Journal of Plant Physiology 174 (2015) 36–40
Contents lists available at ScienceDirect
Journal of Plant Physiology journal homepage: www.elsevier.com/locate/jplph
Short communication
Crystalloids in apparent autophagic plastids: Remnants of plastids or peroxisomes? Alessio Papini a,∗ , Wouter G. van Doorn b,∗ a b
Dipartimento di Biologia Vegetale, Università di Firenze, Via La Pira 4, 50132 Florence, Italy Mann Laboratory, Department of Plant Sciences, University of California, Davis, CA 95616, USA
a r t i c l e
i n f o
Article history: Received 25 August 2014 Received in revised form 20 October 2014 Accepted 20 October 2014 Available online 29 October 2014 Keywords: Apparent autophagous plastid Crystalloids Plastids Ovary Peroxisomes
a b s t r a c t Plant macroautophagy is carried out by autophagosome-type organelles. Recent evidence suggests that plastids also can carry out macroautophagy. The double membrane at the surface of plastids apparently invaginates, forming an intraplastidial space. This space contains a portion of cytoplasm that apparently becomes degraded. Here we report, in Tillandsia sp. and Aechmaea sp., the presence of almost square or diamond-shaped crystalloids inside what seems the intraplastidial space of autophagous plastids. The same type of crystalloids were observed in chloroplasts and other plastids, but were not found in the cytoplasm or the vacuole. Peroxisomes contained smaller and more irregularly shaped crystalloids compared to the ones observed in ‘autophagous’ plastids. It is hypothesized that plastids are able to sequester chloroplasts and other plastids. © 2014 Elsevier GmbH. All rights reserved.
Introduction It has been reported that plastids seem able to sequester a portion of the cytoplasm. The outer plastid membranes apparently invaginate, forming what was called an ‘intraplastidial space’. During this process a portion of the cytoplasm is taken up into the plastid, filling the intraplastidial space. Since various stages of degradation are found inside the intraplastidial space, it is possible that plastids are able to degrade parts of the cytosol and some organelles. However, the evidence that plastids can take up parts of the cytosol and organelles for subsequent degradation is mainly based on 2D TEM images of ultrathin sections. Only one first attempt has been made to use tomography for a 3D impression. These data have not yet proven that the ‘intraplastidial’ space becomes fully closed. Although the data suggest that plastids can function as an autophagosome and autolysosome, this idea is a hypothesis with only circumstantial support (Nagl, 1977; Gärtner and Nagl, 1980; van Doorn et al., 2011; van Doorn and Papini, 2013). During leaf senescence the chloroplasts are dismantled. Some authors found what seemed to be remnants of chloroplasts in the vacuole (Wittenbach et al., 1982; Wada et al., 2009; Mulisch and Krupinska, 2013) but proof of dismantling in the vacuole seems as
∗ Corresponding authors. Tel.: +31 6 10788443; fax: +31 317 413322. E-mail addresses: alessio.papini@unifi.it (A. Papini), [email protected] (W.G. van Doorn). http://dx.doi.org/10.1016/j.jplph.2014.10.010 0176-1617/© 2014 Elsevier GmbH. All rights reserved.
yet inadequate. Hörtensteiner (2006) was unable to corroborate chloroplast dismantling in the vacuole. Chloroplasts are at least partially degraded while still in the cytoplasm (e.g. Evans et al., 2010; Yamane et al., 2012). Compared to chloroplasts, little is as yet known about the breakdown of other plastids. The data do not exclude the possibility that chloroplasts and other plastids also can become degraded in autophagous plastids. In yeasts and animal cells the degradation of peroxisomes is mainly or entirely due to both macroautophagy and microautophagy (Till et al., 2012). In Ricinus communis Vigil (1970) showed sequestering of peroxisomes (called microbodies) by plantlike autophagosomes/autolysosomes. Shibata et al. (2013) and Yoshimoto et al. (2014) corroborated autophagy of peroxisomes in plants. These data do not exclude that peroxisomes could possibly also become degraded in autophagous plastids. Here we report the presence of crystalloids in what seems the intraplastidial space of autophagous plastids. Such crystalloids were also found in peroxisomes and plastids, but the same shape was only observed in plastids. The data suggest the hypothesis that plastids can become degraded in autophagous plastids. Materials and methods Flowers were collected from Aechmaea sp. and Tillandsia sp. (both genera in the Bromeliaceae) growing in the wild, between Mexico City and Cuernavaca, Mexico. Tissues studied were the central bundle cells of anthers, the pollen at the tetrad stage of
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
37
Fig. 1. Peroxisomes in central bundle cells of an anther of Tillandsia sp. (A) A peroxisome (the lowermost organelle) close to a mitochondrion, which is located next to a chloroplast. (B) A peroxisome (uppermost organelle at the right) close to both a chloroplast and a mitochondrion. (C) A peroxisome close to a chloroplast and a mitochondrion. The shape of a crystalloid is rather irregular. (D). Irregularly shaped crystalloid. (E) White dots of the crystalloid surrounded by electron-dense parts, similar to a honey comb (arrow). C: chloroplast; M: mitochondrion; P: peroxisome; V: vacuole. Bar sizes are indicated.
