytosol and imported into the nucleus (Mou et al., 2003). Further phosphorylation of NPR1 in the nucleus promotes interactions with TFs such as WRKY and TGA in a redox-dependent manner, leading to the expression of PR genes. Other proteins that move from the cytosol to the nucleus upon perturbation of redox homeostasis are known in yeast. The AP-1-like TF (YAP1), which is a member of the basic leucine zipper protein family (bZIP), translocates when oxidized from the cytosol to the nucleus, where it activates genes encoding oxidative stress tolerance proteins (Kuge et al., 1997), while the enzyme superoxide dismutase, SOD1, moves to the nucleus to moonlight as a TF (Tsang et al., 2014). Examples from mammalian cells include NRF2, CLK-1, and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (reviewed in Bobik and Burch-Smith, 2015; Monaghan and Whitmarsh, 2015; Min et al., 2016). A second paradigm is organelle to nucleus retrograde signalling pathways that allow cells to adapt to changes in metabolic state, often in a redox-dependent manner (Monaghan and Whitmarsh, 2015; Boukouris et al., 2016). In this review, we discuss plant proteins that are known or suspected to move location in the cell to perform alternative activities. We also present evidence in support of the hypothesis that redox cues and resulting PTMs not only alter the interactome of redox-sensitive proteins but are also central to mechanisms that facilitate the movement of proteins between compartments. Mechanisms of protein movement in plants ROS-triggered PTMs Reduction–oxidation (redox) processes not only drive cellular energy metabolism but they are crucial to cell signalling and communication (Foyer and Allen, 2003). A vast wealth of molecular genetic evidence supports the concept that ROS produced by photosynthesis, respiration, and other metabolic processes, and by specific enzymes such as NADPH oxidases, are essential signalling molecules that control plant growth and stress responses (Schmidt and Schippers, 2015; Foyer, 2018; Mhamdi and Van Breusegem, 2018). This is achieved by redox-mediated PTMs of the Cys residues on key proteins involved in multiple pathways such as primary and secondary metabolism, cell cycle, phytohormone metabolism and signalling, gene expression, translation, and protein production and transport. Chloroplasts, mitochondria, and peroxisomes are thus not only the essential sites of metabolic energy production and utilization, but also important sources of ROS and other redox regulators that influence nearly every aspect of cell biology (Noctor and Foyer, 2016). While cells regulate redox processes in a compartment-specific manner, redox PTMs may also be used to regulate the movement of proteins between compartments, as described for NPR1 above (Tada et al., 2008). Assessing the protein–protein interactions that are involved in the functions of NPR1 and other redox-regulated proteins is not only technically challenging but also entails considerations of the interdependent facets of redox state and oligomeric structure. Moreover, ROS and redox cues modify microtubule orientation and behaviour within plant cells (Dang et al., 2018), as well as the operation of protein import machineries (reviewed in Bölter et al., 2015; Ling and Jarvis, 2015). Oxidative PTMs on protein Cyst residues are formed enzymatically or non-enzymatically via promiscuous reactive species, including ROS, reactive nitrogen species (RNS), and other radicals or electrophilic lipids. There is growing appreciation that redox PTMs are site specific, governed by the microenvironment of the Cys residues, and that they are subject to temporal and spatial control. Studies using small molecule and protein-based fluorescent sensors have shown that eukaryotic cells tightly control the location of reactive species, proteins, and redox state across compartments (Kaludercic et al., 2014). However, rather than being fixed, this balance is flexible and responsive to metabolic and environmental controls. For example, it is shifted during the ageing processes in the model organism Caenorhabditis elegans (Kirstein et al., 2015). As in other organisms, redox PTMs control the activities and binding partners and probably also the compartmentation of many plant proteins, including antioxidant and/or redox-associated enzymes, and new methods, as discussed in detail below, should allow greater understanding of this level of regulation. Genetically encoded protein-based tools to trap sulfenylated proteins in situ S-Sulfenylation (protein-SOH) is a reversible oxidative PTM that acts as a regulatory switch in signal transduction pathways. However the global ‘sulfenome’ is particularly challenging to detect as this PTM is transient, unstable, and prone to overoxidation even during cell lysis. Recently a genetically encoded tool to capture S-sulfenylated proteins was developed (Waszczak et al., 2014). The Cys-rich domain of the yeast TF YAP1 forms disulfides