Abstract Light energy is essential for photosynthetic energy production and plant growth. Chloroplasts in green tissues convert energy from sunlight into chemical energy via the electron transport chain. When the level of light energy exceeds the capacity of the photosynthetic apparatus, chloroplasts undergo a process known as photoinhibition. Since photoinhibition leads to the overaccumulation of reactive oxygen species (ROS) and the spreading of cell death, plants have developed multiple systems to protect chloroplasts from strong light. Recent studies have shown that autophagy, a system that functions in eukaryotes for the intracellular degradation of cytoplasmic components, participates in the removal of damaged chloroplasts. Previous findings also demonstrated an important role for autophagy in chloroplast turnover during leaf senescence. In this review, we describe the turnover of whole chloroplasts, which occurs via a type of autophagy termed chlorophagy. We discuss a possible regulatory mechanism for the induction of chlorophagy based on current knowledge of photoinhibition, leaf senescence and mitophagy—the autophagic turnover of mitochondria in yeast and mammals. Introduction Plants absorb light energy from the sun using Chl pigments and convert the energy from visible light (wavelengths of 400–700 nm) into chemical energy via the photosynthetic electron transport chain, comprising PSII, the Cyt b6f complex, PSI and the ATP synthase complex. These photosynthetic reactions occur in the chloroplast. The conversion of light energy can potentially damage the photosynthetic machinery via a process known as photoinhibition (Aro et al. 1993, Li et al. 2009). Plants concomitantly absorb UV-A (wavelengths of 315–400 nm) and UV-B (280–315 nm) radiation, which can directly damage macromolecules in the cell, such as proteins, DNA and lipids (Takahashi and Badger 2011, Kataria et al. 2014). UV-related damage may enhance photoinhibition (Takahashi and Badger 2011). Reactive oxygen species (ROS) are actively produced during photoinhibition and directly cause further oxidative damage to chloroplasts (Asada 2006). Consequently, plants have developed diverse chloroplast protection systems to quench excess light energy, repair photodamaged proteins and scavenge ROS (Takahashi and Badger 2011); however, the fate of photodamaged, collapsed chloroplasts is not clearly understood. Autophagy: A Major Intracellular Degradation System for Cytoplasmic Components in Eukaryotes Organelle turnover in eukaryotic cells is widely achieved via autophagy-related transport into lytic organelles, including lysosomes in animal cells and the vacuole in yeast and plant cells (Ohsumi 2001). Macroautophagy is a well-characterized autophagic process by which cytoplasmic components are engulfed by double-membrane-bound vesicles known as autophagosomes. The outer membrane of the autophagosome then fuses with the lysosomal or vacuolar membrane and releases the inner membrane-bound autophagic body into the lysosomal or vacuolar lumen (Nakatogawa et al. 2009, Mizushima and Komatsu 2011). During another type of autophagy termed microautophagy, cytoplasmic components are directly engulfed by the invaginated membranes of the lysosome or vacuole, and the sequestered material is subsequently degraded (Li et al. 2012). This process is well characterized in the methylotrophic yeast Pichia pastoris (Oku and Sakai 2016), in which the switch from the use of methanol to glucose as the cell’s energy source activates the microautophagic digestion of peroxisomes. AUTOPHAGY (ATG) genes were originally identified in the budding yeast Saccharomyces cerevisiae (Tsukada and Ohsumi 1993). To date, 41 ATG genes have been identified in yeast, including 15 (ATG1–10, 12–14, 16 and 18) ‘core’ ATG genes that are required for all types of autophagy (Nakatogawa et al. 2009). Core ATGs are classified into four subgroups: (i) ATG1 and ATG13 are components of the ATG1 kinase complex; (ii) ATG6 and ATG14 are components of the autophagy-specific phosphatidylinositol 3-kinase (PI3K) complex; (iii) ATG2 and ATG18 form a complex with membrane-anchored ATG9; and (iv) the remaining core ATGs participate in the two ubiquitin-like conjugation systems that facilitate ATG8 lipidation and autophagosomal membrane elongation (Nakatogawa et al. 2009). Through the two ubiquitin-like cascades, ATG8 is conjugated with a lipid, phosphatidylethanolamine, subsequently forming the autophagosomal membrane (Ichimura et al. 2000). These core autophagy components are mainly involved in autophagosome formation, and their orthologs have been identified in various plant species (Meijer et al. 2007, Chung et al. 2009, Zhou et al. 2015). Autophagy mediates the bulk digestion of cytoplasmic components and facilitates the recycling of released molecules, such as