development, placenta cells in ovaries, and the parenchyma underlying the nectary as well as nectar epithelium. Using a razor blade, transverse sections (0.5 mm thickness, about three per flower) were obtained. These sections were fixed overnight in 1.25% glutaraldehyde at 4 ◦ C in 0.1 M phosphate buffer (pH 6.8), then post-fixed in 1% OsO4 in the same buffer for 1 h. After dehydration in an ethanol series and a propylene
oxide step, the samples were embedded in Spurr’s epoxy resin. Cross sections approximately 80 nm thick were cut with a diamond knife and a Reichert-Jung ULTRACUT E ultramicrotome (about five to ten sections per thicker transverse section), stained with uranyl acetate (Gibbons and Grimstone, 1960), lead citrate (Reynolds, 1963), and then examined with a Philips EM300 TEM at 80 kV.
38
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
Fig. 2. Crystalloids in the stroma of plastids of Tillandsia sp. (A) Plastid in cells of an anther. (B) Chloroplasts in placenta cells. (C) Plastids in parenchyma underlying a nectary. (D) Plastids in pollen at the tetrad stage. M: mitochondrion; P: peroxisome. Bar sizes are indicated.
Results Peroxisomal crystalloids Fig. 1A and B shows examples of peroxisomes, localized close to a mitochondrion and a chloroplast. At the plane of section, these peroxisomes, like most others in the cells studied (about 80%) did not contain a crystalloid. Fig. 1C and D shows examples of peroxisomes with crystalloids. The crystallloids exhibit parallel lines (called striation) as well as a striation that is perpendicular to the first. Fig. 1E shows a higher magnification. The crystalloids were not directly surrounded by a membrane. Crystalloids in chloroplasts and other plastids Crystalloids were also observed in chloroplasts and other plastids. Fig. 2A shows one in a chloroplast. The crystalloid is almost square and shows striation. No striation perpendicular to the first was found. Fig. 2B a plastid shows a large crystalloid that forms a bulge on the organelle. Fig. 2C and D shows examples of crystalloids in what seems a non-photosynthetic plastid, as no thylakoids
are found in the plane of section. The crystalloids were not surrounded by a membrane. A double membrane related to a stack of thylakoids was found at two sides of a crystalloid (Fig. 2A, arrows) or not related to such a stack but still only at one side of the crystalloid (Fig. 2D, arrow). Crystalloids in other organelles or the cytoplasm No crystalloids were observed in vacuoles, mitochondria, nuclei, or any other parts of the cells. Crystalloids in what seems the intraplastidial space of plastids Fig. 3 shows several plastids containing a crystalloid in what is apparently the intraplastidial space. This space had low electron density. The low electron-dense area around the crystalloid varied in size. Rarely, the low electron-dense space appeared to be filled with a network of filamentous material (Fig. 3A). At some places (arrows in Fig. 3B–E) a double membrane can be discerned to delimit the low electron-dense space. The crystalloids were almost square, with rounded edges.
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
39
Fig. 3. Crystalloids in what seems to be the intraplastidial space of plastids. (A) Ovary cells of Tillandsia sp. The intraplastidial space around the crystalloid shows slight electron-density. (B) Septal nectary, secretory parenchyma of Aechmaea sp. The intraplastidial space is virtually electron-tranlucent. (C–E) Sepal nectary of Tillandsia sp. Small intraplastidial space around a crystalloid in a plastid. A double membrane around the crystalloids can be observed, which is likely the invaginated outer plastidial membrane. (C) Parenchyma underlying the secretory epithelium of the nectary of Tillandisa sp. (D) Secretory epithelium of the nectary of Tillandsia sp. (E) Secretory epithelium of the nectary of Tillandsia sp.