with S-sulfenic acid modifications on its cognate signalling protein; fusion of this domain with an affinity tag creates a tool to capture and enrich S-sulfenylated proteins in vivo (Fig. 1A). YAP1 can be expressed in cells, with control cells expressing a catalytically inactive version (YAP1A), and, following cell lysis, downstream affinity purification is used to identify disulfide linked proteins. The authors detected ~100 sulfenylated proteins in Arabidopsis cell suspensions exposed to hydrogen peroxide (H2O2) oxidative stress (Waszczak et al., 2014). Fig. 1. Open in new tabDownload slide Methods to identify the sulfenome. (A) Protein-based probe YAP1C. (B) Small molecule-based probe DYn-2. Fig. 1. Open in new tabDownload slide Methods to identify the sulfenome. (A) Protein-based probe YAP1C. (B) Small molecule-based probe DYn-2. Small molecule-based probes to detect the sulfenome A complementary approach exploits the chemoselective reaction of small molecules based on dimedone with sulfenic acid. Whilst YAP1C recognition of sulfenic acids is dependent on protein–protein interactions, a small molecule is in principle more general and able to access more sulfenylation sites. The Carroll group have pioneered the use of DYn-2, a dimedone probe that is small yet readily appended to affinity tags such as biotin by click chemistry for enrichment of sulfenylated proteins (Fig. 1B) (Paulsen et al., 2011). Akter et al. (2015) applied DYn-2 is Arabidopsis cultures, identifying 226 sulfenylation events in response to oxidative stress, and, more recently, in plants (Akter et al., 2017). Protein import and export The molecular mechanisms of protein import into mitochondria, chloroplasts, and peroxisomes have now been established, and the importance of the accuracy of these processes underscored by the realization that defects result in human disease. Recent work has revealed that protein import can be regulated at several levels; from modification of individual precursor proteins to prevent or alter their targeting, to regulated interaction with binding partners, and modification of the import apparatus by phosphorylation or ubiquitination to alter its activity (Ling et al., 2012; Harbauer et al., 2014; Bölter et al., 2015) (Fig. 2). Such processes allow the location of proteins to change in response to changes in cellular state. For example, in C. elegans, the TF called ATF1 is imported into mitochondria and degraded by a Lon protease but, when import is decreased, ATF1 relocates to the nucleus and induces an unfolded protein response (Nargund et al., 2012). In mammals, import of the protein catalase (CAT) into the peroxisome is redox regulated and under stress conditions the peroxisome import receptor PEX5 retains CAT in the cytosol (Walton et al., 2017). Intriguingly, an old observation that NADPH but not NADH inhibits protein import hints at the importance of redox balance for protein import into plant peroxisomes as well (Pool et al., 1998). Retrograde signalling from organelles to the nucleus to integrate cellular activities is well established, and modulation of chloroplast import activity is important in response to biotic and abiotic stress (de Torres Zabala et al., 2015; Ling and Jarvis, 2015). Fig. 2. Open in new tabDownload slide Potential mechanisms of protein relocation. Proteins can potentially change their cellular localization by a number of mechanisms. Proteins which have been inserted into an organelle membrane can be released by regulated proteolysis as described for the chloroplast envelope-localized PTM1 and ER membrane-localized ANAC013 and ANAC017. All organelles appear to have an ER-associated degradation (ERAD)-like pathway which exports proteins in a ubiquitin-dependent manner for degradation by the proteasome. Whether proteins can be exported and escape degradation to be retargeted elsewhere in the cell is currently unknown. Ubquitination on membrane components can also lead to organelle turnover. Transport by direct organelle contacts is also a possible mechanism. Proteins normally targeted to an organelle can be prevented from import through either modification of the import machinery or modification of the cyctosolic precursor form of the protein. This can include post-translational modifications (PTMs) which can modify the targeting signal or affect interactions with other binding partners. See text for further details. Fig. 2. Open in new tabDownload slide Potential mechanisms of protein relocation. Proteins can potentially change their cellular localization by a number of mechanisms. Proteins which have been inserted into an organelle membrane can be released by regulated proteolysis as described for the chloroplast envelope-localized PTM1 and ER membrane-localized ANAC013 and ANAC017. All organelles appear to have an ER-associated degradation (ERAD)-like pathway which exports proteins in a ubiquitin-dependent manner for degradation by the proteasome. Whether proteins