amino acids, especially under starvation conditions. In addition, specific organelles or proteins are selectively transported into lytic organelles as selective cargoes of autophagosomes under various conditions (Anding and Baehrecke 2017). This selective autophagy process leads to the removal of dysfunctional organelles; for example, dysfunctional mitochondria are removed through a selective autophagy process termed mitophagy in yeast and mammals (Youle and Narendra 2011, Kanki et al. 2015). Chlorophagy Removes Whole Photodamaged Chloroplasts Studies on Arabidopsis thaliana mutants of core ATG genes indicate that the core autophagy machinery for the initiation and elongation of the autophagosomal membrane has been conserved in plants (Li and Vierstra 2012, Y. Liu et al. 2012, Yoshimoto 2012). The establishment of in vivo monitoring methods for plant autophagy based on fluorescent marker proteins of the autophagosomal membrane or organelles has further facilitated studies of the involvement of autophagy in the intracellular turnover of plant organelles (Yoshimoto et al. 2004, Thompson et al. 2005). A recent study investigated the possibility that autophagy participates in the turnover of photodamaged chloroplasts under stress conditions (Izumi et al. 2017). This study revealed that whole chloroplasts are transported into the vacuole following photodamage caused by exposure to strong visible light or UV-B through an autophagic process termed chlorophagy. This phenomenon was observed in true rosette leaves of Arabidopsis plants grown in soil under a 12 h light/12 h dark photoperiod using fluorescent lamps (140 μmol m–2 s–1) at 23°C. When plants grown under these conditions were exposed to strong visible light of various photosynthetic photon flux densities (PPFDs; 800, 1,200, 1,600 and 2,000 µmol m–2 s–1) for 3 h, chlorophagy was only observed after exposure to >1,200 µmol m–2 s–1 PPFD (Izumi et al. 2017). Natural sunlight includes visible light, UV-A and UV-B. Exposure of chamber-grown Arabidopsis plants to natural sunlight also induces chlorophagy (Izumi et al. 2017), through sunlight damage. Methods for Assessing Chlorophagic Activity Fig. 1 shows the current methods used to detect and assess chlorophagic activity in Arabidopsis. When transgenic plants expressing stroma-targeted green or red fluorescent protein (GFP or RFP) are grown under normal conditions without photodamage treatment, all chloroplasts exhibiting Chl autofluorescence produce signals from stroma-targeted fluorescent protein when observed under a confocal microscope (Izumi et al. 2017; Fig. 1A). At 2 d after a 2 h exposure to high levels of visible light (HL; 2,000 µmol m–2 s–1), chloroplasts lacking stroma-targeted fluorescent protein signals that appear to move randomly are observed in the central regions of mesophyll cells (Fig. 1A, arrowheads), specifically in the central vacuole, as chloroplasts lacking stroma-targeted RFP were observed inside the tonoplast (labeled by GFP; Fig. 1B, arrowheads). Transmission electron microscopy (TEM) also revealed that chloroplasts accumulate in the vacuole after HL exposure (Fig. 1C, arrowheads). These chloroplasts have retained their thylakoid membranes but have lost their stromal components, which is consistent with confocal microcopy observations of vacuolar chloroplasts labeled with fluorescent protein markers. It is thought that when chloroplasts are incorporated into the vacuole via chlorophagy, envelope and stromal components are degraded and diffuse before the thylakoid structures, including Chl, are digested; such chloroplasts appear as stromal marker-deficient chloroplasts under confocal microscopy (Fig. 1, arrowheads). TEM images show that vacuolar chloroplasts are partially fragmented, supporting the notion that vacuolar chloroplasts are in the process of being digested (Fig. 1C). Such observations led to the discovery of chlorophagy, a process by which whole photodamaged chloroplasts are transported into the central vacuole (Fig. 1D;Izumi et al. 2017). Fig. 1 View largeDownload slide Images and schematic representation of chlorophagy induced by strong visible light in Arabidopsis. (A) Confocal images of leaf mesophyll cells expressing stroma-targeted GFP under the control of the 35S promoter. The second rosette leaves of non-treated control plants or plants at 2 d after exposure to 2 h of high visible light (HL; 2,000 μmol m–2 s–1) at 10°C were observed. Arrowheads indicate chloroplasts lacking stroma-localized GFP. Chl autofluorescence appears magenta, and GFP signals appear green. In the merged images, overlapping regions of Chl and GFP appear white. Differential interference contrast (DIC) images are also shown. Scale bars = 10 µm. (B) Confocal images of leaf mesophyll cells expressing tonoplast-targeted GFP–δ-tonoplast intrinsic protein (δTIP) under the control of the 35S promoter and stroma-targeted Rubisco small subunit (RBCS)–RFP under the control of the RBCS promoter. The second rosette leaves of non-treated control plants or plants 1 d after exposure to 2 h HL at 10°C were observed. Arrowheads indicate chloroplasts in the vacuolar lumen. Chl autofluorescence appears magenta. In the merged images, GFP and RFP signals appear green. DIC images are also shown. Scale bars = 10 µm. (C) TEM images of leaf mesophyll cells from wild-type plants. The second rosette leaves of non-treated control plants or plants 1 d after exposure to 2 h of HL at 10°C were fixed and observed. Images in the right-hand panels are enlargements of the boxed regions in the left-hand panels. Scale bars = 5 µm. Arrowheads indicate vacuolar chloroplasts resulting from chlorophagy. (D) Schematic model of photodamage-induced chlorophagy. In this model, photodamaged chloroplasts are transported into the vacuolar lumen for degradation via autophagic membrane-associated sequestration. Fig. 1 View largeDownload slide Images and schematic representation of chlorophagy induced by strong visible light in Arabidopsis. (A) Confocal images of leaf mesophyll cells expressing stroma-targeted GFP under the control of the 35S promoter. The second rosette leaves of non-treated control plants or plants at 2 d after exposure to 2 h of high visible light (HL; 2,000 μmol m–2 s–1) at 10°C were observed. Arrowheads indicate chloroplasts lacking stroma-localized GFP. Chl autofluorescence appears magenta, and GFP signals appear green. In the merged images, overlapping regions of Chl and GFP appear white. Differential interference contrast (DIC) images are also shown. Scale bars = 10 µm. (B) Confocal images of leaf mesophyll cells expressing tonoplast-targeted GFP–δ-tonoplast intrinsic protein (δTIP) under the control of the 35S promoter and stroma-targeted Rubisco small subunit (RBCS)–RFP under the control of the RBCS promoter. The second rosette leaves of non-treated control plants or plants 1 d after exposure to 2 h HL at 10°C were observed. Arrowheads indicate chloroplasts in the vacuolar lumen. Chl autofluorescence appears magenta. In the merged images, GFP and RFP signals appear green. DIC images are also shown. Scale bars = 10 µm. (C) TEM images of leaf mesophyll cells from wild-type plants. The second rosette leaves of non-treated control plants or plants 1 d after exposure to 2 h of HL at 10°C were fixed and observed. Images in the right-hand panels are enlargements of the boxed regions in the left-hand panels. Scale bars = 5 µm. Arrowheads indicate vacuolar chloroplasts resulting from chlorophagy. (D) Schematic model of photodamage-induced chlorophagy. In this model, photodamaged chloroplasts are transported into the vacuolar lumen for degradation via autophagic membrane-associated sequestration. Fluorescently labeled stroma-targeted proteins can be used to distinguish easily vacuolar chloroplasts (resulting from chlorophagy) from cytoplasmic chloroplasts (Fig. 1). The direct observation and counting of vacuole-incorporated chloroplasts using plants expressing stroma-targeted fluorescent proteins is a simple, reliable method for assessing chlorophagic activity. In fact, the number of stroma-deficient vacuolar chloroplasts increases in response to greater chloroplast damage, as represented by the larger decline in the maximum quantum yield of PSII (Fv/Fm; Izumi et al. 2017). Studies examining organelle-targeted autophagy frequently involve biochemical assays using organelle marker proteins fused with fluorescent proteins, in which free fluorescent proteins derived from vacuolar degradation of the fusion proteins are detected by immunoblot analysis of protein extracts (Mizushima et al. 2010). For instance, mitophagic activity in yeast has been assessed by detecting free GFP released via the vacuolar degradation of the mitochondria-targeted fusion protein Om45–GFP (consisting of the C-terminus of the mitochondrial outer membrane protein Om45 and GFP; Kanki et al. 2009). The establishment of similar biochemical methods specifically to monitor the occurrence of chlorophagy in combination with other techniques might allow for the future quantitative evaluation of chlorophagy induction under various conditions. The Relationship Between Photoinhibition and Chlorophagy During PTEN-induced putative kinase 1 (PINK1) and Parkin (PINK1/Parkin)-mediated mitophagy in mammals (Fig. 2B), depolarized mitochondria that lose transmembrane potential (ΔΨ) across the inner envelope for ATP synthesis become the targets of selective removal (Youle and Narendra 2011). Similarly, damaged chloroplasts suffering from a specific damage might be selectively removed in individual mesophyll cells during chlorophagy. The decline in Fv/Fm represents the extent of photoinhibition, and