Discussion We observed that peroxisomes were often spatially associated both with chloroplasts and with mitochondria. Such association suggests that the peroxisomes are involved in photorespiration (Gruber et al., 1973).
Crystalloids were found in peroxisomes and plastids. The peroxisomal crystalloids varied widely in shape, and had a typical perpendicular striation pattern. The plastid crystalloids were square or diamond shape, with rounded corners. In a few of these crystalloids a single parallel striation was found. The data on peroxisome crystalloids confirm Vigil (1970), LaBorde and
40
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
Spurr (1973), Buvat (1989) and Evert (2006). Similar crystalloids as those presently observed in plastids also have been reported previously (Newcomb, 1967; Wrischer, 1973; Sprey, 1977). We previously showed evidence for autophagy carried out by plastids of Tillandsia albida, one of the species presently used (Papini et al., 2014). In the present experiments crystalloids were found in what was apparently the intraplastidial space of autophagic plastids. The evidence for their presence in the intraplastidial space was twofold: the space was delimited by a double membrane, visible at least at parts of the circumference, and by the variation from low electron-density to electron-translucent (van Doorn et al., 2011). These data may indicate the sequestering of organelles with a crystalloid, whereby the crystalloid is the last part to be degraded, or whereby it does not become degraded. The crystalloids in the intraplastidial space were square or diamond-shaped, often with rounded edges. In the present tests such crystalloids were only found in plastids. It was no possible to detect the crystal structure of the crystalloids in the intraplastidial space, hence this feature was unable to distinguish crystalloids from peroxisomes or plastids. It should be noted that the present evidence for the sequestering and degradation of plastids by plastids is only rather indirect and only preliminary. Nonetheless, the data give rise to the hypothesis that autophagic plastids can degrade plastids.
References Buvat R. Ontogeny, cell differentiation, and structure of vascular plants. Berlin: Springer; 1989. Evans IM, Rus AM, Belanger EM, Kimoto M, Brusslan JA. Plant Biol 2010;12:1–12. Evert RF. Esau’s plant anatomy. 3rd ed. Hoboken, NJ: Wiley; 2006. Gärtner PJ, Nagl W. Planta 1980;149:341–9. Gibbons IR, Grimstone AV. J Biophys Biochem 1960;7:697–716. Gruber PJ, Becker WM, Newcomb EH. J Cell Biol 1973;56:500–18. Hörtensteiner S. Annu Rev Plant Biol 2006;57:55–77. LaBorde JA, Spurr AR. Am J Bot 1973;60:736–44. Mulisch M, Krupinska K. Biswal B, Krupinska K, Biswal UC, editors. Plastid development in leaves during growth and senescence. Dordrecht: Springer; 2013. p. 307–35. Nagl W. Z Pflanzenphysiol 1977;85:45–51. Newcomb NJ. Cell Biol 1967;33:143–63. Papini A, Mosti S, van Doorn WG. Protoplasma 2014;251:719–25. Reynolds ES. J Cell Biol 1963;17:208–12. Shibata M, Oikawa K, Yoshimoto K, Kondo M, Mano S, Yamada K, et al. Plant Cell 2013;25:4967–83. Sprey B. Zeitschr Pflanzenphysiol 1977;83:159–79. Till A, Lakhani R, Burnett SF, Subramani S. Int J Cell Biol 2012:1–18, ID 512721. van Doorn WG, Kirasak K, Sonong A, Srihiran Y, van Lent J, Ketsa S. Autophagy 2011;7:584–97. van Doorn WG, Papini A. Autophagy 2013;9:1922–36. Vigil EL. J Cell Biol 1970;46:435–54. Wada S, Ishida H, Izumi M, Yoshimoto K, Ohsumi Y, Mae T, et al. Plant Physiol 2009;149:885–93. Wittenbach VA, Lin W, Hebert RR. Plant Physiol 1982;69:98–102. Wrischer M. Protoplasma 1973;77:141–5. Yamane K, Mitsuya S, Taniguchi M, Miyake H. Plant Cell Environ 2012;35:1663–71. Yoshimoto K, Shibata M, Kondo M, Oikawa K, Sato M, Toyooka K, et al. J Cell Sci 2014;127:1161–8.