can be exported and escape degradation to be retargeted elsewhere in the cell is currently unknown. Ubquitination on membrane components can also lead to organelle turnover. Transport by direct organelle contacts is also a possible mechanism. Proteins normally targeted to an organelle can be prevented from import through either modification of the import machinery or modification of the cyctosolic precursor form of the protein. This can include post-translational modifications (PTMs) which can modify the targeting signal or affect interactions with other binding partners. See text for further details. Proteins are exported from mitochondria, chloroplasts, peroxisomes, and the endoplasmic reticulum (ER) (Fig. 2). The best characterized mechanisms of protein export are involved in protein degradation and in organelle quality control via the cytosolic ubiquitin–proteasome system (UPS; Ling and Jarvis, 2016; Bragoszewski et al., 2017; Kao et al., 2018). In peroxisomes, for example, an ER-associated degradation (ERAD)-like system exports the import receptor PEX5 from the peroxisome membrane to perform further rounds of import. PEX5 cycling between the peroxisome and cytosol is regulated by ubiquitination of a conserved Cys, and in mammalian cells reduced glutathione can de-ubiquitinate the receptor (Grou et al., 2009). Although not shown experimentally, this Cys is conserved in plant PEX5 proteins, suggestive of a similar mechanism operating. Mutants in this PEX5 re-export system in Arabidopsis also fail to degrade some peroxisome matrix proteins, suggesting that they are exported for degradation (Burkhart et al., 2013). The release of TFs from cellular membranes by regulated proteolysis is also a well-known response to stress in both animals and plants (Seo et al., 2008; Sun et al., 2011). Potentially, protein export from organelles and retargeting (rather than degradation) could provide a means of signalling and genetic regulation. To date this has only been proposed/described for a handful of proteins, and the mechanism(s) by which this occurs and is regulated are still obscure (Foyer et al., 2014; Bobik and Burch-Smith, 2015). The next section presents proteins which are candidates for regulated relocation. Candidates as a paradigm for redox-regulated movement in plants Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) GAPDH is a quintessential example of a moonlighting protein (Sirover, 2012, 2014). Several GAPDH isoforms exist in different subcellular localizations in plants (Holtgrefe et al., 2008). In animals, it has multiple functions in addition to its classic role in glycolysis, such as DNA stability and control of gene expression, autophagy, and apoptosis. Both the activity and localization of the plant cytosolic GAPDH isoform (GapC) are controlled by the cellular redox state (Bedhomme et al., 2012). Redox PTMs on the cytosolic GAPDH protein in animals block enzyme activity and promote novel cell signalling and transcription functions in the nucleus (Zaffagnini et al., 2013; Yang and Zhai, 2017). The functions of GAPDH in the nuclei of plant cells are not clear, but nuclear GAPDH may have a role as a coactivator for gene expression (Hildebrandt et al., 2015). Since GapC is also localized in the nucleus, it is suggested that redox modification facilitates transfer to the nucleus in plants as it does in animals (Ortiz-Ortiz et al., 2010). However, the mechanism of nuclear translocation of GapC is unknown although it is thought to involve S-sulfhydration, a process that reversibly regulates the function of this protein, in a manner similar to that described in mammalian systems (Aroca et al., 2015). However, GapC undergoes S-nitrosylation, S-glutathionylation, S-sulfhydration, S-sulfenylation, as well as other modifications that all occur on the same Cys residue (Lindermayr et al., 2005; Bedhomme et al., 2012; Waszczak et al., 2014; Aroca et al., 2017). Thus, how each type of PTM modifies GapC to shift location and/or alternatively instigate non-metabolic functions remains to be determined. Catalase (CAT) CAT is a peroxisomal enzyme whose import in mammals is redox regulated (Walton et al., 2017) and in yeast is dependent on carbon source (Horiguchi et al., 2001). In plants, it is classically known as a peroxisomal enzyme, but recent evidence suggests that the compartmentation of this central antioxidant enzyme may be more dynamic than the literature acknowledges. The role of CAT as a central ‘redox guardian’ is well established (Mhamdi et al., 2012). Plant CATs have been shown to interact with a variety of cytosolic proteins including calmodulin (Yang and Poovaiah, 2002), calcium-dependent protein kinase 8 (CDPK8) (Zou et al., 2015), salt overly sensitive 2 (SOS2) (Verslues et al., 2007), lesion simulating disease1 (LSD1) (Li et al., 2013), receptor-like cytoplasmic kinase STRK1 (Zhou et al., 2018), and no catalase activity 1 (NCA1) (Hackenberg et al., 2013; Li et al., 2015) (Fig. 