chlorophagy is activated in response to larger declines in Fv/Fm (Izumi et al. 2017); therefore, we postulate that photoinhibition-associated chloroplast damage is closely related to the selective recognition of the cargo of chlorophagy. Multiple systems prevent the occurrence of photoinhibition in chloroplasts. Excessive light energy absorbed by the PSII light-harvesting complex is quenched as heat energy through a mechanism known as thermal energy dissipation (Havaux and Niyogi 1999). The efficiency of this energy dissipation corresponds to the extent of ΔpH across the thylakoid membrane (Jahns and Holzwarth 2012). Cyclic electron flow around PSI can produce high ΔpH levels during photosynthesis (Shikanai and Yamamoto 2017). Metabolic processes across chloroplasts, mitochondria and peroxisomes, such as the malate–oxaloacetate shuttle and photorespiration, probably help dissipate excessive reducing power (Yamori 2016). When the reducing power produced by excess light energy is not sufficiently dissipated, the photosystems produce ROS, including singlet oxygen (1O2) from PSII or hydrogen peroxide (H2O2) and superoxide (O2–) from PSI (Asada 2006). Chloroplasts have scavenging systems for ROS: 1O2 is detoxified by carotenoids that closely localize around the PSII reaction centers (Ramel et al. 2012), O2– is quickly dismutated to H2O2 by superoxide dismutase (SOD), and H2O2 is detoxified by ascorbate peroxidase (APX; Asada 2006). Accumulated ROS and increasing reducing power primarily damage the D1 reaction center within PSII (Aro et al. 1993). Damaged D1 turns over very rapidly via the co-operative activity of two types of intrachloroplastic proteases, FtsH and Deg, and is replaced by newly synthesized D1 (Kato et al. 2012). Photoinhibition appears when light energy exceeds the capacity of these protection and repair mechanisms. Such conditions are sometimes caused by the interference of additional abiotic stresses, such as drought and low temperatures, with photosynthetic reactions (Yamori 2016). Even under normal light conditions that do not induce strong photoinhibition (100 µmol m–2 s–1), mutants of a major subunit of FtsH (FtsH2) showed compromised D1 degradation and accumulated more ROS in their leaf chloroplasts than the wild type (Kato et al. 2009). Therefore, PSII damage constantly occurs under normal (non-stressed) growth conditions, but photoinhibition of PSII does not occur when the PSII repair system sufficiently restores such damage. It is thought that if a chloroplast sustains local damage that can be sufficiently repaired by intrachloroplastic systems, and chloroplast functions can be maintained, the chloroplast would be subjected to local repair systems instead of total degradation via chlorophagy. Therefore, given that PSII damage occurs constantly and is rapidly repaired by proteases, PSII photoinhibition is unlikely to be the direct trigger of chlorophagy. In contrast to PSII, PSI does not have a quick repair system; PSI repair is a relatively slow process compared with that of PSII, requiring several days for completion (Scheller and Haldrup 2005). PSI damage mainly involves the O2–-induced damage of iron–sulfur (FeS) clusters within the PSI reaction centers. PSI damage was originally considered to occur only in response to specific treatments under experimental conditions, such as exposure to moderate light with chilling treatment (Sonoike 1998); conversely, recent studies have indicated that PSI damage may constantly occur under fluctuating light conditions, such as in natural sunlight (Yamori 2016). PROTON GRADIENT REGULATION5 (PGR5) is a PSI-associated protein that is required for the generation of the ΔpH across the thylakoid membrane through the activation of cyclic electron flow (DalCorso et al. 2008, Shikanai and Yamamoto 2017). The Arabidopsis pgr5 mutant accumulates more severe damage to PSI during HL illumination compared with wild-type plants, and the growth of this mutant is strongly suppressed under experimentally fluctuating light conditions, i.e. exposure to repeated cycles of 5 min of moderate light and 1 min of strong light throughout the day (Suorsa et al. 2012). Thus, the accumulation of PSI damage upon sudden irradiation under fluctuating light conditions probably leads to fatal damage. In the Arabidopsis chloroplast, stromal APX (sAPX) and thylakoid APX (tAPX) help scavenge O2– and H2O2 (Maruta et al. 2012). The possible involvement of O2– and H2O2 accumulation in the induction of chlorophagy was suggested by the observation that UV-B damage-induced chlorophagy is activated in tAPX mutant plants compared with the wild type (Izumi et al. 2017). Therefore, O2–-related damage, including PSI photoinhibition, might be linked to the induction of chlorophagy. Photoinhibition May Damage the Envelope The core autophagy machinery