Contents lists available at ScienceDirect
Journal of Plant Physiology journal homepage: www.elsevier.com/locate/jplph
Short communication
Crystalloids in apparent autophagic plastids: Remnants of plastids or peroxisomes? Alessio Papini a,∗ , Wouter G. van Doorn b,∗ a b
Dipartimento di Biologia Vegetale, Università di Firenze, Via La Pira 4, 50132 Florence, Italy Mann Laboratory, Department of Plant Sciences, University of California, Davis, CA 95616, USA
a r t i c l e
i n f o
Article history: Received 25 August 2014 Received in revised form 20 October 2014 Accepted 20 October 2014 Available online 29 October 2014 Keywords: Apparent autophagous plastid Crystalloids Plastids Ovary Peroxisomes
a b s t r a c t Plant macroautophagy is carried out by autophagosome-type organelles. Recent evidence suggests that plastids also can carry out macroautophagy. The double membrane at the surface of plastids apparently invaginates, forming an intraplastidial space. This space contains a portion of cytoplasm that apparently becomes degraded. Here we report, in Tillandsia sp. and Aechmaea sp., the presence of almost square or diamond-shaped crystalloids inside what seems the intraplastidial space of autophagous plastids. The same type of crystalloids were observed in chloroplasts and other plastids, but were not found in the cytoplasm or the vacuole. Peroxisomes contained smaller and more irregularly shaped crystalloids compared to the ones observed in ‘autophagous’ plastids. It is hypothesized that plastids are able to sequester chloroplasts and other plastids. © 2014 Elsevier GmbH. All rights reserved.
Introduction It has been reported that plastids seem able to sequester a portion of the cytoplasm. The outer plastid membranes apparently invaginate, forming what was called an ‘intraplastidial space’. During this process a portion of the cytoplasm is taken up into the plastid, filling the intraplastidial space. Since various stages of degradation are found inside the intraplastidial space, it is possible that plastids are able to degrade parts of the cytosol and some organelles. However, the evidence that plastids can take up parts of the cytosol and organelles for subsequent degradation is mainly based on 2D TEM images of ultrathin sections. Only one first attempt has been made to use tomography for a 3D impression. These data have not yet proven that the ‘intraplastidial’ space becomes fully closed. Although the data suggest that plastids can function as an autophagosome and autolysosome, this idea is a hypothesis with only circumstantial support (Nagl, 1977; Gärtner and Nagl, 1980; van Doorn et al., 2011; van Doorn and Papini, 2013). During leaf senescence the chloroplasts are dismantled. Some authors found what seemed to be remnants of chloroplasts in the vacuole (Wittenbach et al., 1982; Wada et al., 2009; Mulisch and Krupinska, 2013) but proof of dismantling in the vacuole seems as
∗ Corresponding authors. Tel.: +31 6 10788443; fax: +31 317 413322. E-mail addresses: alessio.papini@unifi.it (A. Papini), [email protected] (W.G. van Doorn). http://dx.doi.org/10.1016/j.jplph.2014.10.010 0176-1617/© 2014 Elsevier GmbH. All rights reserved.
yet inadequate. Hörtensteiner (2006) was unable to corroborate chloroplast dismantling in the vacuole. Chloroplasts are at least partially degraded while still in the cytoplasm (e.g. Evans et al., 2010; Yamane et al., 2012). Compared to chloroplasts, little is as yet known about the breakdown of other plastids. The data do not exclude the possibility that chloroplasts and other plastids also can become degraded in autophagous plastids. In yeasts and animal cells the degradation of peroxisomes is mainly or entirely due to both macroautophagy and microautophagy (Till et al., 2012). In Ricinus communis Vigil (1970) showed sequestering of peroxisomes (called microbodies) by plantlike autophagosomes/autolysosomes. Shibata et al. (2013) and Yoshimoto et al. (2014) corroborated autophagy of peroxisomes in plants. These data do not exclude that peroxisomes could possibly also become degraded in autophagous plastids. Here we report the presence of crystalloids in what seems the intraplastidial space of autophagous plastids. Such crystalloids were also found in peroxisomes and plastids, but the same shape was only observed in plastids. The data suggest the hypothesis that plastids can become degraded in autophagous plastids. Materials and methods Flowers were collected from Aechmaea sp. and Tillandsia sp. (both genera in the Bromeliaceae) growing in the wild, between Mexico City and Cuernavaca, Mexico. Tissues studied were the central bundle cells of anthers, the pollen at the tetrad stage of
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
37
Fig. 1. Peroxisomes in central bundle cells of an anther of Tillandsia sp. (A) A peroxisome (the lowermost organelle) close to a mitochondrion, which is located next to a chloroplast. (B) A peroxisome (uppermost organelle at the right) close to both a chloroplast and a mitochondrion. (C) A peroxisome close to a chloroplast and a mitochondrion. The shape of a crystalloid is rather irregular. (D). Irregularly shaped crystalloid. (E) White dots of the crystalloid surrounded by electron-dense parts, similar to a honey comb (arrow). C: chloroplast; M: mitochondrion; P: peroxisome; V: vacuole. Bar sizes are indicated.