3). All are integral stress signalling proteins. The nca1 mutants, which lack a functional CAT, are hypersensitive to abiotic stresses. Similarly, the cat2 mutant of Arabidopsis, which lacks the predominant leaf isoform that is essential for the metabolism of H2O2 produced by photorespiration, activates a wide range of SA- and jasmonic acid (JA)-dependent responses and displays day-length-dependent localized programmed cell death (PCD) and resistance to pathogens (Queval et al., 2010). CAT can also be a target for pathogen-encoded effector proteins (Mathioudakis et al., 2013; Murota et al., 2017). The fungal effectors PsCRN115 and PsCRN63 both traffic CAT to the nucleus but have opposite biochemical and physiological effects. PsCRN115 stabilizes CAT, decreases H2O2, and reduces PCD, whereas PsCRN63 destabilizes CAT, increases H2O2, and increases PCD (Zhang et al., 2015). How can the interaction of peroxisomal CAT with such a wide variety of cytosolic proteins be explained? Some interactions may be occurring during biosynthesis in the cyotsol before import into peroxisomes. However, the evidence that CAT interacts with different stress signalling and PCD proteins potentially provides a mechanism for protein retention and/or relocation. We speculate that the location of cytosolically synthesized CAT is determined by competition among different potential binding partners as a consequence of reduced import into peroxisomes and/or increased retention of CAT in the cytosol. While sensitivity of peroxisomal protein import to redox status is likely to impact import of all peroxisome proteins, CAT, which has a non-canonical targeting signal (Mhamdi et al., 2012; Rymer et al., 2018), may be more sensitive, and indeed PEX5, the major peroxisome import receptor, has been proposed specifically to retain mammalian CAT in the cytosol under conditions of oxidative stress (Walton et al., 2017). This property, combined with the potential to interact with an array of cytosolic proteins as shown in Fig. 3, could allow swift control of CAT localization between compartments in such a way as to influence various redox signalling pathways. Fig. 3. Open in new tabDownload slide Switching partners: model for regulation of catalase localization through interaction with different binding proteins. Several cytosolic proteins have been reported to interact with plant catalases. Redox-mediated PTMs could alter the affinity of catalase for different binding partners, leading to a change in distribution between peroxisomes, cytosol, and nucleus. See text for further details. Fig. 3. Open in new tabDownload slide Switching partners: model for regulation of catalase localization through interaction with different binding proteins. Several cytosolic proteins have been reported to interact with plant catalases. Redox-mediated PTMs could alter the affinity of catalase for different binding partners, leading to a change in distribution between peroxisomes, cytosol, and nucleus. See text for further details. WHIRLY1 (WHY1) WHY1 is a member of a small family of ssDNA-binding proteins that are specific to the plant kingdom (Desveaux et al., 2004, 2005). WHY1 protein is encoded in the nuclei and targeted to chloroplasts and the nuclei, with the nuclear and processed chloroplast forms having the same molecular mass (Grabowski et al., 2008). Studies using epitope-tagged, transplastomically expressed WHY1 provided evidence that WHY1 can move directly from the chloroplasts to the nuclei (Isemer et al., 2012; Foyer et al., 2014). However, the factors that trigger and regulate this apparently direct movement of this protein from the chloroplasts to the nuclei are unknown. PTM of WHY1 in the cytosol can also regulate the partitioning between the chloroplasts and nuclei. This change in partitioning is regulated by phosphorylation of WHY1 in the cytosol by a serine/threonine SNF1-related protein kinase called calcineurin B-Like-Interacting Protein Kinase14 (CIPK14). Phosphorylation of WHY1 results in transport to the nucleus (Ren et al., 2017). The phosphorylation of WHY1 in the cytosol regulates the intracellular localization with respect to leaf development. WHY1 is predominantly in the chloroplasts of young leaves, while in senescing leaves the protein is localized mainly in the nucleus (Ren et al., 2017). It is perhaps not surprising therefore that WHY1 is a multifunctional protein, with DNA binding properties that are relevant in both cellular compartments. WHY1 is important in the regulation of chloroplast development, plastome copy number and plastome gene expression, chloroplast ribosome formation, and chloroplast to nucleus signalling (Prikryl et al., 2008; Comadira et al., 2015). WHY1 promotes rRNA splicing that is catalysed by other factors within plastids (Prikryl et al., 2008). Nuclear WHY1 is involved in the expression of senescence and defence genes as well as in the maintenance of telomeres (Yoo et al., 2007). ROXY proteins