is limited to the cytoplasm, and the envelope acts as a border between the chloroplast and cytoplasm. During PINK1/Parkin-mediated selective mitophagy in mammalian cells, the modification of the outer envelope is a key induction signal for this process, which follows the loss of ΔΨ across the inner envelope (Fig. 2B). Therefore, it is possible that altered envelope integrity may act as a trigger for the induction of chlorophagy. In support of this theory, recent studies have established that the chloroplast envelope can accumulate damage and that VESICLE-INDUCING PROTEIN IN PLASTIDS1 (VIPP1) plays an important role in maintaining envelope integrity (Zhang et al. 2012). The VIPP1 homolog in Escherichia coli, Phage Shock Protein A, helps maintain plasma membrane integrity. In plants, VIPP1 binds to the membrane and functions in membrane remodeling (Heidrich et al. 2017). VIPP1–GFP fusion protein localizes around the chloroplast envelope in the form of large particles approximately 1 µm in diameter that appear to move quickly around chloroplasts during osmotic stress (Zhang et al. 2012). VIPP1 has an intrinsically disordered region in its C-terminus; deletion of the C-terminal region of VIPP1–GFP fusion protein led to increased aggregation of these particles, thereby inhibiting their active movement and preventing them from protecting the chloroplast membrane (Zhang et al. 2016b). VIPP1–GFP-overexpressing Arabidopsis plants showed enhanced tolerance to heat shock, but the expression of VIPP1 with a truncated C-terminus increased sensitivity to this stress (Zhang et al. 2016b). These reports highlight the importance of protecting the chloroplast membrane during plant stress responses. VIPP1-knockdown Arabidopsis plants have abnormal, swollen chloroplasts, indicating that the integrity of the chloroplast envelopes in these plants is impaired. Swollen chloroplasts are also observed in seedlings of an Arabidopsis mutant of NON-YELLOW COLORING1 (NYC1), encoding an enzyme that degrades Chl (Nakajima et al. 2012); nyc1 seedlings contain chlorotic cotyledons with swollen chloroplasts (Zhang et al. 2016a). This phenomenon is likely to be caused by Chl-related photooxidative damage, since the number of seedlings with chlorotic cotyledons increases with increasing PPFD during growth. Overexpressing VIPP–GFP in nyc1 plants restored their abnormal chloroplast shape and defective cotyledon phenotypes (Zhang et al. 2016a). These results indicate that the envelope is a target of photooxidative damage within chloroplasts and that VIPP1 can alleviate such envelope damage. In UV-B-damaged Arabidopsis leaves, few chloroplasts exhibit ruptured envelopes, similar to those found in the cytoplasm of UV-B-damaged atg plants (Izumi et al. 2017). TEM observations of mesophyll cells in UV-B-damaged atg leaves revealed normal as well as abnormal chloroplasts with altered shapes and disorganized thylakoid membranes. Treatment of tobacco leaf cells with methyl viologen, which enhances the production of O2– within PSI, can lead to the rupture of the envelope (Kwon et al. 2013), indicating that the envelope can suffer ROS-mediated damage. As shown in Fig. 1C, some chloroplasts in HL-damaged mesophyll cells have abnormal shapes. In sum, the extent of envelope damage and the related morphological changes to chloroplasts as a result of ROS producution around PSII and PSI during the induction of chlorophagy should be a major focus of further study. Regulatory Mechanisms of Mitophagy to Remove Damaged Mitochondria in Yeast and Mammals The mitophagy regulatory mechanism for mitochondrial quality control has been extensively studied in yeast and mammals. During PINK1/Parkin-mediated mitophagy in mammals, depolarized mitochondria are eliminated, as mentioned previously. In healthy mitochondria, PINK1 is imported into mitochondria and subsequently degraded by the inner membrane-localized serine protease PARL (Jin et al. 2010). The ΔΨ across the inner membrane is also required for mitochondrial protein import; thus, its loss allows PINK1 to accumulate on the TOM (translocase of the outer membrane) complex (Matsuda et al. 2010, Narendra et al. 2010, Vives-Bauza et al. 2010, Lazarou et al. 2012). The accumulated PINK1 phosphorylates ubiquitin and the ubiquitin E3 ligase, Parkin, to activate Parkin-mediated ubiquitination of mitochondria, thereby leading to the build up of ubiquitin chains on mitochondrial outer membrane proteins (Koyano et al. 2014). PINK1 and Parkin-mediated ubiquitination recruit various autophagic receptors that bind to autophagosome-anchored LC3 (a mammalian homolog of ATG8; Lazarou et al. 2015). These molecular events allow for the transport of depolarized mitochondria as a specific cargo of autophagosomes. Therefore, PINK1- and Parkin-mediated ubiquitination act as inducers, allowing