development, placenta cells in ovaries, and the parenchyma underlying the nectary as well as nectar epithelium. Using a razor blade, transverse sections (0.5 mm thickness, about three per flower) were obtained. These sections were fixed overnight in 1.25% glutaraldehyde at 4 ◦ C in 0.1 M phosphate buffer (pH 6.8), then post-fixed in 1% OsO4 in the same buffer for 1 h. After dehydration in an ethanol series and a propylene
oxide step, the samples were embedded in Spurr’s epoxy resin. Cross sections approximately 80 nm thick were cut with a diamond knife and a Reichert-Jung ULTRACUT E ultramicrotome (about five to ten sections per thicker transverse section), stained with uranyl acetate (Gibbons and Grimstone, 1960), lead citrate (Reynolds, 1963), and then examined with a Philips EM300 TEM at 80 kV.
38
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
Fig. 2. Crystalloids in the stroma of plastids of Tillandsia sp. (A) Plastid in cells of an anther. (B) Chloroplasts in placenta cells. (C) Plastids in parenchyma underlying a nectary. (D) Plastids in pollen at the tetrad stage. M: mitochondrion; P: peroxisome. Bar sizes are indicated.
Results Peroxisomal crystalloids Fig. 1A and B shows examples of peroxisomes, localized close to a mitochondrion and a chloroplast. At the plane of section, these peroxisomes, like most others in the cells studied (about 80%) did not contain a crystalloid. Fig. 1C and D shows examples of peroxisomes with crystalloids. The crystallloids exhibit parallel lines (called striation) as well as a striation that is perpendicular to the first. Fig. 1E shows a higher magnification. The crystalloids were not directly surrounded by a membrane. Crystalloids in chloroplasts and other plastids Crystalloids were also observed in chloroplasts and other plastids. Fig. 2A shows one in a chloroplast. The crystalloid is almost square and shows striation. No striation perpendicular to the first was found. Fig. 2B a plastid shows a large crystalloid that forms a bulge on the organelle. Fig. 2C and D shows examples of crystalloids in what seems a non-photosynthetic plastid, as no thylakoids
are found in the plane of section. The crystalloids were not surrounded by a membrane. A double membrane related to a stack of thylakoids was found at two sides of a crystalloid (Fig. 2A, arrows) or not related to such a stack but still only at one side of the crystalloid (Fig. 2D, arrow). Crystalloids in other organelles or the cytoplasm No crystalloids were observed in vacuoles, mitochondria, nuclei, or any other parts of the cells. Crystalloids in what seems the intraplastidial space of plastids Fig. 3 shows several plastids containing a crystalloid in what is apparently the intraplastidial space. This space had low electron density. The low electron-dense area around the crystalloid varied in size. Rarely, the low electron-dense space appeared to be filled with a network of filamentous material (Fig. 3A). At some places (arrows in Fig. 3B–E) a double membrane can be discerned to delimit the low electron-dense space. The crystalloids were almost square, with rounded edges.
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
39
Fig. 3. Crystalloids in what seems to be the intraplastidial space of plastids. (A) Ovary cells of Tillandsia sp. The intraplastidial space around the crystalloid shows slight electron-density. (B) Septal nectary, secretory parenchyma of Aechmaea sp. The intraplastidial space is virtually electron-tranlucent. (C–E) Sepal nectary of Tillandsia sp. Small intraplastidial space around a crystalloid in a plastid. A double membrane around the crystalloids can be observed, which is likely the invaginated outer plastidial membrane. (C) Parenchyma underlying the secretory epithelium of the nectary of Tillandisa sp. (D) Secretory epithelium of the nectary of Tillandsia sp. (E) Secretory epithelium of the nectary of Tillandsia sp.