The glutaredoxins ROXY1 and ROXY2 are found in both the nuclei and cytosol (Delorme-Hinoux et al., 2016). ROXY1 interacts with the TGA TFs called TOPLESS in the nuclei, in a redox-dependent manner, and with four other TGA TFs. While there is as yet no direct evidence of the redox-regulated movement of ROXY1 between the nucleus and cytosol, there is no other explanation for the dual compartmentation of this protein except that ROXY proteins are small and may pass through the nuclear pores. Heat shock factor (HSF) A8 Like other HSFs, HSFA8 is retained in the cytosol in an inactive form by interaction with heat shock proteins (HSPs), which mask the nuclear location signal and the oligomerization domain. In response to oxidative and other stresses, HSFs oligomerize and are translocated into the nucleus, where they modulate the expression of target genes (Scharf et al., 2012). Redox-mediated nucleocytoplasmic shuttling has been characterized for HSFA8 in Arabidopsis thaliana (Giesguth et al., 2015). Cys24, which is located in the DNA-binding domain of AtHSFA8, and Cys269, which is located in the C-terminal part of the protein, act as redox sensors. Disulfide bond formation between Cys24 and Cys269 is thought to induce release from multiheteromeric complexes and translocation into the nucleus (Giesguth et al., 2015) Membrane-bound transcription factors Membrane-located proteins can be cleaved from their membrane anchor in response to an appropriate signal to release a soluble domain that can be relocated (Fig. 2). Often these proteins function as TFs once liberated from the membrane. ANAC013 and ANAC017 encode Arabidopsis TFs belonging to the NON APICAL MERISTEM/ARABIDOPSIS TRANSCRIPTION ACTIVATION FACTOR/CUP SHAPED COTYLEDON (NAC) family. These TFs mediate ROS-related retrograde signalling originating from mitochondrial complex III. Both proteins are anchored to the ER membrane. They were identified via one-hybrid assays as binding to a conserved cis-acting regulatory sequence termed the mitochondrial dysfunction motif (MDM) which mediates mitochondrial retrograde regulation (MRR) during oxidative stress (De Clercq et al., 2013). Green fluorescent protein (GFP)–ANAC013 was partially processed and nuclear localized, but, while difficult to detect, there was a suggestion that the full-length protein is ER targeted (De Clercq et al., 2013). ANAC017 was also identified in a screen for loss of response to mitochondrial dysfunction (Ng et al., 2013). It is targeted to the ER, and dual tagging experiments showed that it is cleaved upon antimycin A treatment, which inhibits the mitochondrial electron transport chain at complex III. The N-terminal part of ANAC017 locates to the nucleus whilst the C-terminal part remains ER associated. ANAC017 function was essential for H2O2-mediated stress signalling (Ng et al., 2013). Upon perception of redox signals, ANAC013 and ANAC017 are released from the ER and translocated to the nucleus, where they activate MDS genes such as those encoding alternative oxidase (AOX), SOT12, and ANAC013. The latter provides positive feedback regulation of the signalling pathway with enhancement of the signal. The ROS-dependent signalling pathways from chloroplasts and mitochondria merge at RADICAL-INDUCED CELL DEATH1 (RCD1), a nuclear protein that is suggested to suppress the activities of the ANAC013 and ANAC017 TFs (Shapiguzov et al., 2019). Another member of the family, ANAC089, is an ER- and trans-Golgi network-localized membrane protein. Upon treatment with reducing agents, the N-terminal domain of ANAC089 localizes to nuclei where it partially supresses chloroplast stromal ascorbate peroxidase gene expression (Klein et al., 2012). The chloroplast-bound plant homeodomain TF called PTM1 (TF PTM1) was proposed to play a crucial role in chloroplast signalling to the nucleus. The full-length TF PTM1 is located to the chloroplast outer envelope, whereas a truncated form lacking the transmembrane domain was found in the nucleus. Treatments such as high light and Norfluazon were reported to result in cleavage of TF PTM1 and localization of the N-terminal domain to the nucleus. Processed TF PTM1was shown to activate ABI4 transcription (Sun et al., 2011). Mutants defective in this gene show aberrant responses to treatments such as Norfluazon, high light, dibromothymoquonine, and Rose Bengal that affect different ROS and the level of reduction of the plastoquinione pool (Sun et al., 2011), although these results were not observed in a subsequent study (Page et al., 2017), leaving the role of TF PTM1 in chloroplast signalling questionable. Like TF PTM1, ABI4 is no longer considered to be involved in chloroplast to nucleus signalling (Kacprzak et al., 2019). PEX2 is a peroxisome membrane protein with a cytosolically exposed RING domain E3 ligase that regulates the recycling and turnover of the PEX5 import receptor through ubiquitination (Burkhart et al., 2014). Interestingly a mutant of