dysfunctional mitochondria to be selectively eliminated (Fig. 2B). During mitophagy in yeast, ATG32 acts as an autophagic receptor that is directly anchored to the outer membranes of oxidized mitochondria and interacts with ATG8 (Kanki et al. 2009, Okamoto et al. 2009). ATG proteins with ATG8-interacting motifs also participate in the selective turnover of other organelles in yeast. For example, ATG39 and ATG40 were identified (in a co-immunoprecipitation assay of yeast ATG8) as the autophagic receptors of nucleus- or endoplasmic reticulum (ER)-targeted autophagy (nucleophagy or ER-phagy; Mochida et al. 2015). The Roles of Plant ATG8-Interacting Proteins and Chloroplast-Associated Ubiquitination in Organelle Turnover To remove collapsed chloroplasts selectively via chlorophagy in plant cells, these chloroplasts must be recognized by a specific protein that functions in a manner similar to PINK1 and ATG32 during mitophagy in mammalian cells and yeast, respectively. Three AUTOPHAGY8-INTERACTING PROTEINS (ATIs) have been identified in plants. ATI1 and 2 interact with the ER or plastids, forming small vesicles during sugar starvation (Honig et al. 2012, Michaeli et al. 2014), and ATI3 may be involved in ER turnover during ER stress (Zhou et al. 2018). Thus, ATI1–ATI3 are unlikely to be the autophagic receptors that trigger photodamage-induced chlorophagy. A recent genetic screen indicated that the selective removal of chloroplasts involves ubiquitination (Woodson et al. 2015). When etiolated seedlings of the plastid-localized FERROCHELATASE2 Arabidopsis mutant, fc2, are transferred from darkness to light, 1O2 accumulates in their chloroplasts. This ROS accumulation causes the death of photosynthetic cells and impairs plant greening. A suppressor mutant of this inhibited greening phenomenon has an additional single amino acid substitution in PLANT U-BOX4 (PUB4), a cytosol-localized ubiquitin E3 ligase. In double mutants of FC2 and PUB4, the digestion of whole chloroplasts in the cytoplasm is suppressed compared with fc2 single mutants, even though 1O2 accumulation is not affected in these mutants. Therefore, PUB4-related ubiquitination triggers the degradation of 1O2-accumulating chloroplasts. TEM images of greening fc2 plants suggest that entire chloroplasts are digested in the cytoplasm and that these digested chloroplasts interact with the central vacuole via globule-like structures (Woodson et al. 2015). In contrast, during chlorophagy, whole chloroplasts that have retained thylakoid membranes and exhibit Chl autofluorescence accumulate in the vacuolar lumen (Fig. 1). These distinct observations suggest that PUB4-related ubiquitination is not a simple trigger of chlorophagy and that it controls another pathway that specifically degrades 1O2-accumulating chloroplasts. In the cytoplasm, ubiquitinated proteins are generally degraded by the 26S proteasome complex (Vierstra 2012). SUPPRESSOR OF PLASTID PROTEIN IMPORT1 LOCUS1 (SP1) is a ubiquitin E3 ligase that is anchored to the chloroplast outer envelope and induces proteasome-dependent degradation of some proteins of the TOC (translocon on the outer chloroplast membrane) complex (Ling et al. 2012, Ling and Jarvis 2015). To date, only two ubiquitin E3 ligases, PUB4 and SP1, were found to be associated with the ubiquitination of chloroplasts. Eukaryotic genomes generally encode large families of ubiquitin E3 ligases, and Arabidopsis can express >1,500 of these proteins based on genome-wide analysis (Vierstra 2012). Therefore, another as yet unidentified ubiquitin E3 ligase might be involved in the induction of chlorophagy. The Regulation of Chlorophagy During Leaf Senescence Leaf senescence is a developmental process during which cytoplasmic components including chloroplasts undergo massive degradation and the released molecules are remobilized to newly developing organs. Photoinhibition may be enhanced during senescence, since photosynthetic activity decreases due to the degradation of photosynthetic proteins, and ROS accumulation is generally enhanced in senescing leaves (Juvany et al. 2013). Such enhanced ROS accumulation might activate chlorophagy during senescence. However, entire chloroplasts were transported to the vacuole via chlorophagy at later stages of accelerated senescence in individual Arabidopsis leaves when covered with aluminum foil (Wada et al. 2008), which is an experimental condition widely used to analyze phenomena during leaf senescence. Under this condition, another type of chloroplast-targeted autophagy is preferentially activated, in which a portion of the chloroplast stroma is transported to the vacuole as a specific autophagic vesicle termed the Rubisco-containing body (RCB; Ishida et al. 2008, Izumi et al. 2015). Chloroplasts in covered senescing leaves are much smaller than those in young