Discussion We observed that peroxisomes were often spatially associated both with chloroplasts and with mitochondria. Such association suggests that the peroxisomes are involved in photorespiration (Gruber et al., 1973).
Crystalloids were found in peroxisomes and plastids. The peroxisomal crystalloids varied widely in shape, and had a typical perpendicular striation pattern. The plastid crystalloids were square or diamond shape, with rounded corners. In a few of these crystalloids a single parallel striation was found. The data on peroxisome crystalloids confirm Vigil (1970), LaBorde and
40
A. Papini, W.G. van Doorn / Journal of Plant Physiology 174 (2015) 36–40
Spurr (1973), Buvat (1989) and Evert (2006). Similar crystalloids as those presently observed in plastids also have been reported previously (Newcomb, 1967; Wrischer, 1973; Sprey, 1977). We previously showed evidence for autophagy carried out by plastids of Tillandsia albida, one of the species presently used (Papini et al., 2014). In the present experiments crystalloids were found in what was apparently the intraplastidial space of autophagic plastids. The evidence for their presence in the intraplastidial space was twofold: the space was delimited by a double membrane, visible at least at parts of the circumference, and by the variation from low electron-density to electron-translucent (van Doorn et al., 2011). These data may indicate the sequestering of organelles with a crystalloid, whereby the crystalloid is the last part to be degraded, or whereby it does not become degraded. The crystalloids in the intraplastidial space were square or diamond-shaped, often with rounded edges. In the present tests such crystalloids were only found in plastids. It was no possible to detect the crystal structure of the crystalloids in the intraplastidial space, hence this feature was unable to distinguish crystalloids from peroxisomes or plastids. It should be noted that the present evidence for the sequestering and degradation of plastids by plastids is only rather indirect and only preliminary. Nonetheless, the data give rise to the hypothesis that autophagic plastids can degrade plastids.
References Buvat R. Ontogeny, cell differentiation, and structure of vascular plants. Berlin: Springer; 1989. Evans IM, Rus AM, Belanger EM, Kimoto M, Brusslan JA. Plant Biol 2010;12:1–12. Evert RF. Esau’s plant anatomy. 3rd ed. Hoboken, NJ: Wiley; 2006. Gärtner PJ, Nagl W. Planta 1980;149:341–9. Gibbons IR, Grimstone AV. J Biophys Biochem 1960;7:697–716. Gruber PJ, Becker WM, Newcomb EH. J Cell Biol 1973;56:500–18. Hörtensteiner S. Annu Rev Plant Biol 2006;57:55–77. LaBorde JA, Spurr AR. Am J Bot 1973;60:736–44. Mulisch M, Krupinska K. Biswal B, Krupinska K, Biswal UC, editors. Plastid development in leaves during growth and senescence. Dordrecht: Springer; 2013. p. 307–35. Nagl W. Z Pflanzenphysiol 1977;85:45–51. Newcomb NJ. Cell Biol 1967;33:143–63. Papini A, Mosti S, van Doorn WG. Protoplasma 2014;251:719–25. Reynolds ES. J Cell Biol 1963;17:208–12. Shibata M, Oikawa K, Yoshimoto K, Kondo M, Mano S, Yamada K, et al. Plant Cell 2013;25:4967–83. Sprey B. Zeitschr Pflanzenphysiol 1977;83:159–79. Till A, Lakhani R, Burnett SF, Subramani S. Int J Cell Biol 2012:1–18, ID 512721. van Doorn WG, Kirasak K, Sonong A, Srihiran Y, van Lent J, Ketsa S. Autophagy 2011;7:584–97. van Doorn WG, Papini A. Autophagy 2013;9:1922–36. Vigil EL. J Cell Biol 1970;46:435–54. Wada S, Ishida H, Izumi M, Yoshimoto K, Ohsumi Y, Mae T, et al. Plant Physiol 2009;149:885–93. Wittenbach VA, Lin W, Hebert RR. Plant Physiol 1982;69:98–102. Wrischer M. Protoplasma 1973;77:141–5. Yamane K, Mitsuya S, Taniguchi M, Miyake H. Plant Cell Environ 2012;35:1663–71. Yoshimoto K, Shibata M, Kondo M, Oikawa K, Sato M, Toyooka K, et al. J Cell Sci 2014;127:1161–8.