Arabidopsis PEX2 (ted3) was recovered as a suppressor of the photomorphogenesis mutant det1 (Hu et al., 2002). The mechanism of this remains unknown, but an artificially expressed RING domain was found in the nucleus where it interacted with the TF called HY5 (Desai et al., 2014). Possible mechanisms could be cleavage of the RING domain and relocation to the nucleus, alternative transcription/translation sites, or direct movement between the peroxisome and nuclear membrane. Since peroxisomes are important nodes in the cell’s antioxidant network and import is under redox control, we speculate that PEX2 relocation could represent a potential mechanism for sensing the redox state of peroxisomes and relaying this information to the nucleus. The above list is not exhaustive and it may in fact be the tip of the iceberg because there are many proteins in the literature that are suggested to undergo intercompartmental switching in response to appropriate triggers. Arabidopsis hexokinase 1, for example, which is located at the outer mitochondrial membrane, has been suggested to translocate between the mitochondrion and nucleus, upon perception of sugar signals or methyljasmonate, in a manner that is linked to mitochondrial ROS production (Claeyssen and Rivoal, 2007; Xiang et al., 2011). Organelle movement and contact as a mechanism of protein movement Apart from release of proteins from membranes, prevention of import into or promotion of export from organelles, direct transfer of proteins between membrane-bound compartments via membrane extensions and contact sites can occur (Pérez-Sancho et al., 2016) (Fig. 4). The cytoplasm in plant cells is densely packed and mainly constrained by the vacuole and ER to a narrow cortical zone. Protein transfer between organelles requires regulated release and redirection. Redirection through the cytosol may be slow and prevent bulk delivery. Emerging evidence suggests that the physical interaction between organelles is a requirement for the exchange of small molecules, lipids, and proteins in plants, as well as in mammals and yeast (Cohen et al., 2018; Liu and Li, 2019). Coordinated re-arrangement of organelle positioning within the cell could provide a mechanism for shuttling moonlighting proteins between compartments. The ‘protected’ delivery of protein targets from degradation, or potential reversal of the PTM, could be provided through the formation of a microenvironment between organelles that allows for exchanging proteins through a narrow 10–40 nm cytoplasmic zone at the membrane contact site interface. Repositioning of organelles could also allow neighbouring organelles to signal to one another to regulate protein exchange. Fig. 4. Open in new tabDownload slide Organelle interactions through protrusions and membrane contact sites. Organelle–organelle interactions in cells occurs either by the formation of membrane contact sites (MCSs) between organelles or by the formation of tubular structures by one organelle. MCSs are known to occur between mitochondria and plastids, mitochondria and the ER, and plastids and the ER. In addition, mitochondria, plastids, and peroxisomes form tubular structures. In the case of plastids, stromules are formed especially in the direction of the nucleus. Mitochondria form matrixules within ER structures and peroxisomes form peroxules in the vicinity of plastids, mitochondria, and the nucleus. Both MCSs and the tubular structures will mediate communications between organelles by exchanging signalling molecules, metabolites, or potentially even proteins. Fig. 4. Open in new tabDownload slide Organelle interactions through protrusions and membrane contact sites. Organelle–organelle interactions in cells occurs either by the formation of membrane contact sites (MCSs) between organelles or by the formation of tubular structures by one organelle. MCSs are known to occur between mitochondria and plastids, mitochondria and the ER, and plastids and the ER. In addition, mitochondria, plastids, and peroxisomes form tubular structures. In the case of plastids, stromules are formed especially in the direction of the nucleus. Mitochondria form matrixules within ER structures and peroxisomes form peroxules in the vicinity of plastids, mitochondria, and the nucleus. Both MCSs and the tubular structures will mediate communications between organelles by exchanging signalling molecules, metabolites, or potentially even proteins. Redox-dependent formation of stromules, matrixules, and peroxules Chloroplasts, mitochondria, and peroxisomes are pleomorphic, dynamic organelles that produce tubules upon stress. Like membrane contact sites (MCSs, these tubules allow positioning of the organelles in relation to each other within the cell and might be involved in the exchange of metabolites or macromolecules. For example, stroma-filled tubules called stromules (Fig. 5) can extend from the envelope of all plastid types in response to appropriate stimuli. ROS