leaves; therefore, the active separation of stroma via RCBs is thought to result in chloroplast shrinkage, and these small chloroplasts are believed to become whole targets of autophagy (Izumi and Nakamura 2018). In covered leaves that do not acquire light, photodamage does not occur; thus, the idea that senescence-induced chlorophagy and photodamage-induced chlorophagy are differentially regulated appears to be reasonable. In mammals, other forms of mitophagy distinct from the PINK1/Parkin-mediated type have been observed (Fig. 2B). In most mammals, red blood cells lack mitochondria due to the autophagic removal of mitochondria that accumulate the LC3-interacting protein, NIX (Nip3-like protein; also known as BNIP3L), on the outer envelope (Schweers et al. 2007, Sandoval et al. 2008). This form of mitophagy is triggered by the up-regulation of NIX expression during red blood cell differentiation. When ΔΨ in mitochondria declines due to cell hypoxia, another LC3-interacting protein, FUN14 domain-containing 1 (FUNDC1), accumulates on the outer envelope, thereby inducing mitophagy (L. Liu et al. 2012). Hypoxia-induced dephosphorylation of FUNDC1 triggers this mitophagic process. Together, these findings suggest that in plants, chlorophagy might also be regulated by distinct mechanisms in different organ types, conditions or developmental stages. Diverse pathways contribute to the degradation of intrachloroplastic components during leaf senescence without causing the digestion of entire chloroplasts via chlorophagy (Izumi and Nakamura 2018). In addition to the separation of stroma via the RCB pathway, Chls are actively degraded through the autophagy-independent cascade via multistep enzymatic reactions (Hortensteiner and Krautler 2011). Autophagy-independent routes that degrade stroma, thylakoid and envelope components during senescence include the formation of senescence-associated vacuoles, i.e. small vacuoles generated in the cytoplasm in senescing leaves (Martinez et al. 2008) and CHLOROPLAST VESICULATION-containing vesicles, a type of vesicle that mobilizes a portion of the chloroplast into the vacuole (Wang and Blumwald 2014). These active degradation processes of intrachloroplastic components might produce almost empty chloroplasts that have lost photosynthetic activity. Therefore, it is also conceivable that the same proteins that function during photodamage-induced chlorophagy also recognize senescence-induced dysfunctional chloroplasts; however, the initial event that occurs during the induction of chlorophagy in both cases is distinct. Conclusions and Future Perspectives The discovery of photodamage-induced chlorophagy has prompted new questions, including what types of chloroplast damage induce chlorophagy, how the damaged chloroplasts are recognized and recruited to the core autophagy machinery, and whether photodamage-induced chlorophagy and senescence-induced chlorophagy share a common regulatory mechanism (Fig. 2A). Our summary of the process of photoinhibition indicates that damage accumulates in PSII and PSI, which is manifested as ROS accumulation and chloroplast envelope damage. Thus, investigating chlorophagic activity in mutants of the respective systems that alleviate each type of damage may help clarify the direct triggers of chlorophagy within photodamaged chloroplasts. Based on studies of mitophagy in yeast and mammals, we postulate that unknown inducers and autophagic receptors selectively recognize chloroplasts that exhibit specific types of damage and recruit them as cargoes for chlorophagy (Fig. 2). Chloroplasts are approximately 5–7 µm in diameter, which is much larger than mitochondria and typical autophagosomes, which are only approximately 1 µm in diameter (Yoshimoto et al. 2004, Thompson et al. 2005). How chloroplasts are incorporated into the vacuole, i.e. via macroautophagy, microautophagy or other pathways, is another fascinating issue to uncover (Fig. 2). Fig. 2 View largeDownload slide Possible mechanism for the regulation of chlorophagy: lessons from mitophagy regulatory mechanisms in mammals. (A) Possible events leading to photodamage-induced chlorophagy. Plant chloroplasts can accumulate several types of damage during photoinhibition, including PSII and PSI damage, ROS accumulation and envelope damage. Specific types of damage within the chloroplast might act as a direct trigger of chlorophagy. Based on our understanding of mitophagy in mammals (shown in B), unknown proteins that interact with targeted chloroplasts might act as inducers or autophagic receptors for chlorophagy. Outer envelope-associated proteins or ubiquitins might be involved in this induction process. How chloroplasts are incorporated into the vacuole remains unknown. (B) Schematic models of the events leading to three types of selective mitophagy mechanisms in