increase the speed of movement and dynamics of peroxisomes, resulting in membrane extensions (peroxules), which could facilitate contact with other organelles including chloroplasts (Rodríguez-Serrano et al., 2009, 2016; Gao et al., 2016). However, the cargo of the tubular structures and the nature of the potential signals (metabolic or proteinaceous) that are released are largely unknown (Hanson and Hines, 2018). Dynamin-type proteins are thought to be involved in stromule formation, as well as in the formation of vesicles that are shed from the stromule tips (Hanson and Hines, 2018). At least some of the plastid-derived vesicles are found in the vacuole, where they fulfil a role in chloroplast degradation. Fig. 5. Open in new tabDownload slide Stromule formation in N. benthamiana leaves upon transient overexpression of GFP-tagged plastid outer membrane protein AtLACS9 (At1g77560). Agrobacterium tumefaciens carrying AtLACS9–GFP (Breuers et al., 2012) and a second strain carrying the P19 silencing suppressor construct (Takeda et al., 2002) were co-infiltrated at 0.4 OD each into 7-week-old N. benthamiana leaves. Fluorescence imaging was done at 72 h post-infiltration with a Zeiss LSM710 confocal microscope. The image is a maximum projection of 10 optical sections. GFP (green); chlorophyll autofluorescence (red). Fig. 5. Open in new tabDownload slide Stromule formation in N. benthamiana leaves upon transient overexpression of GFP-tagged plastid outer membrane protein AtLACS9 (At1g77560). Agrobacterium tumefaciens carrying AtLACS9–GFP (Breuers et al., 2012) and a second strain carrying the P19 silencing suppressor construct (Takeda et al., 2002) were co-infiltrated at 0.4 OD each into 7-week-old N. benthamiana leaves. Fluorescence imaging was done at 72 h post-infiltration with a Zeiss LSM710 confocal microscope. The image is a maximum projection of 10 optical sections. GFP (green); chlorophyll autofluorescence (red). Stromules allow actin-mediated anchoring of chloroplasts at different locations within the cell to facilitate specific functions. For example, they can extend along microtubules to guide chloroplast movement to the nucleus during innate immunity responses. The application of H2O2 resulted in rapid stromule formation in Arabidopsis leaves (Caplan et al., 2015). The accumulation of ROS, like other pro-defence molecules, is sufficient to induce stromule formation, leading to the development of direct contact points between the chloroplasts and nuclei (Caplan et al., 2015). In addition, other direct contact sites between chloroplasts and nuclei that are induced by high light have been suggested to allow movement of H2O2 to the nucleus from attached chloroplasts (Exposito-Rodriguez et al., 2017). Arogenate dehydratase (ADT) 2 which catalyses the final step in phenylalanine biosynthesis localizes to stromules and also helps in dividing chloroplasts, whilst ADT5 is proposed to traffic to nuclei via stromules, (Bross et al., 2017). Another interesting example of possible organelle to organelle transport of proteins via membrane extensions is the triacyl glycerol lipase SDP1 which is proposed to move from peroxisomes to oil bodies in a tubule- and retromer-dependent process (Thazar-Poulot et al., 2015). Mitochondria produce structures that are partly homologous to the chloroplast stromules, in response to light and other stimuli in an ER-mediated manner (Schmidt et al., 2016). The protrusion-driven movement and positioning is considered to promote the intercompartmental trafficking of metabolites and proteins, but there remains a paucity of data on which proteins are trafficked and the mechanisms involved. ROS and redox cues modify microtubule orientation and behaviour within cells, as well as the operation of protein import and export machineries (Schmidt et al., 2016). So far, it remains to be determined if these organelle-derived tubular structures are involved in direct exchange of metabolites or macromolecules between compartments, or might rather have a supportive function in the communication between organelles by acting as a cellular anchor to temporarily fix their position relative to each other. Conclusions and perspectives Our understanding of ROS functions has been entirely revised in recent decades. Initially confined to oxidative stress and associated cellular damage, ROS are now recognized as signals released from the plasma membrane and organelles to orchestrate plant growth and stress tolerance. Moreover, the same oxidative changes to proteins such as irreversible oxidation or nitrosylation of glutathionylation of Cys residues that were once regarded as damage are now recognized as being instrumental in regulating protein–protein interactions and signalling. However, gaps remain; for example, little is known about how protein carbonylation functions as a PTM in response to cellular redox changes. Literature evidence supports the concept that changes in ROS production alter the redox status of plant cells, exerting a strong influence on metabolism and gene expression. Redox-related PTMs may have important effects on chromatin structure and function, opening up a new area of redox epigenetics (García-Giménez et al., 2012). Histone PTMs have a direct impact on chromatin conformation, controlling important cellular events such as cell proliferation and differentiation. The carbonylation of specific histones (H1, H10, and H3.1 dimers) has been described during DNA synthesis in proliferating NIH3T3 fibroblasts, where it was found to decrease when nuclear proteasome activity was activated, suggesting that this PTM prevents excessive histone accumulation during DNA synthesis (García-Giménez et al., 2012). Until recently the paucity of experimental data on subcellular protein distribution has limited our understanding of the capacity and ability of proteins to move between different intracellular compartments. There has been a step change in our knowledge of proteins that perform more than one cellular function. The term given to such proteins is ‘moonlighting’, but this description is limited as we have discussed above because it does not apply to all proteins that move between different cellular compartments. Moreover, it has become increasingly apparent that protein localization is not fixed, and a high proportion of cellular proteins have the potential to move between compartments in response to specific triggers. In some cases this movement is the basis for an alternative cellular function. At present, however, we have only a fragmented picture with relatively few well-characterized examples of proteins in plants that change compartment in order to moonlight, and the mechanisms by which they do so are largely unexplored. Here we have presented evidence in support of an extension to existing concepts suggesting that redox PTMs are likely to be a key driver for intercompartmental shifts of antioxidant and redox-regulated proteins. Redox cues and associated PTMs are fundamental regulators of alternative protein functions and localization. However, the extent of this phenomenon, what makes proteins move, and the mechanisms by which they do so remain largely obscure. Redox-regulated PTMs that drive intercompartmental protein relocation have the potential to integrate metabolic processes and influence genetic and epigenetic controls of plant growth and stress tolerance. This prospect is already opening up a new intriguing and technically challenging area of research. Although it is widely recognized that ROS act as signals through the redox processing of other molecules, particularly proteins, relatively little is known about the network of proteins that undergo redox-mediated PTMs, highlighting the need for improved redox proteomics approaches. Moreover, a larger tool box of molecular and cell biology techniques is required to fully understand the redox-mediated movement of organelles and the associations/dissociations between different cellular compartments, as well as if and how redox-mediated structural changes facilitate direct movement of proteins from one compartment to another, particularly between chloroplasts, mitochondria, peroxisomes, and nuclei, without the need to transverse the cytosol in between. Author contributions CHF and AB proposed the original idea and wrote the first draft of the manuscript text. All authors polished and contributed to the final manuscript text. AB produced Figs 1 and 2, and Table 1. JS produced Fig. 3 and harmonized the style of all the figures. MW produced Fig. 1 and wrote the sections relating to this figure. Abbreviations: Abbreviations: ADT2 arogenate dehydratase 2 CAT catalase ER endoplasmic reticulum GAPDH glyceraldehyde 3-phosphate dehydrogenase HSF heat shock factor HSP heat shock proteins MDM mitochondrial dysfunction motif MRR mitochondrial retrograde regulation NPR1 NON-EXPRESSOR OF PATHOGENESIS-RELATED GENES 1 PR pathogenesis-related PRX peroxiredoxin PTM post-translational modification ROS reactive oxygen species RNS reactive nitrogen species SA salicylic acid SAR systemic acquired resistance SOD superoxide sismutase TF transcription factor TRX thioredoxin UPS ubiquitin–proteasome system WHY1 WHIRLY1 Acknowledgements The authors gratefully acknowledge Professor Patricia Conklin State University of New York College at Cortland for the image in Fig. 5. CHF thanks BBSRC (UK) for financial support (BB/N004914/1). FVB and AM are supported by the Fonds Wetenschappelijk Onderzoek-Vlaanderen and Fonds De La Recherche Scientifique–FNRS under the EOS project (O018218F). 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TI - On the move: redox-dependent protein relocation in plants
JO - Journal of Experimental Botany
DO - 10.1093/jxb/erz330
DA - 2020-01-07
UR - https://www.deepdyve.com/lp/oxford-university-press/on-the-move-redox-dependent-protein-relocation-in-plants-9H2oM2gTGy
SP - 620
VL - 71
IS - 2
DP - DeepDyve
ER -