mammalian cells. (a) PINK1/Parkin-mediated mitophagy is initiated upon the accumulation of PINK1 on the outer membranes of depolarized mitochondria. PINK1 then phosphorylates ubiquitin to activate the E3 ligase, Parkin, thereby leading to the accumulation of ubiquitin chains on the outer envelope. Several types of autophagic receptors that bind to LC3 (a mammalian homolog of ATG8), including NDP52, optineurin and p62, interact with ubiquitinated mitochondrial proteins and autophagosome-anchored LC3, which induces the sequestering of depolarized mitochondria by the autophagosome. (b) NIX acts as a mitophagy receptor that directly binds to LC3 on the outer envelope to induce mitophagy during red blood cell differentiation. This phenomenon is triggered by the up-regulation of NIX expression. (c) Dephosphorylation of FUNDC1 on the mitochondrial outer envelope in response to hypoxia allows the protein to interact directly with LC3, thereby inducing mitophagy. Fig. 2 View largeDownload slide Possible mechanism for the regulation of chlorophagy: lessons from mitophagy regulatory mechanisms in mammals. (A) Possible events leading to photodamage-induced chlorophagy. Plant chloroplasts can accumulate several types of damage during photoinhibition, including PSII and PSI damage, ROS accumulation and envelope damage. Specific types of damage within the chloroplast might act as a direct trigger of chlorophagy. Based on our understanding of mitophagy in mammals (shown in B), unknown proteins that interact with targeted chloroplasts might act as inducers or autophagic receptors for chlorophagy. Outer envelope-associated proteins or ubiquitins might be involved in this induction process. How chloroplasts are incorporated into the vacuole remains unknown. (B) Schematic models of the events leading to three types of selective mitophagy mechanisms in mammalian cells. (a) PINK1/Parkin-mediated mitophagy is initiated upon the accumulation of PINK1 on the outer membranes of depolarized mitochondria. PINK1 then phosphorylates ubiquitin to activate the E3 ligase, Parkin, thereby leading to the accumulation of ubiquitin chains on the outer envelope. Several types of autophagic receptors that bind to LC3 (a mammalian homolog of ATG8), including NDP52, optineurin and p62, interact with ubiquitinated mitochondrial proteins and autophagosome-anchored LC3, which induces the sequestering of depolarized mitochondria by the autophagosome. (b) NIX acts as a mitophagy receptor that directly binds to LC3 on the outer envelope to induce mitophagy during red blood cell differentiation. This phenomenon is triggered by the up-regulation of NIX expression. (c) Dephosphorylation of FUNDC1 on the mitochondrial outer envelope in response to hypoxia allows the protein to interact directly with LC3, thereby inducing mitophagy. Elucidation of the chlorophagy induction mechanism is still in the initial stages. To improve our understanding of this mechanism, additional studies should investigate chloroplast function and compare organelle-selective autophagy among different eukaryotes. Funding This work was supported by Japan Society for the Promotion of Science (JSPS) [KAKENHI grant Nos. 17H05050 and 18H04852 to M.I., and 16J03408 to S.N., and Research Fellowship for Young Scientists to S.N.]; Japan Science and Technology Agency (JST) [PRESTO grant No JPMJPR16Q1 to M.I.]; and the Program for Creation of Interdisciplinary Research at Frontier Research Institute for Interdisciplinary Sciences, Tohoku University, Japan [to M.I.]. Acknowledgments We thank Maureen R. Hanson for stroma-targeted GFP-expressing plants, Hiroyuki Ishida for RBCS–RFP-expressing plants, and Youshi Tazoe for critical reading of the manuscript. Disclosures The authors have no conflicts of interest to declare. References Anding A.L. , Baehrecke E.H. ( 2017 ) Cleaning house: selective autophagy of organelles . Dev. 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Google Scholar CrossRef Search ADS PubMed Abbreviations Abbreviations APX ascorbate peroxidase ATG AUTOPHAGY ATI AUTOPHAGY 8-INTERACTING PROTEIN ER endoplasmic reticulum FC2 FERROCHELATASE 2 FUNDC1 FUN14 domain containing 1 GFP green fluorescent protein HL high visible light NIX Nip3-like protein X NYC1 NON-YELLOW COLORING 1 PINK1 PTEN-induced putative kinase 1 PPFD photosynthetic photon flux density PUB4 Plant U-Box 4 RCB Rubisco-containing body RFP red fluorescent protein ROS reactive oxygen species SP1 SUPPRESSOR OF PLASTID PROTEIN IMPORT1 LOCUS 1 TEM transmission electron microscopy VIPP1 VESICLE-INDUCING PROTEIN IN PLASTIDS 1 © The Author(s) 2018. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. 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Plant and Cell Physiology – Oxford University Press
Published: May 14, 2018
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